Arthropoda

Chapter 3

Arthropoda Chapter 3a

Arthropoda, Diptera, Nematocera Larry R. Cruthers, MS, PhD1 and Marco Pombi, PhD2 1

LCruthers Consulting, Chesapeake, VA, United States, 2Dipartimento di Sanita` Pubblica e Malattie Infettive, Sapienza University of Rome, Rome, Italy

Arthropoda Diptera Nematocera The order Diptera is composed of three main groups: Nematocera (Culicidae—mosquitoes, Simulidae—black flies, Ceratopogonidae—biting midges, and Psychodidae—sand flies), Brachycera (horse and deer flies), and Cyclorrhapha (Muscidae—houseflies, Sarcophagidae—flesh flies, Calliphoridae—blow flies, and Oestridae—bot flies). All three contain bloodsucking species. The name of the order Diptera, that is, insects with two wings, is derived from the Greek di meaning two, and ptera meaning wings. Adult flies of this order are typically equipped with a single pair of membranous wings attached to the dorsolateral angle of the second thoracic segment. Only the forewings are used to fly, while the second pair of hindwings is reduced into a pair of club-like balancing organs known as halters. All flies have a complete metamorphosis (i.e., egg, larval stage with several instars, pupa, and adult). Adult flies escape through a circular opening in the anterior of the puparium or pupal case using a ptilinum, a bladder-like structure inflated with hemolymph (inflatable facial sac), that projects from the frontal suture and is then withdrawn into the head. The antennae are three-segmented with an anteriorly directed arista (frond-like structure near its proximal end) or bristle on the last segment.

Parasiticide Screening, Vol 1. DOI: https://doi.org/10.1016/B978-0-12-813890-8.00003-1 © 2019 Elsevier Inc. All rights reserved.

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Culicidae Mosquitoes (Culex, Aedes, Anopheles) Disease-vectoring mosquito species can acquire and transmit pathogens that cause human and animal disease, such as malaria, yellow fever, West Nile, Zika, Chikungunya, dengue fever, eastern and western encephalitis, and filariasis, to name a few.

Biology and life cycle The mosquito goes through four separate and distinct stages of its life cycle (complete metamorphosis): egg, larvae, pupa, and adult. How long each stage lasts depends on temperature, food availability, and species characteristics. Eggs are laid one at a time or attached together to form “rafts.” They float on the surface of water or they are placed to the borders of the potential larval site. One feature that all oviposition sites have in common is protection of the female from the action of wind and wave. In the case of Culex species, the eggs are stuck together in rafts up to 200. Anopheles and Aedes, as well as many other genera, do not make egg rafts but lay their eggs singly. Culex and Anopheles lay their eggs on the water surface, while many Aedes lay their eggs on damp soil that will be flooded by water. Most eggs hatch into larvae within 48 h or the embryo may remain quiescent for extended periods of time so that the egg will hatch only under the proper conditions; others might withstand subzero winters before hatching. High moisture and humidity are necessary features of their habitat to be suitable for hatching. The larvae are commonly known as wrigglers. The larvae live in water and come to the surface to breathe. Larvae shed (molt) their cuticle four times, growing larger after each molt, the last molt resulting in a pupa. About 7 days (but sometimes longer depending on environmental conditions and the species) are required to complete the larval stage under optimum food conditions. Most genera have siphon tubes for breathing and hang upside down from the water surface, with the exception of Anopheles larvae which do not have a siphon and lie parallel (horizontally) just beneath the water surface, suspended particularly by means of palmate hairs, to get a supply of oxygen through a breathing opening (spiracle). The larvae feed on suspended microorganisms, filtrating both plant and animals and organic matter in the water. The pupae are commonly known as tumblers. The pupal stage is a resting, nonfeeding stage of development, but pupae are remarkably mobile and sensitive to light changes and disturbances of the water, moving (tumbling) with a flip of their tails toward the bottom or protective areas. After a few moments, they rise with little motion to the surface of the water, where they breathe through a pair of breathing trumpets situated dorsally on the

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cephalothorax. The pupal stage is quite short, usually 2
3 days. The pupal stage of the mosquito changes into an adult in a process similar to the metamorphosis seen in all insects belonging to olometabola (e.g., butterflies) when the adult butterfly develops from the cocoon stage of a caterpillar. When mosquito development is complete, the pupal skin splits and the adult mosquito emerges. The newly emerged adult rests on the surface of the water for a short time to allow itself to dry and all its body parts to harden. The wings have to spread out and dry properly before it can fly. Blood feeding and mating do not occur for a couple of days after the adults emerge. The mouthparts of male mosquitoes are not suited to piercing; hence, they are not bloodsuckers. Their nourishment is normally derived from nectar and plant juices. With a few exceptions, female mosquitoes pierce the skin and feed on blood. Normally, female mosquitoes require a blood meal before oviposition, except for the first oviposition in some species (e.g., Culex molestus, Culex quinquefasciatus), in which the energetic reserves accumulated during larval development are exploited to develop the first egg raft. Also nonbiting mosquitoes belonging to Toxorhynchitinae, in which larvae are predators and adults of both sexes feed on nectars, adopt the latter strategy. Male mosquitoes usually remain alive for but 6
7 days, although longer (B1 month or slightly longer) under laboratory conditions. Females with ample food may live for several weeks, up to 2
6 months, particularly under hibernating conditions.

Rearing method(s) This procedure describes a method for the production of Aedes aegypti using a sedated or restrained live animal as a blood source. A review article by Gonzales and Hansen1 discusses older and recent studies that were aimed at the development of artificial diets for mosquitoes in order to replace vertebrate blood. Aedes aegypti Linnaeus, 1762—yellow fever mosquito Temperatures and humidity controls are probably the most important factors in the successful rearing of mosquitoes. The temperature should remain at B25 C
29 C at all times, and the humidity should remain at 70% or greater whenever adult mosquitoes are present. Photoperiod affects the development of the various stages in the life cycle of the mosquito. A. aegypti perform at an optimum level when subjected to L14:D14 hour cycle. Every reference to water which is used for the rearing process described next should be untreated (e.g., unchlorinated), using distilled or untreated well water and this water should sit for at least 24 h prior to use.

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Procedure to complete the life cycle when starting with collected eggs that have been collected on filter paper1: 1. Immerse the eggs on filter paper in a container of water. Rectangular plastic trays are excellent choices (e.g., cat litter boxes); however, clear glass containers similar to baking dishes are also useable. 2. The eggs will generally begin to hatch within 24 h. Remove the strip of filter paper containing the eggs after 24 h immersion and rewrap to use again later, since not all of the eggs will have hatched by this time. See no. 7 for method of obtaining eggs. 3. The hatched larvae should be at a concentration of one larva per 5
10 mL of water and fed daily. Fresh food should be made at least every third day. Refrigerating the food after mixing will keep it fresh. Larvae food is prepared as follows: Several protocols for preparing the food mixture are available, including ground fish food, liver powder, rodent food 1 Brewer’s yeast 1 milk protein, etc. We report next an example of one food type as a reference. Mix together ground dog food and Brewer’s yeast at the ratio of two parts dog food to one part Brewer’s yeast. Add 1 g of this food mixture to 200 cm3 of water. Feed each container of larvae B15
22 cm3 per B14
21 g of this food daily. If the larval water develops a foul odor, transfer the developing larvae to fresh water or change half of the water from the container with clean water and reduce the daily food amount. 4. The larvae should begin to pupate 4
7 days after hatching. Remove the pupae from the trays/containers described earlier using a piece of screen or a pipette. Place the pupae into a container, such as a sample specimen cup (B120 cm3); fill the cup to the top with water; and place the cup into a secure mosquito cage. 5. Adult mosquitoes emerge within 24
48 h. Provide the adult mosquitoes with sugar solution ad libitum. Prepare sugar solution as follows: Prepare a 10% sucrose (table sugar) solution (10 g sugar per 100 cm3 water). Pour the sugar solution into a vial (e.g., 7- or 20-dram vial) and fill B3/4 full. Form a wick from cotton and/or gauze and insert the wick into an opening made on the vial top. Slowly add the wick and lid to the vial. The wick should become saturated with sugar but not to the point where sugar drips onto the floor of the cage. If the sugar is allowed to drip onto the floor, the mosquitoes may get stuck in it and die. The sugar should be checked daily and changed ideally every 4 days.

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6. The mosquitoes must first be given a blood meal before eggs will be laid. Only the females will feed on blood. The mosquitoes will mate 2
3 days following their emergence as adults. The afternoon prior to the day when the mosquitoes will be offered a blood meal, remove the sugar from the cage, replacing it with a source of water for hydration. This will encourage better intake of the blood meal. The next morning, the mosquitoes should be offered the blood meal. The preferred method is to use a rabbit/guinea pig that has been shaved. The animal bait is placed on its shaved back in the mesh fabric/hammock at the top or side of the cage. If necessary, the host can be sedated according to its weight (i.e., using 0.5 cm3 of acepromazine). The rabbit should be observed at all times and held in place if necessary. The mosquitoes should be allowed to feed for B20
30 min. Following the blood meal, the sugar solution is returned to the cage. 7. To obtain the mosquito eggs, a shiny black cup containing a rolled strip of filter paper is placed inside the mosquito cage. The cup should be 1/ 2
3/4 filled with water. The filter paper should extend beyond the top of the cup by at least 3 in. Leave the cup in the cage for 4
9 days. At this time, the paper should be coated with mosquito eggs, which are black in color. 8. Remove the strip of filter paper and place it on a damp paper towel. Prepare a 0.1% solution of a viricide (e.g., Roccal-D) in water at a ratio of 1 cm3 Roccal-D to 1000 cm3 water. Using a pipette, place B1
3 cm3 of this solution onto the filter paper containing the mosquito eggs. This will prevent molding of the paper. Wrap the filter paper and paper towel in plastic wrap and seal. Label with date and species of mosquitoes. The eggs can be stored at RT for up to 6 months. The egg sheets should be checked periodically (monthly) to ensure they have not dried out. If the paper towel is dry, it should be remoistened at this time. Procedures for collecting adult mosquitoes for subsequent screening: 1. Mosquitoes are fasted (sugar solution removed) overnight prior to collection. 2. Collect the required number of unfed female mosquitoes for the screening procedure. Do not allow mosquitoes to sit for an extended period of time in the collection jar prior to use. a. Each genus and species of mosquito has various distinguishing characteristics (see Clements2 for details). Generally speaking, the following three items are true of most mosquitoes. b. Male mosquitoes tend to have very plumed antennae, whereas the females tend to have very few hairs distributed along the antennae.

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c. The males have palpi which tend to be as long as the proboscis. Females tend to have shorter palpi. A. aegypti females have very short stubby (almost unseen) palpi. d. The male proboscis may be as long as the female, but the mouthparts are usually not well developed or may even be absent. The proboscis of male A. aegypti has a “feathery” appearance. A low suction aspirator machine is used to collect the mosquitoes into a collection jar. The aspirator is fitted with plastic tubing and a 10 cm3 pipette, which has had the pointed tip removed. Use the least amount of vacuum possible when aspirating the mosquitoes to help prevent mortality. Place the pipette tip through the stockinette on the front of the mosquito cage to collect the mosquitoes. The female mosquitoes are counted as they are collected. Male mosquitoes should be avoided and are not counted if they are collected. Placing a gloved hand on the outside of the cage may help to keep the mosquitoes still as they may be attracted to the hand and will land on the cage. During and after collection of the mosquitoes, check the collection jar to ensure the mosquitoes look viable. Check the tubing to ensure mosquitoes are not stuck in the tubing, or in the connecting piece that runs from the tubing through the jar lid. Once the required number of mosquitoes is collected, turn off the aspirator. Check the tubing once again to ensure all mosquitoes are out of the line and in the jar. If mosquitoes are in the tubing, turn the aspirator back on to vacuum the mosquitoes into the jar.

A publication by Cosgrove et al.3 described a convenient, electronically controlled, in situ, membrane feeding system for feeding pair-mated A. aegypti with defibrinated, refrigerated pig blood. This membrane system demonstrated no significant difference from mouse-fed controls in the rate of egg maturation, fecundity, or pupal yield. This is an improvement of membrane-fed mosquitoes that showed a lower degree of fecundity and fertility than those fed on live animals.4

In vitro method(s) Larvicidal assays As specified in the WHO Guidelines for Laboratory and Field Testing of Mosquito Larvicides,5 the objective of laboratory testing is to evaluate the biological activity of a mosquito larvicide. Laboratory-reared mosquito larvae of known age or instar are exposed for 24
48 h or longer in water treated with the potential larvicide at various concentrations within its activity range, and mortality is recorded. It is important to use homogeneous populations of mosquito larvae or a given instar. The aims of laboratory tests are

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to establish dose
response line(s) against susceptible vector species; to determine the lethal concentration (LC) of the larvicide for 50% and 90% mortality (LC50 and LC90) or for 50% and 90% inhibition of adult emergence (IE50 and IE90); to establish a diagnostic concentration for monitoring susceptibility to the mosquito larvicide in the field; and to assess cross-resistance with commonly used insecticides.

The WHO guidelines describe the following assay to determine biological activity against mosquito larvae. Materials required for testing G G G G G G G

One pipette delivering 100
1000 μL Disposable tips (100, 500 μL) for measuring aliquots of dilute solutions Five 1 cm3 pipettes for insecticides and one for the control Three droppers with rubber suction bulbs A strainer Disposable test cups Graduated cylinder

Preparation of stock solutions or suspensions and test concentrations The technical materials of many organic compounds are insoluble in water. These materials have to be dissolved in appropriate organic solvents, such as acetone or ethanol (or DMSO), in order to prepare dilute solutions for laboratory testing. The formulated materials are, however, miscible with water. Suspending or mixing these formulations in water requires no special equipment, homogeneous suspensions can be obtained by gentle shaking or stirring. The volume of stock solution should be 20 cm3 of 1%, obtained by weighing 200 mg of the technical material and adding 20 cm3 solvent. It should be kept in a screw-cap vial, with aluminum foil over the mouth of the vial. Shake vigorously to dissolve or disperse the material in the solvent. The stock solution is then serially diluted (10-fold) in ethanol or other solvents (2 cm3 solution to 18 cm3 solvent). Test concentrations are then obtained by adding 0.1
1.0 cm3 (100
1000 μL) of the appropriate dilution to 100 or 200 cm3 chlorine-free or distilled water. When making a series of concentrations, the lowest concentration should be prepared first. Small volumes of dilutions should be transferred to test cups by means of pipettes with disposable tips. The addition of small volumes of solution to 100, 200 cm3, or greater volumes of water will not cause noticeable variability in the final concentration. When a test is carried out using formulated materials, distilled water is used in the preparation of the 1% stock solution or suspension and in subsequent serial dilutions, according to the content of the active ingredient. WHO bioassay Initially, the mosquito larvae are exposed to a wide range of test concentrations and a control to determine the activity range of the

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materials under test. After determining the mortality of larvae in this wide range of concentrations, a narrower range (of 4
5 concentrations, yielding between 10% and 95% mortality in 24 or 48 h) is used to determine LC50 and LC90 values. Batches of 25 third or fourth instars are transferred by means of a strainer or dropper to small disposable test cups or vessels, each containing 100
200 cm3 water. Small, unhealthy, or damaged larvae should be removed and replaced. The depth of the water in the cups or vessels should remain between 5 and 10 cm; deeper levels may cause undue mortality. The appropriate volume of dilution is added to 200 cm3 water in the cups to obtain the desired target dosage, starting with the lowest concentration. Four or more replicates are set up for each concentration and an equal number of controls are set up simultaneously with tap water, to which 1 cm3 alcohol (or the organic solvent used) is added. Each test should be run three times on different days. For long exposures, larval food should be added to each test cup, particularly if high mortality is noted in the control. The test containers are held at 25 C
28 C and preferably a photoperiod equal to the one used for the rearing of the mosquito strain used (e.g., 14 h light followed by 10 h dark). After 24 h exposure, larval mortality is recorded and confirmed after 48 h. For slow-acting insecticides, 48 h assessment is required. Moribund larvae are counted and added to dead larvae for calculating mortality. Dead larvae are those that cannot be induced to move when they are probed with a needle in the siphon or the cervical region. Moribund larvae are those incapable of rising to the surface or not showing the characteristic diving reaction when the water is disturbed. Larvae that have pupated during the test period will negate the test. If more than 10% of the control larvae pupate in the course of the experiment, the test should be discarded and repeated. If the control mortality is between 5% and 20%, the mortalities of treated groups should be corrected according to Abbott’s formula6: mortality (%) 5 [(X 2 Y)/X] 3 100, where X is the percentage survival in the untreated control and Y is the percentage survival in the treated sample. A paper published by Pridgeon et al.7 described the development of a high-throughput assay, using 96-well plates and the first instars of A. aegypti, thus expediting the screening process and allowing several hundreds of compounds to be easily tested in a short time period. This assay is a sensitive predictor of general ectoparasiticidal activity. In addition, Derua et al.8 described the effect of ivermectin on the third instars of Anopheles gambiae Giles, 1902 and C. quinquefasciatus Say, 1823. Adulticidal flying insect spray assays9 Glass chamber: This test is conducted in a glass chamber measuring 70 3 70 3 70 cm. A total of 20 laboratory-reared sucrose-fed adult female

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mosquitoes, 2
5 days of age, are released into the chamber. The insecticide is sprayed into the chamber by using a manual/electric atomizer. The discharge rate (g/spray) of the sprayer is predetermined. Based on the dosage required, an estimated time of spray is discharged into the glass chamber. Knockdown of mosquitoes is observed at the indicated intervals up to 20 min. After 20 min, all mosquitoes are then collected and placed in cylindrical polyethylene containers with a 10% sucrose pad. A further, single assessment of knockdown is made after 60 min. Mortality is observed after 24 h PT. All tests are to be conducted at a temperature of 26 C
28 C and RH of 65%
85%. A minimum of three tests are conducted. The knockdown values (KT50 and KT95) and regression slope are obtained using probit analysis. Mean percentage of insect mortality value is subjected to arcsine transformation followed by comparison of means using the LSD test. Peet-Grady chamber: This test is conducted in a Peet-Grady chamber measuring 180 3 180 3 180 cm. A total of 50 laboratory-reared sucrose-fed adult female mosquitoes, 2
5 days of age, are released into the chamber. The insecticide is sprayed into the chamber by using a manual/electric atomizer. The discharge rate (g/spray) of the sprayer is predetermined. Based on the dosage required, an estimated time of spray is discharged into the PeetGrady chamber through two introduction ports of the chamber. Knockdown and mortality are observed as described earlier (glass chamber). Adulticidal repellency assay The most common goal in mosquito control is to prevent the mosquito female from landing and/or biting the host and its possible transmission of a disease agent. Thus there are numerous articles found in the literature that describe various in vitro bioassays for the evaluation of adult mosquito repellency and fewer articles in the literature on the evaluation of adulticidal agents against female mosquitoes biting the host. A number of in vitro assays for the quantitative evaluation of mosquito repellents with and without the use of blood cells have been reported in the literature. Many of these in vitro bioassays are intended for the discovery of novel human-use mosquito repellents. An article published by Huang et al.10 described the development of an efficient and safe in vitro bioassay system using a multiple-membrane blood-feeding device and a cocktail meal. The multiple-membrane blood-feeding device facilitated the identification of new insect repellents by the high-throughput screening of candidate chemicals. A cocktail meal was developed as a replacement for blood for feeding the female A. aegypti. The cocktail meal consisted of a mixture of salt, albumin, and dextrose, to which adenosine triphosphate was added to induce engorging. The feeding rates of A. aegypti on the cocktail meal and pig blood, respectively, did not differ significantly but were significantly higher than the feeding rate on citrate phosphate dextrose-adenine 1 solutions, which had

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been used to replace blood meals in previous repellent assays. Dose-dependent biting inhibition rates were analyzed using probit analysis. The RD50 (the dose producing 50% repellence of mosquito feeding) values of DEET, citronella, carvacrol, geraniol, eugenol, and thymol were 1.62, 14.40, 22.51, 23.29, 23.83, and 68.05 μg/cm2, respectively. This in vitro assay could be used to identify new repellents having potential use as mosquito repellents on animals. An article by Klun et al.11 described a new in vitro bioassay system for the discovery of novel human-use mosquito repellents. The authors adapted an earlier test module (that was used for quantitative measurement of the efficacy of mosquito repellents on human volunteers), by coupling the module with a membrane
blood reservoir. The module consisted of six test cells and permitted the simultaneous comparison of up to five repellent doses or chemical types and a control using a complete randomized block design with minimal treatment interaction and with $ 6 replicates per human subject. Performance of insect repellents in the new in vitro was compared with their performance on humans against mosquitoes. For each compound, in vitro dose
response assays were conducted with compounds applied to cloth positioned over blood reservoirs covered with Baudruche membrane against A. aegypti. The repellents were also tested in vitro against Anopheles stephensi Liston, 1901 and A. aegypti at a fixed dose of 24 nmol compound/cm2 cloth over the Baudruche and Edicol collagen membranes. Concurrently, the repellents were tested at the fixed dose using human volunteers. The observed proportions of both mosquito species deterred from biting in the fixed doses in the in vitro assays were similar to those obtained using humans, being clearly able to distinguish controls from repellents and differing only in the ranking of the effectiveness of some of the repellents. Dose
response relationships of the in vitro and in vivo systems were also very similar, although not directly comparable because the data were not collected concurrently. This new in vitro assay system can be used in high-throughput screening of compounds to identify new repellents having potential for use as repellents on animals, as well as humans. Grieco et al.12 described, in great detail, a modular and novel assay system for rapid mass screening of chemical compounds for spatial repellent actions (as well as contact irritant actions) against adult A. aegypti. This system detected contact irritancy and spatial repellency activity with reproducible results and provided baseline data for determining minimum effective concentrations for other chemicals. The system was compact in size, easy to decontaminate after use, and required only a minute quantity of chemical compound for testing. Adult mosquito contact assay7 The purpose of this procedure is to describe a contact assay using A. aegypti. G

Fifty cubic centimeter conical centrifuge tubes (Falcon) are sealed with gauze held with an elastic band and 100 μL of a 1% solution is dispensed on to the gauze and air dried for 15 min in a fume hood.

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Eight to 10 mosquitoes (previously immobilized on ice) are placed in the tube and the treated gauze replaced. The tubes containing the mosquitoes are placed in the insectary at 28 C. The mosquitoes are encouraged to rise to the top of the tube and sit on the gauze with a CO2 stimulus caused by breathing over them. Mortality is assessed at 60 min. Actives are compounds which prevented 70% of the mosquitoes from sitting on the gauze. Knockdown effects are also recorded. All dose titrations are performed at least twice to confirm activity.

Additional contact assays have been described in the literature, including tarsal contact with treated filter paper, mosquito net impregnation, and cone bioassays by Darriet et al.13 as well as toxic sugar baits.14 In addition, there are a number of mosquito studies using wind tunnel, olfactometer, and electroantennogram assays, in particular for pheromones (semiochemicals), and the reader is directed to Chapter 3c, Arthropoda, Diptera, Cyclorrhapha, on Cyclorrhapha for examples of such assays.

In vivo method(s) Laboratory efficacy test of cutaneous repellent product on human hosts This protocol is based on WHO guidelines15 for testing products for human use against mosquitoes, but it can be generalized for other vectors of medical interest. The main objective is to estimate both the effective dose (ED50 and ED99.9 protection from mosquito landing and/or probing) and the complete protection time (CPT, the time before the first mosquito landing and/or probing) provided by the repellent product. The test cage where the insects will be released should have a metal frame for ease of decontamination, with an expected size of about 40 cm per side. The bottom and top should be solid, while the back should be of net. The lateral sides should be transparent for viewing (e.g., made with acrylic sheet), and on the front, there is a fabric sleeve for access of the hand and specimens. Before the test, the skin area is washed with unscented soap, rinsed first with water, followed by 70% ethanol or IPA and dried. Mosquitoes (or other target insect species) should be host-seeking and of uniform age (e.g., 5
7 days postemergence). Serial dilutions of repellent are made with ethanol or another diluent and tested to identify an effective dose range between 10% and 90% (2
3 dosages below and above 50% repellent response, respectively). Five successive applications of increasing dose of repellent are used on the test forearm by the host, starting from the lower dose to the higher one. A single test comprises continuous use of the same insect by the same volunteer and is completed in one day.16 At least three test replicates are needed, repeating this process using different batches of insects over several days. One mL of ethanol or the same diluent used in the preparation of the

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test repellent is applied evenly using a pipette to B600 cm2 of the skin and allowed to dry. Before insertion of the arm into the cage containing 50
100 host-seeking insects previously released, the nontest skin area is protected by a material through which the insects cannot bite. The first step is to insert the forearm applied with diluent into the cage and to count the number of insects that land on and/or start probing the skin during a 30-s period. During testing, the volunteer should avoid movement of the arm. For the test to proceed, the biting rate must be $ 10 landings and/or probings in the 30 s period. After the control test, the same arm is treated with the lowest dose of repellent in 1 mL diluent and allowed to dry and a test is performed as earlier. This procedure is repeated for each additional incremental repellent dose-repeating test in sequence without delay among them. The repellent dose at each test is calculated as the sum of the doses applied to arrive at the cumulative dose for each test. As a final control of the host-seeking activity of the insect tested at the conclusion of the dose
response experiment, 1 mL of alcohol/diluent is applied on the other forearm and allowed to dry. This forearm is inserted in the cage for 30 s to verify that the number of landings and/or probings is B $ 10/30 s, as was observed at the beginning of the experiment. If the rate is ,10 insects in 30 s, the whole experiment should be discarded. Protection (p) is expressed as a proportion of the number of landings and/ or probings on the treated arm (T) in relation to the number of landings and/ or probings on the control arm (C) of the same individual:   T ðC
T Þ p51
ð3a:1Þ 5 C C where C is the average of the landings/probings on the two untreated arms (the diluent applied at the beginning and at the end of the experiment). Data are analyzed by probit-regression analysis from which the ED50 and ED99.9 and their confidence limits can be estimated.16 After this test, the estimation CPT can be determined starting from the ED99.9 calculated earlier. One mL of the repellent at the ED99.9 level is compared to 1 mL of a standard repellent as a positive control (e.g., 20% ethanolic DEET). Two mosquito cages (as described earlier) each containing 200
250 nonblood-fed insects are prepared: one for testing the candidate repellent and the other for the positive control. During testing, the nontest skin area is protected from bites as earlier. As a first step, the willingness to bite of the tested insects is assessed by inserting an untreated arm into the two cages for 30 s or until 10 landings/probings are counted. If this level of landing and/or probing is not achieved in either cage, the experiment should be discarded. Thirty min before starting the test, 1 mL of the candidate repellent prepared in alcohol/diluent solution is applied to one arm and 1 mL of DEET standard solution is applied to the other arm. Then, the new repellenttreated arm is inserted into the cage and exposed for 3 min to determine

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landing and/or probing activity. Subsequently, the DEET-treated arm is exposed. This procedure is repeated at 30 or 60 s intervals and should be used consistently throughout the experiment. The occurrence of at least one landing and/or probing in a 30 s test interval concludes the test for that repellent dose. CPT is calculated as the number of min elapsed between the time of repellent application and the first mosquito landing and/or probing. The median CPT and confidence intervals can be estimated from the Kaplan
Meier survival function.

Laboratory artificial challenge on animal hosts in cages Several studies may require the direct challenge of laboratory-reared hematophagous insects on a host (e.g., a dog), in particular those involving cutaneous products with insecticidal/repellent activity. In these studies, antifeeding and mortality rates can be measured. Laboratory-reared insects of the same age (e.g., 2
5-days old for mosquitoes and sand flies) are deprived of sugar 24 h before the test, giving only water to prevent dehydration. A defined number of insects (generally 50/animal host) are transferred in separated bottles or cups per each host and transported from insectary to the challenge room. The sedated host is placed in the test cage lying on its side before the release of the insects on it. All the cages of the tested animals are located in the same challenge room, setting the temperature between 20 C and 30 C. Release the precounted insects in the cage leaving them free to exit spontaneously from the container. Remove and (possibly) replace immediately any moribund/dead specimens. Exposure time may vary according to protocol specifications (30
60 min in general). After the exposure time, alive insects are collected from the cage (by mouth aspirator or electric pooter). Moribund/Dead insects are collected separately and counted. According to the test purpose (antifeeding or insecticidal activity of the compound tested, see next), the proportion of fed/alive specimens per cage is compared between test and control animals. Identification of feeding status of specimens is done by a stereomicroscope, if needed, including fed and partially fed specimens in the same group as opposed to unfed. The same is for moribund/dead as opposed to alive specimens. Moreover, a specific repellent test based on a mice host and mosquitoes is presented, but it can be adapted to other hosts and insects.17 Five hosts are randomly treated with one control solution (ethanol or other solvents) and four serial dilutions of the test repellent in ethanol. Treatments are applied with a pipet over the whole body, if possible. Hosts are transferred to a test cage (as described earlier) containing 100 mosquitoes (or other host-seeking species). The number of insects biting each host is recorded at 2-min intervals for a 30-min period. Total biting events are cumulated at the end of the test. Dose ranges are selected according to the concentration of the repellent,

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from the lower to the higher. At least three replicates are needed for each dose. The analysis can be done as described for human repellent tests.

Simuliidae Simulium spp. Latreille, 1802—black flies, buffalo gnats, turkey gnats Bloodsucking black flies are a serious problem of worldwide medical, veterinary, and economic significance. Adult female simuliid species can acquire and transmit pathogens, such as nematodes, protozoans, and viruses, that cause human and livestock diseases such, onchocerciasis, leucocytozoonosis, vesicular stomatitis, and trypanosomiasis. Other species are voracious biters that can cause discomfort and annoyance (nuisance), morbidity, and even death.

Biology and life cycle Black flies go through four separate and distinct stages of its life cycle (complete metamorphosis): egg, larvae, pupa, and adult. The length of each stage depends on both regional temperature and species characteristics. There can be anywhere from one to six generations in a year. Eggs usually numbering 200
500 eggs per female and for most species are deposited in or on flowing water. Some females attach them to wet surfaces, such as blades of aquatic grasses, logs, or water-splashed rocks. A common method is for the female to drop eggs while flying over the water surface; some species will hover and oviposit through a thin film of water that covers sand, rock, or vegetation; others will settle and oviposit on waterlapped surfaces or on surfaces at the water’s edge. As for several other insect groups, the shiny eggs are at first creamy white, changing to almost black when the chorion is sclerotized. The black fly usually spends the winter not only in the egg stage but also in the last larval stage as described next. The length of time it takes an egg to hatch varies greatly from species to species. Eggs of most species hatch in 4
30 days, but those of certain species may not hatch for a period of several months or longer. The duration of larval development ranges from a few days to 9 months, depending in part on water temperature and food supply. The last instar can pass through winter attached underwater to rocks, driftwood, and man-made concrete surfaces, such as dams and channels. The number of larval stages ranges from four to nine, with seven being the usual number. Larvae remain attached to stationary objects in flowing water, held on by silken threads extruded from glands located at the end of the bulbous abdomen (caudal sucker). Depending on the species, mature larvae range from 5 to 15 mm in length and may be brown, green, gray, or nearly black in color. They possess

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a large head that bears two prominent structures known as “labral fans” that project forward. Labral fans are the primary feeding structures, filtering small crustaceans, protozoa, algae, bacteria, and organic debris out of the water current. The pupal stage is formed in the following spring or summer, typically in the same site as the last instar, but may occur downstream following larval “drift” with the current. They are typically orange and appear mummy-like because the developing wings and legs are tightly attached to the body. Pupae in many species produce a delicate, silken “cocoon” of varying density, weave, and size that partially or nearly encloses them; other species produce hardly any cocoon at all. The pupal period is quite short in some species, requiring 2
6 days; in others, it may last 3
4 weeks. Temperature influences the duration of this stage with cooler temperature retarding the emergence of adults. Adults emerge from the pupal stage in 4
7 days and can live for 2
3 weeks. When the adult is ready to emerge, it can be seen through the translucent pupal integument. In B1 min, the fly breaks through the T-shaped emergence slit and rises to the surface quickly in a bubble of gas. Adult black flies are small insects that measure 1
5 mm in length and possess a shiny thorax that ranges in color from black to various shades of gray, or brown or yellow. The female fly has bladelike and piercing mouthparts that are more or less rudimentary in the male. Nectar from flowers provides both males and females with carbohydrates for flight energy and females, in addition, usually require a blood meal for ovarian development. Males never suck blood, whereas most females are vicious bloodsuckers. Males may form a hovering swarm, and mating occurs when females fly into or near such swarms. At other times, however, mating occurs apparently by accidental contact on the ground or elsewhere, without the formation of swarms.

Rearing method(s) The collection and rearing of black flies have been under study for many years and have long been impeded because of the difficulty in working with them in the laboratory. Earlier attempts were made, without success, to establish continuous colonies of blood-feeding adult black flies. Adults, however, were successfully reared from eggs, first instars or pupae. In order to overcome the earlier difficulties of colonizing simuliids in the laboratory, membrane systems were developed to feed the adults on whole blood. A method of rearing large numbers of black flies by Bernardo and Cupp18 described a highly efficient, mass-scale in vitro membrane feeding system for black flies that was proven effective for five Simulium species exhibiting a wide range of blood-feeding preferences. Factors affecting the feeding rate, such as type of membrane or blood used, addition of ATP, age of flies, and temperature differential, were examined for the species tested.

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Gray and Noblet19 have maintained and operated the only colony of black flies (Simulium vittatum Zetterstedt, 1838) in the world for over 26 years, the past 18 years at the University of Georgia (UGA) Black Fly Rearing and Bioassay Laboratory. They provided detailed descriptions of the aquatic rearing units and all associated systems. In addition, the operation and maintenance of all systems were described as well as troubleshooting measures. The UGA colony was initiated in 1991 in collaboration with Dr. Ed Cupp’s group at the University of Arizona and now maintains the only version of Cupp’s initial efforts at Cornell in 1981. The primary purpose of the colony is to produce uniform larvae for a variety of research activities, including product development and a larval bioassay associated with insecticidal proteins of Bacillus thuringiensis subsp. israelensis. The UGA colony is operated in similar fashion to the Cornell Automated Rearing System. This system incorporates a closed-circulation trough system in which water is pumped over a wooden runway creating ideal larval habitat. Each week, B200,000
300,000 larvae are produced. The continuous rearing process of black flies is extremely difficult and time-consuming and requires a number of trained personnel. For this reason, it appears that the UGA black fly rearing laboratory is currently the only one-of-a-kind operation. The UGA colony is operated with associated systems that automatically feed the larvae, capture emerging adults and provide suitable substrates for oviposition. Larval feeding is accomplished by grinding a 1:1 mixture of rabbit chow and soybean meal and washing it through a 53 μm sieve (No. 270). The resulting food slurry is stored in tanks that are kept in modified refrigerators adjacent to the rearing units. Each tank includes two submersible pumps, one to stir the food solution and one to pump it into the rearing unit. Larval development occurs on a wooden runway (0.3 m 3 1.2 m). Upon significant pupation, an emergence hood is placed over the rearing unit to capture emerging adults and initiate the mating process. Adults move toward the light coming through a glass funnel located at the apex of the hood and the associated piece of Tygon tubing that is attached. Mating occurs in this emergence tube and a smaller mating tube where all adults that have been collected during the day are confined. Adults are provided 10% sucrose and distilled water via cotton pads that are placed on the screened portion of the adult container. The oviposition process is initiated the following week when the adults are removed from refrigeration and egg development is allowed to resume. After 24 h, adults are released into insect cages (0.3 m 3 0.3 m 3 0.3 m) that are situated over disks of cloth (0.15 m diameter, light green) which are bathed in a film of water pumped from 38 L aquariums via submersible pumps and a tubing apparatus. The insect cages are covered in black cloth, so the only light available is from a fluorescent light, on the bottom edge of the oviposition platform, below the disks. Females are attracted to the

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moisture and light and lay their eggs on the moist, disks of cloth. Egg sheets are pinned to the upper surface of a clean runway and the rearing cycle continues. The development and maintenance of this black fly colony have eliminated the need to obtain eggs and larvae from the environment and reduce the potential for experimental variability. An excellent review article on rearing black flies was published by Edman and Simmons.20

In vitro method(s) Larvicidal assays Black fly larvae are the most commonly used stage for in vitro screening purposes. The two methods briefly described next use larvae derived from existing closed systems. A scientific note by Carlos et al.21 described a method for the evaluation of black fly larvicides that was easy to maintain and low on special requirements. Healthy larvae selected for an evaluation were separated in groups $ 50 larvae, and each group was then placed in a 300 cm3 plastic container (5recipient) filled with 200 cm3 of water. The water must be from a natural breeding site in order to eliminate the need of adding food into the system. Each recipient must have a pump for circulation of water to allow the larvae to filter feed. Each recipient filled with larvae was accompanied by another recipient filled with the same water volume, but without the larvae. Since the recipients are closed systems, the larvae must be removed from the original recipient container after the desired exposure time to the larvicide and placed in the corresponding recipient without the larvicide. The same procedure must be applied to the control group, to standardize any side effects that the transference may have on the larvae. Therefore this method allowed the adjustment of the desired exposure time in the laboratory and thereby presented a simpler methodology for the reproduction of field conditions. The orbital shaker bioassay developed at Clemson University as described in Gray and Noblet19 has been used for subsequent bioassay evaluations. Overmyer et al.22 determined that fipronil was more toxic than imidacloprid to a colony of simuliid larvae in a 48 h orbital shaker bioassay. Overmyer et al.23 followed this work, demonstrating that the enantiomers of fipronil did not produce significantly different levels of mortality in a 48 h orbital shaker bioassay using the colony larvae. In this study, the toxicity of fipronil and its enantiomers to S. vittatum was assessed using a 48-h modified orbital shaker toxicity test. Larval mortality was assessed at 20 3 magnification due to observations made during the previous testing with fipronil

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in which slight twitching movements of the labral fans were difficult to observe with the naked eye. Six insecticide concentrations were prepared in 100 cm3 volumetric flasks by spiking test water with insecticide stock solution. The maximum volume of stock solution used for preparation of the treatment solutions was 100 μL (0.07%). The contents were emptied into 250 cm3 amber bottles until treatment. During treatment, 5 cm3 of the prepared solutions was added to the appropriate flasks creating test concentrations of 2.0, 1.0, 0.5, 0.25, 0.125, and 0.06 μg/L inside the flasks. Approximately 300 cm3 of spiked water was collected before and after each test from the highest and lowest concentrations tested and stored in amber bottles with Teflon lids at 220 C for analysis of fipronil concentrations. Six insecticide concentrations and two controls [water and a carrier (acetone control)] were tested on one shaker with 5 flasks/concentration and control, bringing the final totals to 40 flasks and 600 larvae. The carrier control had an acetone content equivalent to the maximum volume of stock insecticide used in the preparation of the treatment solutions. Each of the insecticide concentrations was tested on a separate shaker on the same day and replicated three times over the course of 3 weeks. Tests were conducted with an ambient air temperature of B20 C and a 16L:8D hr photoperiod. Tests were considered valid if control mortality was ,10%. Water quality (temperature, dissolved oxygen, conductivity, and pH) was measured in the controls before and after each evaluation. The orbital shaker bioassay has also been used to evaluate the efficacy of B. thuringiensis
produced insecticide proteins.

Adulticidal repellency assays Bernardo and Cupp18 describe preliminary experiments to test various commercial black fly repellents using an in vitro membrane feeding system. The design of a membrane feeding apparatus enabled large numbers of females to feed simultaneously. Because of their ease of preparation, in comparison to 2-week-old chicks, or Badruche membranes, stretched Parafilm membranes were used for the screening of repellents with this feeding apparatus. Membrane feeding apparatus An incubator served as the basic frame with three holes cut in the side to allow passage of tubing and electrical cords. An immersion heater-recirculator was placed in a 15-L plastic bin filled with water outside of the incubator, and Tygon tubing from the recirculatory was passed into the interior of the incubator to connect with water-jacketed glass membrane feeders.24 Stretched Parafilm as well as two other membrane types were used to cover the bottom of the feeder and were held in place with small rubber bands. A humidifier was placed at the bottom of the incubator, but rarely used, since RH had no apparent effect on feeding rates.

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Feeding cages for female black flies were constructed of plastic or Plexiglas cylinders 2 cm tall, with a diameter slightly larger than that of the glass feeders. The top and bottom of the feeding cages were covered with polyester screening large enough for flies to extrude their mouthparts but preventing their escape. Mesh size varied according to fly species. A coverable hole bored into the side of the cage (cylinder) allowed for the transfer of flies. Laboratory-reared females were collected on the day of emergence and were aspirated into a 0.5-L cardboard cage with screened ends. Cotton pads soaked in 10% sucrose and tap water were offered to the flies, and the cages were placed in polyethylene bags (to maintain humidity) and placed in a refrigerator at 7 C. Before feeding, females were removed from cold storage and kept 4
6 h at ambient temperature in the dark without sugar solution or water. They were next transferred, without anesthetization, from holding cages to feeding cages. A set number of females were put into each feeding cage and the glass feeders were lowered directly on top of the cages and clamped so that the membrane firmly contacted the mesh screening. Blood (or distilled water) was pipetted into the central chamber of the feeder (the bottom which was covered by the membrane). The heaterrecirculator warmed the water to 38 C
39 C, prior to the introduction of the blood or distilled water. The temperature inside the incubator was maintained at 16 C
17 C for the feeding process. Flies were usually given the opportunity to engorge for B2 h and after feeding, the flies were anesthetized with CO2, and removed. For studies in which the number of bites through the membrane was recorded, the feeder was emptied of blood or distilled water and turned upside down to examine the membrane surface with a stereomicroscope at 40 3 magnification. Individual punctures were readily visible at this magnification. The term “biting” is used to denote touching of the mouthparts to the membrane following the piercing of the membrane and presumably tasting the fluid. In addition, the number of blood-fed flies can also be determined by visually examining the females for engorgement. Factors affecting biting and engorgement were analyzed using chi-square determinations or Pearson correlations. In order to use this system for the evaluation of fly repellents, the following steps were performed. A line was drawn down the center of the Parafilm feeding membrane; half was treated with a repellent while the other half was untreated. Six membranes were treated in this way with six commercial repellents. The entire experiment was replicated three times. Each repellent had the same basic components, with varying amounts of the active ingredients. Adult female Simulium pictipes Hagen, 1880 were used for these experiments, with procedures for handling and feeding identical to those described earlier. Feeding began B4 h after treatment of membranes with repellent

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and lasted for 2.5 h. At the end of the experiment, feeders were emptied and the membranes were examined at 40 3 magnification for bites. The number of bites through the untreated control portion (Bc) versus the number through the treated portion (Br) of the membranes was recorded and compared using the following formula: percent reduction of biting activity 5 100 2 [(Br/Bc) 3 100]. Percent reduction of biting was plotted on the probit scale versus the logarithm of the percent active in the repellents tested (i.e., DEET in this study).

In vivo method(s) Laboratory efficacy test of cutaneous repellent product on human host The protocol described earlier for mosquitoes can be also used for black flies. Laboratory artificial challenge on animal host in cages The protocol described earlier for mosquitoes can be also used for black flies.

Ceratopogonidae Culicoides spp. Latreille, 1809—biting midges, “no-see-ums,” punkies Many species of Culicoides never attack man but certain ones constitute a serious economic problem by their annoyance to campers, swimmers, and fishermen. In addition to the annoyance factor, these bloodsucking midges have known roles in the transmission of human and animal pathogens, with at least five helminths involved (e.g., Onchocerca reticulata, the causative organism of fistulous withers in horses). Several protozoan and viral diseases of domestic and wild animals, poultry and waterfowl, and possibly man are related to ceratopogonid transmission (e.g., bluetongue).

Biology and life cycle In most Culicoides species, the life cycle is poorly understood. During the summer months, the development from egg to adult occurs in 2
6 weeks. Some larvae and pupae overwinter in protected breeding places and continue development in warmer weather. Culicoides species undergo complete metamorphosis. Adult midges usually live for B20 days to more than 90 days, depending on ambient conditions. The adults fly and copulate in swarms. The mean distance for female flight is 2 km, less than half of that distance for males. However, these species can exploit wind to increase their dispersal. Eggs can be cigar, banana, or sausage shaped and B0.25 mm long. They are white when first laid, but later turn brown or black. The eggs are laid on moist soil and cannot withstand desiccation. Some species can lay up to

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450 eggs/batch and as many as 7 batches in a lifetime. The number of eggs produced varies among species and size of blood meal. Eggs typically hatch within 2
10 days of being laid; time to hatch is dependent on the species and temperatures. The larvae are worm-like, creamy white, and B2
5 mm long. Larvae develop through four instars; the first instar possesses a functional spinebearing proleg. Larvae require water, air, and food and are not strictly aquatic or terrestrial. They cannot develop without moisture. The larvae are present in and around salt marsh and mangrove swamps, on shores of streams and ponds, and in muddy substrates. They feed on small organisms. Most species cannot exist more than a few inches below the air
water interface. In tropics, the larval habitat of many species is in rotting fruit, bromeliads, and other water-holding plants. Other habitats include tree holes and slime-covered bark. While some larvae can develop in wet manurecontaminated areas, they do not develop inside the animal. The larvae also do not develop inside humans or other animals. The pupal stage typically lasts B2
3 days. Pupal color can be pale yellow to light brown to dark brown. They are 2
5 mm in length with an unsegmented cephalothorax that has a pair of respiratory horns that may bear spines or wrinkles. During this stage, the insects possess a spiny integument which can be used to identify the fly to species level. The adult “no-see-ums” are gray and ,3.17 mm long. The two wings possess dense hairs and give rise to pigmentation patterns which are used to identify species. The mouthparts are well developed with cutting teeth on elongated mandibles in the proboscis of bloodsucking females, but not in males. Male Culicoides typically emerge before the females and are ready to mate when the female emerges from the pupal stage. Mating typically occurs in flight when females fly into swarms of males and the insects are oriented end to end with the ventral parts of the genitalia in contact. Some species mate without swarming; instead, the males go to hosts the female is likely to feed on blood; mating occurs when she finishes feeding. Both males and females feed on nectar, but the females require blood for their eggs to mature. The females will blood-feed primarily around dawn and dusk; however, there are some species that prefer to feed during the day.

Rearing method(s) The procedures and equipment developed by the Arthropod-Borne Animal Diseases Research Laboratory, USDA-ARS, Laramie, WY to produce and maintain immature (eggs, four instars and pupae) and adults of Culicoides sonorensis Wirth & Jones, 1957 have been described in detail by Hunt.25 A generalized version of these rearing procedures, described in detail by Hunt et al.,26 follows next.

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Production of eggs Blood feeding of laboratory-reared female C. sonorensis is done at least 3 days/week. Males and females (24
48-h old) are anesthetized for B15 s with CO2; adults usually recovering within a few minutes. Up to 5 cm3 (B5000 adults) is added to each blood-feeding cone. An artificial bloodfeeding apparatus27 allows the females access to the blood for at least 3 h. The apparatus consists of a water-jacketed glass cylinder with a reinforced silicone feeding membrane attached to the central well where B10 cm3 of sheep blood is mechanically stirred and maintained at 36.5 C, the natural skin temperature of sheep. Insectary personnel periodically exhale on the cones for CO2 to stimulate blood-feeding activity. After blood-feeding, the engorged females along with the males and any unengorged females are maintained up to 4 days in adultholding cages at 27 C 6 1 C, 40%
50% RH, and 13L:11D h photoperiod. An adult-holding cage consists of a 3.8-L cardboard container and lid, fine mesh polyester organdy, clear plastic film, plastic container with an oviposition substrate (filter paper positioned on top of moistened sterile cotton), and three vials and dental cotton wicks used to hold 10% sucrose solution and DI distilled water. Up to 70,000 eggs are collected from each cage during the 2
4 day oviposition period. Oviposition substrates contain thousands of eggs and are stored in a small amount of DI water inside Petri dishes at 4 C. This water provides a temporary rearing medium for first instars if the eggs hatch. The eggs may be stored in the refrigerator for up to 3 days; however, the viability and hatchability of the eggs under refrigeration decrease significantly after 30 days. Production of larvae Insect-rearing pans are prepared once per week. A 3700 cm2 rearing pan consists of two Dacron islands used as substrates for the immature stages, two plastic paddles used to slowly circulate the rearing medium and to minimize the formation of bacterial scum, and four metal bars used to hold the islands in place during the movement of the medium. After the rearing pans have been assembled, B9.5 L of DI water, 6 cm3 of bacterial inoculum, 4 cm3 of nutrient broth fluid concentrate, 3 g of “Kalf” medium (composed of 140.0 g high-protein supplement, 135.0 g alfalfa, 10.0 g albumin, 10.0 g brain heart infusion medium, and 10.0 g yeast extract medium), and 10.0 cm3 algicide are added to each pan. The islands must be located at the water surface so that the larvae can maneuver easily on and off the islands. The C. sonorensis larvae feed on the various microorganisms contained in the inoculum and on the subsequent formation of detritus. Oviposition papers containing B10,000 eggs are placed on both islands. Because the viability and hatchability of the eggs

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significantly decrease during cold storage, the appropriate number of eggs must be considered during the setup of rearing pans, large and robust adults are produced when the optimal level of B10,000 larvae is achieved for each pan. For maintenance of the rearing pans, 4 cm3 of nutrient broth fluid concentrate is added to each pan every other day for B24 days. If an investigator requires a particular instar, then B2000 fourth instars/1 cm3 are collected using a No. 70-size sieve, B3000 third instars/1 cm3 are collected using a No. 80-size sieve, or B8000 first and second instars/1 cm3 are collected using a No. 140-size sieve. Production of pupae C. sonorensis pupae are collected 4 days/week to produce a continuous supply of adult biting midges. Flotation screens are placed on each island containing pupae so that the islands become submerged. Pupae and larvae float to the water surface and congregate primarily along the edges of the pan. These are aspirated using a vacuum pump device and then poured into a No. 40-size sieve to separate the larvae from the pupae. The larvae are transferred back to an established rearing pan. All screens must be removed and the islands must be properly refloated following the pupae collection. About 4 cm3 of pupae is prepared for each adult-holding cage which includes moistened sterile cotton as the emergence substrate inside a plastic container. Production of adults About 1500 male and female typically emerge from the 4 cm3 of pupae during the next 3 days.

In vitro method(s) Larvicidal assay28 Larval biting midges are handled much like mosquitoes for laboratory larvicidal screening tests. G

G

G

G

G

One cubic centimeter of acetone solution of each of a number of insecticides in serial dilution is pipetted into 190 cm3 of distilled water in 250 cm3 flasks with 1 cm3 acetone serving as a check. After 30 min, 10 cm3 of distilled water containing 25 fourth instars is added to each beaker. The beakers are held at B25 C
30 C for 24 h after which time the mortality is checked. A minimum of three tests should be conducted for each of three to four concentrations with mortalities between 10% and 90%. Slopes, LC50s and LC90s are calculated.

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Larvae are considered dead if they are incapable of swimming normally or do not respond to when probed with a needle. Larvae that pupate during the test are discarded. If more than 10% of the control larvae pupate, or the control mortality is more than 20%, the test should be discarded.

Adult assays28 Biting midges are attracted to light and frequently enter houses. They are small enough to pass through window screens. Spraying window screens with an insecticide can prevent annoyance by killing the midges that alight on the screen in the process of entering the room. The toxicity of insecticides to adult biting midges can be measured using standard WHO susceptibility kits in which the standard screen through which the midges can pass is replaced with a fine mesh screen or nylon stocking. Formulations can also be tested in small chambers with one end of the chamber covered by a transparent plastic and the other end opaque. A screen is inserted between the light and dark ends of the container. The adult midges are introduced into the container at the dark end. They are attracted to light and quickly alight on or pass through the treated screen at which time they are exposed to the insecticide. The duration of the effectiveness of such treatments can be measured by preparing a large number of screens in the same way and exposing them to weathering. Evaluation of effectiveness is measured by determining the times from passing through the screen (immediately after they have been introduced) until mortality. Snyder et al.29 reported on the use of insecticidal sugar baits (ISBs) for adult midges (C. sonorensis). Assays were conducted using disposable paper cups (0.28 L) covered by a piece of nylon organdy screening fastened by a lid. Midges were immobilized with CO2 and were subsequently allowed to recover from immobilization at least 1 h before trial initiation. Approximately, equal numbers of midges were sorted into each cup and provided with either a control noninsecticidal sugar solution, insecticidal sugar solution, or both. The treatment was presented by inserting a 9-dram lip vial filled with the solution into the side of the disposable paper cup. A 3-cm cotton wick inserted into the vial moved the insecticidal or control noninsecticidal solution out of the vial by capillary action. Insecticides were mixed in a sucrose solution of 10% sugar (weight to volume) using deionized water and diluted to obtain proper concentrations of commercial formulation. Both treated and untreated sugar solutions were left within the container for 24 h because individual midges fed at different times. Mortality rates were determined for various concentrations of commercial insecticides (0.01%, 0.05%, 0.1%, 1%, 2%, and 3%) and observed at 1, 4, 10, and 24 h postexposure to the ISB. In one assay, laboratory-reared midges

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were fed sugar ad libitum and then exposed to insecticide-treated sugar solutions to measure mortality. In a second assay where competitive feeding was assessed, midges were provided with a sugar control solution (10% sucrose) in one vial, and a sugar and insecticide solution in another. Biting midges were not deterred from feeding on the 1% ISB solutions despite the presence of an insecticide-free alternative source of sugar. Willingness to feed on a treated sugar solution source in the presence of a nontreated source demonstrated that the insecticidal bait had no repellency and may indicate how effective it would be in the field when competing with natural sources.

In vivo method(s) Laboratory efficacy test of cutaneous repellent product on human host The protocol described earlier for mosquitoes can be also used for midges. Laboratory artificial challenge on animal host in cages The protocol described earlier for mosquitoes can be also used for midges.

Psychodidae Phlebotomus Loew, 1845—Old World sand flies; Lutzomyia Franc¸a, 1924—New World sand flies Phlebotomine sand flies [genus Phlebotomus (Old World) and Lutzomyia (New World)] are of considerable public-health importance because of their ability to transmit several viral (arboviruses causing sand fly fever, meningitis, and vesicular stomatitis), bacterial (bartonellosis), and protozoal diseasecausing organisms (cutaneous, visceral, or mucocutaneous leishmaniasis) of humans and animals. Confusion with other types of biting flies is often caused because the common name “sand fly” is also used for other biting flies of the genera Ceratopogon and Culicoides. There are about 700 species of phlebotomine sand flies of which about 70 are considered to transmit disease agents to people and animals.

Biology and life cycle Sand flies go through complete metamorphosis, progressing from egg, to larva, pupa, and adult in B5
11 weeks (B36
74 days), depending upon the temperature and larval food quality. The eggs are elongated oval, dark brown in color with polygonal sculpturing over the chorion. The eggs are B0.31 mm long and 0.10 mm wide. The average duration of the egg stage ranges between 6 and 13 days. The first instars are pale cream in color with a body length measuring 0.40
0.68 mm. While the color remains similar to the first instar, the second and third instars are larger in size and the fourth instar measures

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2.2
2.5 mm. Setae and strong mandibles are present. The larvae feed in organic debris. The average duration of the four instars ranges from 21 to 51 days. Pupae are obtect, measuring B2.6 mm in length and look much like a butterfly chrysalis. They are glued to the substrate by the fourth instar exuvia. The pupae generally require B10 days (range 7
32 days) to develop. Adults are small, silvery-brownish, long-legged flies with narrow bodies. The dense-hairy wings are ,3 mm long and are held in a vertical “V” over the body. They are nocturnally active insects with weak, direct fight capability, typically not dispersing .0.5 km, unless passively transported by wind. Adults shelter during the day in dark, humid places, such as tree holes, animal burrows, or under rocks. Sexual dimorphism is marked between the male and female flies. Males have conspicuous external terminalia with a relatively small and slender abdomen compared to the female. Mandibles are absent in the males. The females require vertebrate blood for maturation of the follicles. The first oviposition begins between 4 and 8 days after blood ingestion. The males and females feed on nectar and other plant juices, but males do not take a blood meal. Eggs are laid (dispersed singly) in small batches in terrestrial humid microhabitats rich in organic matter that provides food for the larvae.

Rearing method(s) Of the known sand fly species, only a few of them have been successfully reared continuously in large numbers to provide sufficient material for experimental work. The first successful colonization of phlebotomine sand flies was by Grassi in 1907.30 Since then, a variety of laboratory rearing methods have been described.31
33 Nine species of phlebotomine sand flies in the genera Lutzomyia, Phlebotomus, and Sergentomyia have been reared and maintained in closed laboratory colonies for six or more generations.34 A 2011 publication by Volf and Volfova35 summarized their experiences with the establishment and maintenance of sand fly colonies and their use in infective experiments: techniques for the collection and handling of wildcaught females, rearing larvae and adults, and experimental infections of sand flies by Leishmania using membrane feeding. Collection in the field Sand flies of both sexes can be collected by several methods, either while resting during the day or foregoing at night. Sampling from resting sites, such as walls of houses, animal dwellings, caves, and tree holes, can be done by several forms of mouth aspirators; similarly, active catches can be done from bait animals or on human bait. The dimensions of each component of an aspirator can be varied according to the preference of the collector. However, since sand flies are fragile, the body of the aspirator should be

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wider than the opening. On the other side, aspirators are closed with a piece of fine mesh gauze or sinter filter. Light traps (e.g., CDC light trap) are used extensively in field studies of sand flies and can be left overnight to collect sand flies. The distance at which these traps are attractive to sand flies is not great. The traps should be collected early in the morning and the sand flies are removed by aspiration or directly transferred in a cage as soon as possible before they suffer from heat and desiccation. The transport to the laboratory of alive specimens should take into account the high humidity requirements of the sand flies in order to maintain them alive. Initiation of the colonies from field collected females Initiation of new colonies is a more difficult step than the routine maintenance of colonies already established in the laboratory for many generations. Refer to Volf and Volfova35 for the time-consuming and laborious details in the establishing of the first laboratory generation. Maintenance and rearing sand fly colonies Colonies are maintained either in a room with controlled temperature or in an incubator at 25 C
28 C. Photoperiod is probably not a crucial factor but 14L:10D or 16L:8D is used in many laboratories. High humidity (70%
90%) is ensured by wrapping cages in plastic bags or large plastic containers with a wet tissue or cotton wool inside, the surrounding humidity in the insectary or incubator (60%
70%) can be ensured by commercially available humidifiers. Different types of plastic containers (pots) with a large hole cut in the bottom and the walls roughened with sandpaper were used for larval rearing. The bottom is filled with a 1 cm thick layer of white plaster of Paris, and when set, a second very thin layer is plastered on the sides of the container. Plaster ensures the humidity in the container and provides a resting surface without water condensation. The container is closed with a fine gauze and snap-cap or screw-cap lid (dependent upon type of container used), the center of which is cut out. The gauze should be fine enough to prevent the escape of the larvae. Females are introduced through a small hole in the gauze using an aspirator (or tweezers, if anesthetized) and the hole is then plugged with a cotton wool pad. A small container (diameter 6 cm) is used for up to 20
40 gravid females, and a large container (diameter 14 cm) is used for 100
150 gravid females. Some colonies, like Lutzomyia longipalpis, Lutz & Neiva, 1912 develop well in “crowded” containers while others, such as Phlebotomus arabicus, Theodor, 1953 prefer less-crowded conditions. At 25 C
26 C, eggs are usually laid 6
10 days postblood meal. The length of the gonotrophic cycle is affected mainly by the speed of blood

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digestion, which varies within species and also depends on external factors, such as temperature and blood meal source. Larvae hatch from eggs usually after another 6
10 days. The day before hatching, the egg tooth and caudal setae are visible in the eggs as a dark spot and a dark line, respectively. Just before hatching is expected, a very small amount of food is sprinkled in several spots near the eggs. Larval food is a composted mixture of rabbit feces and rabbit pellets. Both feces and pellets are ground, but the feces must be first air-dried. (The resulting powder can be stored at 220 C for several months before composting.) A mixture of equal parts of ground feces and fecal pellets is spread in a thin layer (,0.5 cm) inside plastic trays and saturated with distilled water and is composted at RT, under aerobic conditions, for 3
4 weeks. Fermentation is controlled once a week and CO2 or other gas metabolites are released during the inspection of the fermentation process. The food is then dried in open air, scraped from the trays and ground in a mixer. It should be free of fungal spores but full of mycelia and products of cellulolytic enzymes from mold. Food can be stored at 220 C for several months or at 4 C for several weeks. The quality of larval food is a critical factor during the early larval stages. For the first instars, the food must be a very fine dust. This could be ensured by putting fine gauze over the mouth of a vial with food. Containers with larvae are maintained in plastic boxes with a bottom filled with about 1 cm of fine sand dampened with distilled water. For maintenance of colonies, sand is better than filter paper because of its higher water capacity. Sand, moreover, could be easily recycled as it is washed and sterilized before use. A weekly shaking of the sand layer prevents fungal growth on the bottom of the container. The containers are checked at least three times weekly and the food is replenished according to the number of larvae and their size. Excess food and humidity lead to fungal growth, its shortage to cannibalism and unequal development. The amount of food given to larvae and the moisture maintained are very important details during larval rearing. Before pupation, fourth instars empty their guts, making them more opaque. By this time, they do not need any additional food. The larval period (four instars) usually lasts for B3
4 weeks. The pupal period lasts for B7
10 days, with the pupal age distinguished by eye coloration. Dark eyes and wings are visible in pupae ready to emerge. In some colonies, especially those from temperate areas, fourth instars may diapause for unknown reasons, even in the presence of fixed artificial conditions of photoperiod and temperature. At least three times a week, emerged adults are released from the containers into nylon cloth cages supported with a steel frame. For the maintenance of colonies, several cage sizes (20 3 20, 40 3 40, or 50 3 50 cm) are used. Suspended cages inserted in a rigid frame prevent damage of adults due to crawling into splits and are removable for washing. Both sexes feed on sugar solutions or honey may be used as well. A 30%
50% sugar

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solution in distilled water can be sterilized in a microwave oven and stored at 4 C for several weeks or prepared weekly in order to prevent mold or bacterial growth. A small piece of cotton wool soaked with a sugar solution is offered in a Petri dish and changed at least three times a week. Honey solution is changed daily as it generates more fungal growth. Females are offered a blood meal on an anaesthetized animal once or twice a week. For most colonies, mice or hamsters are used, but feeding preferences of various species differ. Fed females are left undisturbed in a large cage for 24 h and then transferred into a small cage (20 3 20 cm) for defecation. When they are ready to lay eggs (for most species 5
6 days postblood feeding), they are transferred to moist oviposition containers using an aspirator. This two-step procedure prevents the early contamination of rearing containers by fungi. Selecting blood-fed females is laborious but it is necessary in colonies where only a small proportion of females take a blood meal. For larger colonies with high feeding rate, a one-step procedure is frequently used. Females are left to defecate in the first cage and then all females, together with males, are transferred directly to the rearing containers 24 h after blood meal. Cotton wool soaked with sucrose solution is placed on top of the nylon screen of the rearing container and changed at least every second day. Within the week of transfer, most females lay eggs and die. The dead females are removed with tweezers; the few survivors can be placed back into the big cage if necessary. Usually, the most critical generations are the third to fifth when the colony is adapting to laboratory conditions. By this time, even if females readily took blood and larvae are not diapausing, the generation time is 10
20 days longer than in an established colony. Colonies are in danger of infestation by mites, ascogregarines, pathogenic bacteria, and fungi. Care should be taken to avoid cross-contamination with these pathogens and cleanliness of the cages and containers is a prerequisite for successful rearing. An alternative method for the mass rearing of Lutzomyia cruzi and the establishment of a colony in a laboratory setting was reported by Oliveira et al.36

In vitro method(s) Phlebotomus argentipes Annandale & Brunetti, 1908, the established vector of kala-azar, is presently being controlled by indoor residual spray of DDT in kala-azar endemic areas in India. The search for nonhazardous and nontoxic biodegradable active molecules from botanicals was reported Dinesh et al.37 A bioassay was conducted with larvae and adult P. argentipes with different plant extracts collected in distilled water, hexane, ethyl acetate,

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acetone, and methanol. The larvicidal and adult bioassays reported by these authors may be useful for the determination of efficacy of other natural or chemical entities. Adult bioassay. Laboratory-reared 3-day-old female P. argentipes were exposed to plant extracts soaked in Whatman No. 1 filter paper along with deltamethrin at a concentration of 20 mg/cm2 as a positive control and distilled water and the respective solvent as negative controls. Crude extracts (50 μL) of plant samples were blotted on a 10 cm2 area of filter paper, dried at 40 C, and placed in a 50 mL centrifuge tube. Ten sand flies were exposed in the bioassay tube for 1 h and kept for 24 h in recovery tubes with 10% glucose solution soaked in a cotton ball along with control, positive control or negative control. All bioassay experiments were conducted in five replicates at 25 C 6 2 C and 72%
80% RH. The observed percentage of mortality was corrected using Abbott’s formula. Larvicidal bioassay. The powdery form of an extract was mixed with the larval food in different ratios of 1:1, 1:3, and 1:7. Rearing containers (pots) were prepared with a plastic tray (45 3 30 cm) having cubic wells of 3 cm2 (13 3 8 wells) with a thin layer of plaster of Paris at the bottom. Ten fourth instars were exposed in different rearing containers and were fed food with the plant extract at different concentrations with the negative and positive controls. Larval mortality was examined under a microscope and mortality from 0% to 100% was calculated using Abbott’s formula.6 Adulticidal bioassay. The toxicity of insecticides to adult sand flies can be measured using the CDC bottle bioassay.38 This assay is an inexpensive and portable alternative to the WHO exposure kit bioassay, especially in regions where there is little money to implement the WHO bioassay. (The WHO bioassay is a standardized protocol that consists of an exposure kit containing tubes lined with filter papers that are impregnated with a specific concentration of an insecticide and despite its accepted use worldwide, it is expensive and the filter papers are not available for some insecticides and/or concentrations.) In addition, the CDC bottle assay requires fewer test insects than the WHO bioassay. The protocol consists of coating the interior of a glass bottle with an insecticide that has been diluted in a solvent. The solvent is then allowed to evaporate, leaving the insecticide coated to the glass surface. Once the bottles are treated, insects are introduced into the bottles and exposed to the insecticide for a specified amount of time. Insect mortality can be scored at distinct time intervals during the exposure test (e.g., every 15 min for 1 h), and percent mortality at each time interval is plotted. The CDC bottle bioassay can also be used as an end-point assay where mortality is only measured at the end of the exposure test. G

Preparation of exposure bottles. On the day prior to exposing the sand flies, 1892.5-cm3 glass bottles or 1000-cm3 glass bottles were prepared

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by coating them with insecticide. For both bottle sizes, the concentration of insecticide in each bottle was determined to be X μg/bottle. For a 250-cm3 bottle, 1 cm3 of insecticide at 10 μg insecticide/cm3 acetone gives a concentration of 10 μg/250 cm3 bottle. To maintain an equivalence of 10 μg insecticide/250 cm3 bottle to compensate for the larger bottle sizes, 4 cm3 of 10 μg insecticide/cm3 acetone is needed to coat the interior of the 1000-cm3 bottle, and 7.57 cm3 of 10 μg insecticide/cm3 acetone is needed to coat the interior of the 1892.5-cm3 bottle. The bottles were coated with insecticide by swirling the acetone: insecticide solution on the bottom, on the sides, and on the lid. The bottle was then placed on a mechanical bottle roller under a chemical hood for 30 min to dry. During this time, the lids were slowly loosened to allow the acetone to evaporate. After 30 min, the caps were removed, and the bottles were rolled until all of the acetone had evaporated. The bottles were then left open to dry overnight. For each test replicate, one bottle serving as a control was coated with either 7.57 or 4.0 cm3 acetone, depending on its volume. All bottles were reused throughout the duration of the experiment. Insecticide exposure tests. Approximately 12 h after the bottles were prepared with insecticide, adult sand flies at least 2 days postecolsion were aspirated from the main colony and gently blown into each bottle: 40
50 flies into each 1892.5 cm3 bottle and 20
30 flies into each 1000 cm3 bottle. Approximately equal numbers of unfed female and male flies were used for each replicate. At least three replicates were completed for each concentration of every insecticide.

A range of exposure times was used (30
120 min), depending on unexpected and actual sand fly survival rates. The sand flies were captured after insecticide exposure via mechanical aspiration, released into cardboard containers (B473 cm3) with a fine mesh screen top, and kept under the same temperature, light, and humidity environment as the main untreated colonies. A cotton ball saturated with 30% sugar-water was placed in the top of each container as an energy/water source. Sand flies were held in these containers for 24 h prior to mortality being recorded. Mortality was scored as a complete cessation of movement. A 24 h holding period was used because in some preliminary experiments, many of the sand flies that appeared physically affected and would have been scored as dead at the end of a 30, 60, or 120 min exposure recovered after this 24 h period. LCs (LC50, LC90, and LC95) were calculated for each insecticide. If mortality in the control group ranged between 5% and 20%, mortalities in the experimental bottles of that test group were corrected using Abbott’s formula,6 but not corrected for experimental mortalities ,5%. If control mortalities exceeded 20%, the entire test replicate was not used.

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The CDC bottle bioassay can also be used to assess insecticide resistance.39 Ovicidal, larvicidal, adulticidal combined bioassay The objective of an in vitro bioassay40 was to evaluate the effect of neem oil on eggs, larvae, and adults of the vector. The insects were captured in the field and kept in the laboratory at B27 C and 80% RH. Five treatments with different concentrations were performed using two negative controls (distilled water and Tween 80) and a positive control. The eggs were sprayed with the oil at different concentrations and the number of hatched larvae evaluated at 10 days. Mortality of larvae was observed to pupation and adult mortality was observed after 24, 48, and 72 h. Statistical analysis was performed by Tukey test at 5% probability. When using larvae of L. longipalpis, 40 eggs were transferred to plastic pots internally coated with sterile plaster containing a thin layer of food substrate for hatching. Six days after hatching, the larvae were counted and sprayed with oil solutions in various concentrations, dependent upon the genus source of the oil. In in vitro tests against adult L. longipalpis, for each oil concentration (1 mL), cypermethrin or Tween 80 was applied to the inner surface and bottom of each pot using a pipette. Thirty adults (15 of each sex) were placed inside of the pots after application of the oils, and various oil concentrations were used. The parameters observed were insect mortality after 24, 48 and 72 h mortality rate differences between male and female insects and the number of eggs obtained from females subjected to the oils. Repellency Spatial repellency test of sand flies can be conducted by the method described by Rowton et al.41 using the redesigned Grieco “high-throughput” modular assay chamber. General insect repellency testing methods and methods for field testing topical applications of compounds as repellents may be applicable.42
44

In vivo method(s) Laboratory efficacy test of cutaneous repellent product on human host The protocol described earlier for mosquitoes can be also used for sand flies. Laboratory artificial challenge on animal host in cages The protocol described earlier for mosquitoes can be also used for sand flies. Experimental transmission studies Dogs as reservoir hosts of Leishmania species and experimental transmission studies have been conducted. The capacity of infected dogs to transmit the

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Leishmania braziliensis to sand fly vectors (Lutzomyia trapidoi, Lutzomyia gomezi, L. longipalpis, Lutzomyia youngi) was tested by allowing the flies to feed on the lesion borders.45 Parasites were isolated by culture in Senekjies culture medium at 25 C and identified using monoclonal antibodies. Dogs were anesthetized with xylazine and the lesion borders exposed for 30 min to the bites of sand flies. Between 30 and 40 flies took a blood meal on each dog. No infections were detected upon dissection of engorged flies. A single peri- and sublesional injection of 1
2 mL of pentavalent antimony in the dog with ear lesions resulted in clinical cure 6 weeks PT. Although dogs are susceptible to L. braziliensis, their reservoir competence could be low. However, there is sufficient epidemiological and experimental evidence supporting dogs as the main reservoir hosts of Leishmania infantum for human infection. Therefore it is plausible to conclude that the domestic dog plays an important role in the epidemiology of zoonotic visceral leishmaniasis.46

References 1. Gonzales KK, Hansen IA. Artificial diets for mosquitoes. Int J Environ Res Pub Health 2016;13(12):1267. 2. Clements AN. Biology of mosquitoes: development. Nutrition and reproduction. Netherlands: Springer; 1992. p. 509. 3. Cosgrove B, Wood RJ, Petric D, Evans DT, Abbott RHR. A convenient mosquito membrane feeding system. J Am Mosq Control Assoc 1994;10(3):434
6. 4. Bunner BL, Scott RL, Dobsin SE, Anderson LM, Boobar LR. Comparison of artificial membrane with live host blood feeding of Aedes aegypti (L) (Diptera: Culicidae). J Entomol Sci 1989;24:198
203. 5. WHO guidelines for laboratory and field testing of mosquito larvicides, 2005. 6. Abbott WS. A method for computing the effectiveness of an insecticide. J Econ Entomol 1925;18(2):265
7. 7. Pridgeon JW, Becnel JJ, Clark GG, Linthicum KJ. A high-throughput screening method to identify potential pesticides for mosquito control. J Med Entomol 2009;46(2):335
41. 8. Derua YA, Malongo BB, Simonsen PE. Effect of ivermectin on the larvae of Anopheles gambiae and Culex quinquefasciatus. Parasit Vectors 2016;9:131
7. 9. Zairi J, Lee YW. Laboratory and field evaluation of household insecticide products and public health insecticides against vector mosquitoes and house flies (Diptera: Culicidae: Muscidae). Proceedings of the Fifth International Conference on Urban Pests; 2005;477
82. 10. Huang TH, Tien NY, Luo YP. An in vitro bioassay for the quantitative evaluation of mosquito repellents against Stegomyia aegypti (5Aedes aegypti) mosquitoes using a novel cocktail meal. Med Vet Entomol 2015;29(3):238
44. 11. Klun JA, Kramer M, Debboun M. A new in vitro bioassay system for discovery of novel human-use mosquito repellents. J Am Mosq Control Assoc 2005;21(1):64
70. 12. Grieco JP, Achee NL, Sardelis MR, Chauhan KR, Roberts DR. A novel high-throughput screening system to evaluate the behavioral response of adult mosquitoes to chemicals. J Am Mosq Control Assoc 2005;21(4):404
11.

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13. Darriet F, Chandre F. Efficacy of six neonicitinoid insecticides alone and in combination with deltamethrin and piperonyl butoxide against pyrethroid-resistant Aedes aegypti and Anopheles gambiae (Diptera: Culicidae). Pest Manag Sci 2013;65:905
10. 14. Khalid K, Qualls WA, Revay EE, Allan SA, Arheart KL, Krachenko VD, et al. Attractive toxic sugar baits: control of mosquitoes with the low-risk active ingredient dinotefuran and potential impacts on nontarget organisms in Morocco. Environ Entomol 2013;42 (5):1040
5. 15. WHO/HTM/NTD/WHOPES/2009.4 16. Rutledge LC. Mathematical models of the effectiveness and persistence of mosquito repellents. J Am Mosq Control Assoc 1985;1:56
62. 17. Reifenrath WG, Rutledge LC. Evaluation of mosquito repellent formulations. J Pharm Sci 1983;72:169
73. 18. Bernardo MJ, Cupp EW. Rearing black flies (Diptera: Simuuliidae) in the laboratory: mass-scale in vitro membrane feeding and its application to collection of saliva and to parasitological and repellent studies. J Med Entomol 1986;23(6):666
79. 19. Gray EW, Noblet R. Black fly rearing and use in laboratory bioassays. Rearing animal and plant pathogen vectors. Boca Raton, FL: CRC Press; 2014. 20. Edman JD, Simmons KR. Rearing and colonization of black flies (Diptera: Simulidae). J Med Entomol 1985;22(1):1
17. 21. Carlos JPC, Araujo-Coutinho, Figueiro R, Viviani AP, Nascimento ES, Cavados CFG. A bioassay method for black flies (Diptera: Simuliidae) using larvicides. Neotrop Entomol 2005;34(3):511
13. 22. Overmyer JP, Mason BN, Armbrust KL. Acute toxicity of imidacloprid and fipronil to a nontarget aquatic insect, Simulium vittatum, Zetterstedt cytospecies IS-7. Bull Environ Contam Toxicol 2005;74:872
9. 23. Overmyer JP, Rouse DR, Avants JK, Garrison AW, Delorenzo ME, Chung KW, et al. Toxicity of fipronil and its enantiomers to marine and freshwater non-targets. J Environ Sci Health 2005;Part B42:471
80. 24. Rutledge LC, Ward RA, Gould DJ. Studies on the feeding response of mosquitoes to nutritive solutions in a new membrane feeder. Mosq News 1964;24:407
19. 25. Hunt GJ. A procedural manual for the large-scale rearing of the biting midge, Culicoides variipennis (Diptera: Ceratopogonidae). Washington, DC: ARS-121. USDA, Agricultural Research Service; 1994. 26. Hunt GJ, Mullens BA, Tabachnick WJ. Chapter 2: Colonization and maintenance of species of Culicoides. In: Maramorosch K, Mahmood F, editors. Maintenance of human, animal and plant pathogen vectors. New Delhi: Published by Mohan Primlani for Oxford & IBH Publishing Co. Pvt. Ltd.; 1999. 27. Hunt GJ, McKinnon CN. Evaluation of membranes for feeding Culicoides variipennis (Diptera: Ceratopogonidae) with an improved artificial blood-feeding apparatus. J Med Entomol 1990;27(5):934
7. 28. Carmichael G, Alvarez CG, Mulla MS, Mount G, Jamnback H. Analysis of specialized pesticide problems invertebrate control agent-efficacy test methods. Mosquitoes, black flies, midges and sand flies. 1977;49
52. 29. Snyder D, Cernicchiaro N, Allan SA, Cohnstaedt LW. Insecticidal sugar baits for adult biting midges. Med Vet Entomol 2016;30:209
17. 30. Grassi B. Richerche sui flebotomi. Mem Soc Ital Sci Ser 1907;14:353
94.

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31. Killick-Kendrick R, Leaney AJ, Ready PD. The establishment, maintenance and productivity of a laboratory colony of Lutzomyia longipalpis (Diptera: Psychodidae). J Med Entomol 1977;13:429
40. 32. Modi GB, Tesh RB. A simple technique for mass rearing Lutzomyia longipalpis and Phlebotomus papatasi (Diptera: Psychodidae) in the laboratory. J Med Entomol 1983;20 (5):568
9. 33. Endris RC, Young DG, Butler JF. The laboratory biology of the san fly Lutzomyia anthophora (Diptera: Psychodidae). J Med Entomol 1984;21(6):656
64. 34. Endris RG, Perkins PV, Young DG, Johnson RN. Techniques for laboratory rearing sand flies (Diptera: Psychodidae). Mosq News 1982;42(3):400
7. 35. Volf P, Volfova V. Establishment and maintenance of sand fly colonies. J Vector Ecol 2011;36(1):S1
9. 36. Oliveira EF, Fernandes WS, Oshiro ET, Oliveira AG, Galati EAB. Alternative method for the mass rearing of Lutzomyia (Lutzomyia) cruzi (Diptera: Psychodidae) in a laboratory setting. J Med Entomol 2015;52(5):925
31. 37. Dinesh DS, Kumari S, Pandit V, Kumar J, Kumari N, Kumar P, et al. Insecticidal effect of plant extracts on Phlebotomus argentipes (Diptera: Psychodidae) in Bihar, India. Indian J Med Res 2015;142(1):S95
100. 38. Denlinger DS, Lozano-Fuentes S, Lawyer PG, Black IV WC, Bernhardt SA. Assessing insecticide susceptibility of Laboratory Lutzomyia longipalpis and Phlebotomus papatasi Sand Flies (Diptera: Psychodidae: Phlebotominae). J Med Entomol 2015;52(5):1003
12. 39. Denlinger DS, Creswell JA, Anderson JL, Reese CK, Bernhardt SA. Diagnostic doses and times for Phlebotomus papatasi and Lutzomyia longipalpis sand flies (Diptera: Psychodidae: Phlebotominae) using the CDC bottle assay to assess insecticide resistance. Parasit Vectors 2016;9:212
22. 40. Maciel MV, Morais SM, Bevilaqua CM, Silva RA, Barros RS, Sousa RN, et al. In vitro insecticidal activity of seed neem oil on Lutzomyia longipalpis (Diptera: Psychodidae). Rev Bras Parasitol Vet 2010;19:7
11 [In Portuguese]. 41. Rowton ED, Grieco JP, Coleman RE. High throughput testing for toxic and behavior modifying chemicals against Phlebotomine sand flies. In: DoD tri-service pest management workshop; 2007. 42. World Health Organization. Guidelines for efficacy testing of spatial repellents Geneva. 2013. ISBN 978 92 4 150502 4. 43. Rutledge LC, Mehr ZA, Debboun M. Chapter 8: Testing methods for insect repellents. In: Debboun M, Frances SP, Strickman D, editors. Insect repellent handbook. 2nd ed. Boca Raton, FL: CRC Press; 2014. p. 159
78. 44. American Society for Testing and Materials (ASTM) E939-94. Standard test methods for field testing topical applications of compounds as repellents for medically important and pest arthropods (including insects, ticks and mites): Mosquitoes. 2006. 45. Travi BL, Tabres CJ, Cadena H. Leishmania (Viannia) braziliensis infection in two Colombian dogs: a note on infectivity for sand flies and response to treatment. Biomedica 2006;26(1):249
53. 46. Dantes-Torres F. The role of dogs as reservoirs of Leishmania parasites, with emphasis on Leishmania (Leishmania) infantum and Leishmania (Viannia) braziliensis. Vet Parasitol 2007;149:139
46.

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Chapter 3b

Arthropoda, Diptera, Brachycera Larry R. Cruthers, MS, PhD1 and Marco Pombi, PhD2 1

LCruthers Consulting, Chesapeake, VA, United States, 2Dipartimento di Sanita` Pubblica e Malattie Infettive, Sapienza University of Rome, Rome, Italy

Arthropoda Diptera Brachycera

Tabanidae Tabanus Linnaeus, 1758—horse flies; Chrysops Meigen, 1803—deer flies Often considered pests for the bites that many inflict, horse and deer flies are among the world’s largest flies. They are known to be extremely noisy during flight. Tabanids are very good vectors of the equine infectious anemia virus, as well as some Trypanosoma species. Species in the genus Chrysops are biological vectors of Loa loa, transmitting this parasitic filarial worm among humans. They have also been known to transmit anthrax in cattle and sheep, and tularemia between rabbits and humans. Tabanids occur worldwide, being absent only at extreme northern and southern latitudes. They are often encountered as bee-like insects that follow people outdoors and bombard them, acting as a nuisance factor. They tend to favor the head and upper body, where they usually alight completely unnoticed by the victim until the painful bite gives them away.

Biology and life cycle As with all other true flies, tabanids go through complete metamorphosis (egg, larvae, pupae, and adult). The eggs are usually laid in large, layered clusters of 25
1000 on vegetation or other objects overlying water or moist soil. The incubation period is greatly influenced by weather conditions, but during the warmer months, the usual range is from 5 to 7 days. Depending on the species, the larvae may be aquatic, semiaquatic, or terrestrial; they hatch from the eggs and drop to the water or soil below, where they become voracious predators of other invertebrates or small vertebrates. The head contains two sharp, slender mandibles with a hollow canal for transmitting venom into their prey. The larva undergoes

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several molts as it grows, and depending on the species, the larval stage may last several months or as long as 2
3 years. Once the larva is fully developed, it moves into drier soil, usually 25 mm or so below the surface to pupate. Depending on the species, the pupal stage lasts B5
21 days, and then the adult flies emerge from the soil. Mating occurs shortly after the adults emerge. Female horse and deer flies are bloodsucking insects whereas male flies feed on nectar and are of no consequence as animal or human pests. Horse flies range in size from B19 to 32 mm long and usually have clear or solidly colored wings and brightly colored eyes. Deer flies, which commonly bite humans, are smaller with dark bands across the wings and colored eyes similar to those of the horse flies. Female horse flies and deer flies are active during the day. These flies are apparently attracted to such things as movement, shiny surfaces, CO2, and warmth. Once on a host, they use their knife-like mouthparts to slice the skin and feed on the blood pool that is created (pool-feeders). It has been estimated1 that horse flies would consume 1 cm3 of blood for their meal, and 20
30 flies feeding for 6 h would consume 100 cm3. Bites can be very painful and there may be an allergic reaction to the salivary secretions released by the insects as they feed. Horse and deer flies are intermittent feeders because their painful bite generally elicits a response from the victim, so the fly is forced to move to another host. This intermittency increases the likelihood to be involved in the mechanical transmission of disease. Following the blood meal, oviposition commences.

Rearing method(s) The authors found no methods reported in the literature for the successful mass-rearing and maintenance of tabanids. One of the main reasons is the persistent difficulty of rearing larvae in the laboratory. These difficulties include (1) the larval stage development period that is very long, lasting from 1 to 3 years, (2) due to the fact that the larvae are cannibalistic, they must be reared individually, (3) they need living prey for food, and (4) they have to be kept free from contamination by various pathogens. Thompson et al.2 attempted to establish the economically important (vector of equine infectious anemia) and locally available tabanid, Hybomitra lasiophthalma Macquart, 1838 in their laboratory. Although no eggs were laid by any of the field-collected females exposed to blood in the laboratory, 44 egg masses were laid by trapped females that were partially engorged at the time of collection. Most of these egg masses failed to hatch, 51 larvae from a total of 4 lots did develop and reached maximum

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growth in B2 months after which time they entered diapause. Results showed that the larvae of H. lasiophthalma could be reared to maturity en masse without the effect of cannibalism becoming restrictive. However, for these insects to complete development, a way had to be developed to prevent diapause. The prospect of maintaining adults with the hope of mating was not encouraging. Middlekauff and Lane,3 for taxonomic purposes, identified techniques useful for extracting immature tabanids from soil or vegetation and for the rearing of larvae in glass beads, agar, sand, paper toweling, filter paper, and soil from the natural habitat, with the conclusion that for observing larval activities or determining the number of molts, the use of filter paper or paper toweling as a medium was recommended. No specific information was provided as to how the larvae were reared in these media. Ferreira and Rafael,4 in order to study the distribution and taxonomy of tabanids, reported on a novel rearing method of immature horse flies by using a substrate of bryophytes and sand. Details provided in this report were insufficient to reproduce their work. The work of Matsumura5 on the laboratory rearing of Tabanus nipponicus Murdoch & Takahasi, 1969 was more encouraging. Engorged females were collected on pasture after they had taken a blood meal. They were kept individually on cages 11 cm 3 8.5 cm and fed a 5% honey water solution. Each cage was provided with a small plant pierced into artificial moss (hydrophilic plastic) as an oviposition site. Egg masses laid on the underside of the leaves were placed individually on a vinyl-chloride net stretched over a shallow dish filled with water, thus ensuring that the newly hatched larvae would fall into the water. Larvae were placed in groups of 10 individuals on plastic, cylindrical tubes for the first 20 days after hatching, after which time these larvae were placed individually in plastic cups containing artificial moss and fed earthworms cut into pieces proportional to the larval body size. The plastic media was replaced when it became dirty or moldy. After an extended outdoor storage period, the rearing containers were checked daily for pupation and emergence of adults. All stages were reared in the natural photoperiodicity and at RT except for the storage outdoors in winter. Results of this work revealed that the engorged females were brought into the laboratory from pasture oviposited from 4 to 13 days after bloodfeeding. In the laboratory, the eggs were laid in mass and the larvae hatched simultaneously in each egg mass; the average hatchability was 65.2%. The larvae grew rapidly, molting 5
7 times. The last instar was attained in autumn and hibernated until the next spring. After hibernation, they did not molt again until pupation. During the group rearing period, the number of larvae decreased dramatically due to cannibalism, the survival rate of early

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instars was 20% after 3 weeks. Thereafter, the survivability increased with individual rearing. Of the 500 larvae that hatched, only 33 adults emerged (6.6% survival rate). In addition, Magnarelli and Anderson,6 studied the oviposition, fecundity, and fertility of the salt marsh deer fly Chrysops fuliginosus Weidemann, 1821. Virgin females, reared from larvae or pupae collected from a salt marsh and reared individually in tightly lidded plastic cream cups (volume 5 30 cm3) with 6 g of sieved soil medium (composed of fine-textured vegetative and mineral components), and maintained at 23 C 6 3 C, oviposited in the sides of cups and beneath the lids within 9 days of emergence. Egg counts ranged from 72 to 209. Also, 48 females captured in copula from the salt marsh were transferred to similar plastic cream cups and reared as the virgin laboratory-reared flies until egg deposition or death. The oviposition habits and fecundity of inseminated females were similar to those observed for virgin females in the laboratory. In conclusion, large numbers of fertile eggs and early instars can easily be obtained from inseminated field-caught females and can be used for rearing, taxonomic, or other purposes. Since virgin females that are reared in the laboratory readily oviposit, it may also be possible to obtain viable eggs and larvae after successful mating in the laboratory by forced copulation, thereby eliminating the need to capture copulating females from the field.

In vitro method(s) The authors found no in vitro screening methods for immature or adult horse or deer flies. This may be due to the fact that they have not been successfully reared en masse in the laboratory and are thus unavailable in significant numbers for testing. In vivo method(s) Efficacy test of cutaneous repellent product on naturally infested animal host A study performed on cows naturally infested with Haematobia irritans7 can be used as a reference study design for other animal and insect models, such as horse/deer flies. The animals were confirmed to be naturally infested based on counts at TD-2 and -1. The sample size of animal hosts required in the study was statistically justified considering both the efficacy and the safety aspects of the study. Animals included in the study did not share the same pastures with other animals. Animals and the test environment were not treated with an ectoparasitide or biocide, respectively, within a time frame that might impact the study outcome. Two

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groups of at least five animals placed in two different areas received an appropriate dosage of repellent solution distributed on the head, neck, and dorsal region of the body (in case of large animals as cows) or the whole body, if possible. All animals of the same group were placed in the same environment. All insects of the target species naturally present on the animals were counted just before (TD-2 and -1) and after treatment (1, 2, 3, 6, 9, 24, 48, and 72 h PT). Data were analyzed by ANOVA with post hoc analysis (Tukey’s test) to highlight differences in time points and test groups. Efficacy test of pour on insecticide products on naturally infested animal host The recommendations reported earlier on the animal and environment conditions must be followed also in this case. The evaluation of a topic product can be done as reported by Lopes et al.8 for calves naturally infested by H. irritans. This test can be easily adapted to other hosts according to the product of interest and the ectoparasite. An appropriate number of animals (e.g., two groups of at least five) were selected according to high and comparable infestation levels at TD-2 and confirmed at TD-1. Then, the animals were divided into two groups: one group was treated with the pour on test product, while the second group was treated with the solvent as a control. The two groups were physically separated in different paddocks to prevent any contact between insects infesting animals from different groups. Then, insects were counted on the entire body surface of each animal on TD 1, 3, 7, 14, 21, and 28 PT. Data were analyzed by ANOVA with post hoc analysis (Tukey’s test) to highlight differences in time points and test groups. Tabanids are considered to be one of the most challenging livestock flies to control, due to a number of factors, mainly associated with their life cycle. Female tabanids spend only 4 min feeding on the host in order to generate eggs, which develop into adults the following year. Direct observation of flight behavior is not possible and adult life history is often calculated (albeit insufficiently) on light trap catches. A more important factor is the tendency to generalize about a family of over 4000 species and at least 137 genera. Due to the complex life cycle which is partly independent of livestock, integrated control strategies are required to reduce the impact of these pests.9 1. Chemical control9: The potential for area-wide control of tabanid populations using insecticides has been studied. As with most other flies, control of the larval habitat is most effective if such control is possible. Studies in Africa and

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the United States demonstrated that larval tabanid populations and subsequent adult populations could be dramatically reduced by the widespread application of persistent insecticides on larval habitats. However, this area of research has received little attention due to the demonstrated harmful environmental impacts associated with such treatments. Areawide control of adult tabanids has met with less success, and this is often blamed on the mobility of adult tabanids. The effect on the tabanid burden of the use of insecticides and repellents applied to livestock has been studied to some degree. The majority of the products used for equine protection have historically contained synergized pyrethrins with some combination of known repellents. Due to limited residual activity and expense, the use of pyrethrins is normally confined to valuable stock under intensive management. Many of the newer formulations for equine control contain one of the several pyrethroids, such as permethrin, resmethrin, or cypermethrin. The pyrethroid fenvalerate applied using a high-pressure spray as a model for determining the effects of pyrethroids on tabanid feeding success has been studied. Feeding time was found to be significantly lower (35% reduction) on treated cows for the horse fly species observed and the amount of blood consumed was significantly reduced (by 30%). The efficacy of pyrethroid ear tags and sprays has been evaluated against tabanids under field conditions in Louisiana. The overall fly mortality rates were 3% in controls, 9% using permethrin tags, 15% using fenvalerate tags, 67% using fenvalerate spray at 0.015, 79% using fenvalerate spray at 0.02%, and 100% using L-cyhalothrin tags. 2. Cultural control9: Animal management can influence the incidence of tabanids on livestock. Few tabanid species will enter barns or other structures; the species of horse flies which do enter structures are usually active during crepuscular or nocturnal periods. Tabanid attack can be reduced even when cattle are stanchioned beneath roofs supported by posts and with open sides. Given assess to suitable structures, free-roaming livestock will seek shelter from tabanid attack. Fewer problems are encountered on pastures located well away from wooded areas. The use of vegetative barriers has been proposed to prevent movement of salt marsh tabanids (Tabanus nigrovittatus) into strategic areas. Water management is also a tool which can be used for managing tabanid larval habitats. Unfortunately, there is such a paucity of information on larval habitats in general that rational decisions cannot be made regarding the potential effects of draining.

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3. Mechanical control9: The salt marsh greenhead (T. nigrovittatus) is considered to be an important human pest along the Atlantic coast of the United States and the use of permanent traps (and possibly treated silhouette traps) for adult control has been advocated. However, long-term population suppression is not considered possible. T. nigrovittatus females are autogenous, and trapping does not prevent the first oviposition. 4. Biological control9: When considering biological control, it should be recognized that tabanids are no exception among insects; there are many natural infectious diseases, parasites, and predators at all life stages of tabanids. However, manipulation of natural mortality factors has rarely been explored for the suppression of tabanid populations.

References 1. Webb JL, Wells R.W. Bulletin no. 1218. Washington, DC: US Department of Agriculture; 1924. 36 pp. 2. Thompson PH, Hogan BF, Del Var Petersen H. Rearing of Texas Tabanidae (Diptera). III. Trapping, survivorship, and limited rearing of Hybomitra lasisphthalma (Macquart). Southwest Entomol 1980;5(3):191
5. 3. Middlekauff WW, Lane RS. Adult and immature Tabanidae (Diptera) of California. Bulletin of the California insect survey, vol. 22. University of California Press; 1980. p. 3
4. 4. Ferreira RLM, Rafael JA. Rearing immature horse flies (Diptera: Tabanidae) by using a substrate of bryophytes and sand. Neotrop Entomol 2006;35(1):141
4. 5. Matsumura T. Development of Tabanus nipponicus (Diptera, Tabanidae) confirmed by laboratory-rearing. Appl Entomol Zool 1995;30(1):57
65. 6. Magnarelli LA, Anderson JF. Oviposition, fecundity, and fertility of the Salt Marsh Deer Fly, Chrysops fuliginosus (Diptera: Tabanidae). J Med Entomol 1979;15(2):176
9. 7. Klauck V, Pazinato R, Stefani LM, Santos RC, et al. Insecticidal and repellent effects of tea tree and andiroba oils on flies associated with livestock. Med Vet Entomol 2014;28 (1):33
9. 8. Lopes WDZ, Chiummo RM, Vettorato LF, de Castro Rodriques D, Sonada RB. The effectiveness of a fixed-dose combination pour-on formulation of 1.25% fipronil and 2.5% fluazuron against economically important ectoparasites and associated pharmacokinetics in cattle. Parasitol Int 2017;66(5):627
34. 9. Foil LD, Hogsette JA. Biology and control of tabanids, stable flies and horn flies. Rev Sci Tech Off Int Epiz 1994;13(4):1125
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Chapter 3c

Arthropoda, Diptera, Cyclorrhapha Alan A. Marchiondo, MS, PhD1, Philip J. Scholl, PhD2 and Ronnie L. Byford, PhD3 1 Adobe Veterinary Parasitology Consulting LLC, Santa Fe, NM, United States, 2Retired Entomologist, Oxford, FL, United States, 3Center for Animal Health & Food Safety, New Mexico State University, Las Cruces, NM, United States

Arthropoda Diptera Cyclorrhapha—flies The Cyclorrhapha feed upon or are associated with livestock and domestic animals, and they tend to breed on decaying plant and animal tissues and manure. The slender larval instars of the families Muscidae, Sarcophagidae, and Calliphoridae are usually referred to as maggots, whereas the robust larvae of the family Oestridae are called bots or grubs.

Muscidae The family Muscidae contains a number prominent species used in screening insecticides and parasiticides: Musca domestica, the common housefly; Musca autumnalis, the face fly, Fannia canicularis, the lesser housefly; Glossina spp., tsetse fly; Stomoxys calcitrans, the stable fly; Haematobia irritans, the horn fly; and Melophagus ovinus, the sheep ked.

Musca domestica Linnaeus, 1758—housefly Biology and life cycle The housefly is a cosmopolitan species commonly associated with livestock facilities housing poultry, swine, sheep, horses, and cattle as well as human garbage. The housefly is a nuisance species and can constitute a public annoyance. Movement of Musca flies between feces and food makes them ideal transmitters of human and animal disease. Houseflies harbor up to 100 different pathogenic organisms and generally propagate disease by mechanically transmitting pathogens that cause cholera, typhoid, bacillary dysentery, bovine mastitis, and conjunctivitis (Moraxella bovis). These flies may also carry pathogens externally on their body and transmit anthrax and glanders in this way. Adult houseflies are medium-sized insects, ranging from 5 to 8 mm in length (females 6
8 mm and males 5
6 mm in length). They are nonmetallic in color with a black-and-gray-striped thorax, and the abdomen has a

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yellowish-brown background color with a black medium longitudinal stripe. The head of the adult flies has reddish eyes and sponging mouthparts. Solid foodstuff is first liquefied with saliva before being ingested. After an adult housefly emerges from its puparium (pupal case) with the aid of its ptilinum, it seeks a resting site to expand its wings and to allow the exoskeleton to harden. Feeding occurs within 24 h after emergence. Most female houseflies become receptive B36 h after emergence and mate once, storing sperm in their spermathecae. Female houseflies can produce B150 eggs at a time laying eggs singly in successive batches at 2
4 day intervals. The entire life cycle from egg to egg can occur in as few as 10
14 days under warm (40 C) and humid (90% RH) conditions. Eggs are oval, slightly curved, whitish in color, and measure B1 mm in length. Eggs are laid singly, often piled in small groups, in moist animal feces, manure piles, garbage, and other decaying organic matter. First instars emerge from eggs within 8
20 h under warm conditions. There are three larval instars (maggots) that develop in B3
8 days under optimum temperature of 30 C
37 C. Early instars measure 3
9 mm in length, creamy white in color, cylindrical in shape but tapered toward the head. Instars possess one pair of dark hooks on the head and the posterior spiracular openings are sinuous slits completely surrounded by an oval black border. The third instar (8
15 mm in length) pupates in relatively dry areas. Pupae are barrel shaped and red to brown in color. The pupal stage lasts 3
10 days depending on temperature and RH.

Rearing method(s) Housefly rearing SOP Medium preparation Two sticks of malt extract (B120 mL) are placed in a 1000 mL graduated beaker. Warm water is added to the 1000 mL mark and the contents stirred until the malt has been dissolved. Add one packet of dry yeast. Stir the contents for a few minutes to allow the yeast to begin bubbling to the surface. Pour the solution into a plastic rearing dish containing two scoops of CSMA fly rearing medium. Rinse the beaker with another 1000 mL of warm water and add this to the rearing dish. Hand mix, wearing plastic gloves, the medium until all the CSMA has been wetted. Cover the rearing dish with cheesecloth. Place the dish in a fume hood overnight to allow CO2 production to dissipate. Egg collection Fold two paper towels in half long ways and place in half Styrofoam cups (4 oz). A milk solution made from dry milk is prepared and poured into the half cups to thoroughly saturate the towels. Excess milk is drained off, so there is no standing milk. These egging cups are placed one per fly cage and left overnight. The next day the egging cup is removed

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from the cage. The towels and cup are rinsed with water through a No. 100mesh screen (0.149 mm) and the eggs (1
2 mL) are placed in the rearing medium. The eggs are allowed to develop at 18 C for 8
11 days. Pupae collection When pupae have been developed on the surface of the rearing medium, add warm water to the dish and stir the medium well. Skim the surface of the dish with a beaker to collect all floating material (pupae and chaffe). Pour the skimmed material through a no. 16-mesh sieve (1.19 mm) and rinse the contents of the sieve with warm water to rinse away as much chaffe as possible. Repeat the skimming step one or two more times to collect as many pupae as possible. Place the collected pupae in a dryer until they are dry. Adult flies Place 22
25 g of pupae in a half cup and place it in a fly cage. Also place water in a half cup and a half cup of fly food (two parts dried milk/one part sugar) into the cage. Check water and fly food twice a week. Adult flies will emerge from the pupae, feed, and mate. Colony care protocol for the common housefly Adults: Numerous methods have been published on the rearing of housefly colonies in the laboratory.1
6 Basically, adult flies are housed in 12 3 12 3 12 in. screen cages fitted with stockinette sleeves (Bioquip No. 1450). Flies are provided with food in the form of a mixture of sucrose (80 cm3) and nonfat dry milk (400 mL), mixed together in equal volumes and placed in plastic weigh boats or 236 mL cups. Addition of 3.5 mL of 37% formalin may be added as a preservative. A strip of 3 3 10 in. cellucotton or paper towel (3- or 4-panel accordion fold) is placed in cups. The strip allows flies a resting place during feeding and provides a good oviposition site. Water sources consist of 150 mL beakers of water inverted on Petriplate halves covered with filter paper disks. Cotton balls placed on the rims of the beakers facilitate water flow. In cages with low numbers of flies, one food and one water source is used. In cages with higher numbers, two of each may be needed. Flies will die within a few days without water. Adult flies survive in large numbers for up to 1 month. After that time, they begin to die off. When numbers in a given cage are too low to justify continuing that cage, the water is removed and the flies are allowed to die. They are then discarded, and the cage cleaned. Eggs: Adult flies prefer “less wet” areas for oviposition and also narrow slits where they can probe with their ovipositor. They will not produce large batches of eggs until they are 5
6-days old; thereafter, eggs lose viability as the flies age. Mash and ammonia will stimulate oviposition, but flies are generally on an oviposition cycle where more eggs are deposited on a night cycle and 48 h peak rhythm. Eggs are collected in plastic weigh boats or

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from the strips placed in the cups. Moist egg masses can be placed into a centrifuge tube (10
15 mL) and the volume measured (0.1 mL contains 500
700 eggs). There are numerous recipes for housefly rearing medium. Two recipes for housefly larval medium are provided: (1) 3000 mL CSMA medium [commercial diet made of wheat bran (33.3%), alfalfa meal (26.7%), and brewer’s grain (40%), 3.25 oz dry yeast, and 25 mL malt mixed with a small amount of warm water to dissolve the yeast] is added to 2125 mL warm water and (2) 380 mL deionized water in a 500 mL flask, is completely dissolved 5 g sucrose in the water, then add 2.5 g dry yeast, continue to mix until dissolved, slowly add the sugar water
yeast mixture to 170 g dry Fly Larva Medium 5060, and mix well with a tongue depressor— the medium may be sterilized by autoclaving using a 1-L Nalgene brand PMP plastic jar. There is also the Gainesville HouseFly Diet that performs as well or better than the CSMA medium and can be mixed, bagged, and delivered by a local feed mill within 3 days.7 By adding pelleted peanut hulls 1:1 by volume, the housefly diet becomes suitable for rearing S. calcitrans. Larval medium is placed into weigh boats or cups. A small depression is made in the medium and the eggs are poured into the depression. The eggs in cups are covered with the larval rearing medium. The cups are covered with double cheesecloth secured with two rubber bands. The cups or weigh boats are placed in an adult fly cage and incubated overnight at 25.6 C
27.8 C and 60%
75% RH. The weigh boat is removed, and placed in a plastic bowl covered with a screen lid or stocking, or simply placed in a lidded plastic box. The lid of the box is modified by cutting two 2.5 in. circular holes in it, which are covered with screening. Larvae emerge in B18
24 h after eggs are laid. Instars: Instars may remain in the weigh boat or cup for 2
3 days. After a few days, they will become too crowded due to growth and will begin to run short of food. At this time, they are placed in a cylindrical plastic bowl filled 1/4
1/3 with larval medium. If the larvae in the weigh boat seem too crowded, some may be discarded. Do not place an entire batch of larval medium in the bowl all at once, as it may mold before it can be consumed. Unused medium may be stored in the refrigerator until needed. The larval bowl should be covered with a screen, stocking, or left in a plastic box as described earlier. As the larvae grow, they will need to be fed again. The timing and amount of food will depend on the number of larvae present. In some cases, it will be necessary to remove half the larvae and medium to a second bowl or cup in order to provide enough additional medium. Some larvae may need to be discarded in order to prevent crowded conditions. Overcrowded larvae develop into smaller adults. If the larvae are not too crowded, they may be given additional food by removing half or more of the old food from the top and replacing it with fresh medium using a tongue depressor. This may need

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to be done 2
3 times as the larvae develop. The maggots tend to tunnel down into the food to avoid the light. Throughout the larval growth phase, the containers should be checked on a daily basis to assess whether the larvae need more food or have begun to pupate. When the larvae begin to reach full size, their containers should be placed in plastic boxes with animal bedding covering the bottom and placed in a darkened area. As the larvae near pupation, most will crawl out of the containers and into the bedding to pupate. However, some will pupate in the top portion of the spent media. As larvae ascend for pupation in cups, B2
6 oz of fine vermiculite (No. 4 mesh, 4.76 mm) is added. The larval period is B7
9 days under laboratory conditions. Pupae: Puparia are formed in 2
3 days and should be collected after they have darkened to a maroon color. When most of the larvae have pupated, the animal bedding should be poured into a plastic bowl and placed in a cage. Puparia in cups can be sifted through a No. 10 mesh sieve (2 mm). Average size puparia weigh B2.1
2.5 g/100; 500
600 puparia fit in 30 mL of a volumetric cylindrical. Pupae are placed in cups that are placed in fly rearing cages. The bowls are allowed to remain in the cages until all (or enough) of the adults have emerged in B10
12 days. Once that happens, the bowls are covered with lids, removed, and frozen in order to kill any remaining flies. Once frozen, the spent medium should be placed in a ziploc bag which is placed in a sterilizable autoclave or biohazard bag for autoclaving. Rearing tips: Watch for insect contamination and fumigate, if necessary, with a mixture of carbon tetrachloride:ethylene dichloride (1:3 ratio). Watch for fungal growth. Keep rearing room clear of flies with fly ribbons, fly swatter, and, if necessary, 0.3% Dipterex liquid bait. No eggs: flies too young, humidity too low. Hard crust on larval medium: humidity too low, too few maggots, medium too dry. Mold: all medium has some mold, but excessive mold is due to medium being too wet or too dry. Cold medium: yeast is too old, yeast made with cold water or placed in a cold area. No larvae: dry eggs used, old yeast. After 10
14 days, the number of emerged adult flies is counted and the percent mortality calculated.

In vitro method(s) Contact—filter paper—adult The objective of this bioassay is to determine the concentration of a parasiticide residue on a filter to provide knockdown and mortality of adult houseflies. Filter papers placed in the lids of a Petri dish cover are treated with serial dilutions of test compound (0.0108
4.3 μg/cm2) dissolved in acetone and allowed to dry at RT in a fume hood. Ten houseflies (anesthetized with CO2) are placed in the bottom dish cover and the treated

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lid cover is applied. Once the flies revive, the Petri dish is inverted so the flies are in contact with the treated filter paper. The dishes are incubated at 27.5 C and 60%
75% RH. Knockdown and mortality are recorded daily up to 3 days of exposure. Contact—glass—adult The objective of this bioassay is to determine the concentration of a parasiticide residue on a glass surface to provide knockdown and mortality of adult houseflies. Petri dish lid covers are treated with serial dilutions of test compound (0.0108
4.3 μg/cm2) dissolved in acetone and allowed to dry at RT in a fume hood. Ten houseflies (anesthetized with CO2) are placed in the bottom dish cover and the treated lid cover is applied. The dishes are incubated at 26.7 C
27.5 C and 60%
75% RH. Knockdown and mortality are recorded daily up to 3 days of exposure. Contact—glass chamber/spray—adult This test8 is conducted in a glass chamber measuring 70 3 70 3 70 cm. A total of 20 laboratory cultured sucrose-fed adult female houseflies are released into the chamber. The test chemical is sprayed into the chamber by using a manual/electric atomizer. The discharge rate (g/spray) of the sprayer is predetermined. Based on the dosage required, an estimated time of spray is discharged into the glass chamber. Knockdown of houseflies is observed at the indicated intervals up to 20 min. After 20 min, all flies are then collected and placed in cylindrical polyethylene containers with 10% sucrose pad. A further, single assessment of knockdown is made after 60 min. Mortality is observed after 24 h PT. All tests are to be conducted at a temperature of 26 C
28 C and 65%
85% RH. A minimum of three tests are conducted. The knockdown values (KT50 and KT95) and regression slope are obtained using probit analysis. Mean percentage of insect mortality value is subjected to arcsine transformation followed by comparison of means using the LSD test. Contact—Peet-Grady chamber/spray—adult fly This test also by Zairi and Lee8 is conducted in a Peet-Grady chamber measuring 180 3 180 3 180 cm. A total of 50 adult female houseflies are released into the chamber. The test chemical is sprayed into the chamber by using a manual/electric atomizer. The discharge rate (g/spray) of the sprayer is predetermined. Based on the dosage required, an estimated time of spray is discharged into the Peet-Grady chamber through two introduction ports of the chamber. Knockdown of houseflies is observed at the indicated intervals up to 20 min after which all flies are then collected and placed in cylindrical polyethylene containers with 10% sucrose pad. A further, single assessment of knockdown is made after 60 min. Mortality is observed after 24 h PT. All

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tests are to be conducted at a temperature of 26 C
28 C and 65%
85% RH. A minimum of three tests are conducted. The knockdown values (KT50 and KT95) and regression slope are obtained using probit analysis. Mean percentage of insect mortality value is subjected to arcsine transformation followed by comparison of means using the LSD test. Contact—spray—adult The objective of this bioassay is to determine the knockdown and mortality of a parasiticide spray applied directly to adult houseflies. Test cages are constructed of cylindrical screens [No. 20-mesh (0.841 mm) stainless steel] 1.5 in. in diameter by 4 in. long with size D polyurethane foam tube plugs on each end. Ten adult houseflies (anesthetized with CO2) are counted and placed into cages. The cages are inserted into a plastic socket mounted on a turntable in a spray hood. The flies are sprayed with test and positive control (tetrachlorvinphos) compounds using a siphon type atomizer. Flies are provided sucrose solution by draping a paper wick over the outside of the screen cylinder. Knockdown and mortality are recorded daily up to 3 days PT. Contact/Ingestion—dental wick—adult IRAC Susceptible Test Method 026, Version 1, May 2011. Serial dilutions of compounds are prepared in 20% w/v sucrose. Compound solution (B5 mL) is required for each concentration. The sucrose solution is also used as a control. A piece of dental wick (2 cm) is placed in a 175 mL plastic container and treated with 1.2 mL of the sucrose solution with or without compound. Ten uniform size female flies (immobilized with CO2) are placed into each container (three replicates/concentration) and the containers are covered with ceaprene foam plugs. Mortality is observed and recorded after 48 h at 23 6 2 C, 50% RH, and 12L:12D h photoperiod. At assessment, flies are classified as either (1) unaffected, giving a normal response (making a coordinated move) when gently stimulated by dropping down the containers, or (2) dead or affected, the latter giving an abnormal response to stimulation or showing abnormal movement. Thus percent response or mortality will include both dead and affected flies. Mortality of untreated flies should be recorded. Express results as percent mortality and correct untreated mortality using Abbott’s formula.9 Precautions and notes 1. Disposable plastic equipment is preferred provided that it is not affected by the formulation constituents; glass equipment may be used but must be adequately cleaned with an appropriate organic solvent before use. To avoid higher control mortality, it is necessary to use containers with adequate volume as recommended. Smaller containers lead to higher control mortality.

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2. Insecticide products (formulations) may contain varying concentrations of active ingredient(s). Ensure insecticide dilutions are based on active ingredient content. 3. Where possible, bioassays to measure the variation in insecticide susceptibility should run in parallel with a bioassay to measure the susceptibility of a known susceptible standard population of houseflies (e.g., WHO-N).

Contact/Ingestion—larval rearing medium—eggs or first instars The objective of this bioassay is to determine the effective concentration of a parasiticide in larval rearing medium to provide 100% mortality against eggs or first instars of the housefly. Experimental parasiticides with potential insect growth regulating activity are formulated and mixed in housefly larval rearing medium. Fly rearing medium (100 g) is weighed out in a Styrofoam testing cup (6
8 oz). Serial dilutions of the parasiticides are mixed in with the rearing medium. Treated and untreated cups are seeded on the surface with 25 housefly eggs or first instars. The cups are covered with two-layer cheesecloth affixed with a rubber band. The cups are incubated at 28 C
30 C for 14 days until adult fly emergence is completed. The treated and control cups are frozen (10 min in a freezer) to kill the live flies, opened, and the adult flies are counted. Efficacy is determined by comparing the number of adult flies in the treated cups to the number of adult flies in the control cups (quality control: no more than 5% death). Endpoint data recorded as ED100 in mg/g.

Contact/Ingestion—treated bovine manure—first instars The objective of this bioassay is to determine the effective concentration of a parasiticide in treated bovine manure to provide mortality against first instars of the housefly. Experimental parasiticides with potential insect growth regulating activity and diflubenzuron as a positive control are formulated on a lactose carrier mixed in fresh bovine manure at 0.2, 1, 5, 25, and 125 ppm. Twenty first-instars (2
4-h old) are seeded onto duplicate 80 g manure samples of each treated concentration plus the positive control, untreated, and lactose carrier control samples. The samples are stored at 25 C and 60%
75% RH with a 12L:12D h cycle. Moisture is provided as needed to allow normal larval development (2
3 weeks). Upon eclosion of adult houseflies from the untreated control samples, the number of pupae and eclosed adult houseflies from each treatment and control groups is counted. The percent reduction in development and emergence is determined and dose-response LD50 and LD90 values are calculated by log probit analysis, if sufficient data are generated. Diflubenzuron typically provides 90% reduction in adult fly emergence at 1 ppm in susceptible houseflies.

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Contact/Ingestion—treated poultry manure—first instars The objective of this poultry oral larvicide bioassay is to determine the effective concentration of a parasiticide as an oral feed-through larvicide to provide mortality against first instars of the housefly. Standard chicken laying ration is mixed with a candidate and positive control (cyromazine) compound. The formulated feed mixture is fed to laying hens (white Leghorn hens, B23
25 week old) for 3 days within the treatment (candidate compound and positive control) and control groups. Fecal collection boards are placed under the cages of the hens. Fecal samples are collected from each bird at 48 and 96 h intervals. Individual fecal samples are homogenized and either moistened or allowed to dry depending on their moisture content. Eighty grams of each sample are placed in Styrofoam cups. Twenty first instars (2
4-h old) of M. domestica are placed on the surface of the feces. The cup is covered with a double layer thickness of cheesecloth and incubated at 26.7 C
27.5 C and 60%
75% RH for 2
3 weeks. Upon eclosion of adult houseflies from the untreated control samples, the number of pupae and eclosed adult houseflies from each treatment and control groups are counted. The percent reduction in development and emergence is determined and dose-response LD50 and LD90 values are calculated by log probit analysis, if sufficient data are generated. Contact/Ingestion—treated poultry manure—second instars The objective of this bioassay is to determine the effective concentration of a parasiticide in treated poultry manure to provide mortality against second instars of the housefly. Experimental parasiticides with potential insect growth regulating activity and diflubenzuron as a positive control are mixed in fresh poultry manure at 0.8, 1.6, 3.1, 6.2, 12.5, 25, 50, and 100 ppm. Triplicate 50 g samples of fresh poultry manure of each treated concentration plus the positive control and untreated control are seeded with 35 second instars (2-day old). Second instars are easier to handle and have less unexplained natural mortality, probably from handling, than first instars. The poultry manure samples are stored at 25 C and 60%
75% RH with a 12L:12D hour cycle. Adult fly emergence is recorded 3 weeks postseeding of the samples with larvae. The percent reduction in development and emergence is determined and dose-response LD50 and LD90 values are calculated by log probit analysis, if sufficient data are generated. The LD50 value of diflubenzuron is B3 ppm. Repellency Y-tube olfactory assay—filter paper—adult Flies were individually introduced into the base of a 20 cm straight glass tube section of a Y-tube olfactometer without anesthetization.10 Flies introduced into the lower port traveled upwind at a 20 degree incline and made a locomotor choice at the

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Y split. Flies were recorded as having made a definitive choice when they had traveled 15 cm up to a Y-tube arm. With the Y-tube olfactometer mounted at a 20 degree incline and the flow rate of incoming, charcoalfiltered line air was 0.2 LPM. The 20 degrees was sufficient to elicit a negative geotaxis behavior in flies and resulted in translocation up toward the arm-end of the Y-tube. To eliminate effects of visual stimuli, the entire Y-tube and airflow regulating apparatus was housed in a light-exclusion box. The Y-tube was back-lit with red (660 nm) LEDs and fly choice inside the enclosed apparatus was monitored with a video camera. Prior to the introduction of each fly, a 2.5 cm diameter filter paper disk impregnated with either treatment or solvent control solutions was inserted into each Y-tube arm odor source adapter. Flies were allowed 2 min to travel 15 cm up into the end of one arm of the Y-tube. Flies that failed to make a choice within 2 min were discarded. Test solution placement was alternated between arms with each trial; 60 male and 60 female flies were assayed for each test solution. All glassware was rinsed with ethanol and DI H2O prior to each round of testing per treatment pinene (1, 2) concentration and cleared of odors for B2 min between each fly tested. Two replicates consisting of 30 male and 30 female flies were run for each of the 7 tested pinene (1, 2) concentrations for each enantiomer, the 29% DEET solution, and the ethanol:H2O (95:5 v/v) solvent control. Y-tube arm choice responses were analyzed using a G-test at an error rate of 5%. Olfactometer bioassay—filter paper—adult Tests are conducted according to the methods and test conditions of Carlson et al.11,12 Basically, filtered, humidified outside air is conducted through two ports in the front face of a Plexiglas cage (90 3 45 3 54 cm) and out through a screen that forms the rear face of the cage. A horizontal glass cylindrical trap fitted with a vertical screen at its midpoint and an inverted screen funnel at its inner end was inserted into each port. A hexane solution containing 50 pg of the test compound was applied to filter paper positioned in the outer end of one of the cylindrical traps, while 50 pg of muscalure on filter paper was similarly placed in the other trap. Incoming air passed sequentially over the samples, through the traps, and past 300 male, 24
36-h-old Cradson-P houseflies in the cage before exiting. Counts were made of flies entering the traps in 30 min. The flies were allowed 30 min for recovery between tests and were used for 2 days. The activity score for each analog was calculated by dividing the percentage of flies trapped by the analog by the percentage of flies trapped by muscalure (85% cis, 15% trans) and multiplying by 100. It therefore represents activity relative to that of muscalure, which was assigned an activity score of 100. In another series, each chemical in one trap was tested against a blank trap (no chemical) and captures were compared with those made by muscalure identically tested.

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Contact—filter paper—adult Flies are introduced, without anesthetization, into the lower end of a contact repellometer, which consisted of a 20 3 2.5 cm section of PVC pipe mounted on a ring stand at a 20 degrees incline. Flies introduced into the lower end of the contact repellency assay tube travel up a 20 cm segment of PVC tube toward a light source at a 20 degree incline. The inner lateral sides of the terminal perforated 8.5 cm section of tube are lined with treatment and control solution-saturated filter paper. The amount of time flies spend in contact with each filter paper-lined side of the perforated tube section was recorded. A perforated section of 8.5 3 2.5 cm plastic cylinder is affixed in line to the upper end of the inclined pipe. Two 3.75 3 8.5 cm pieces of filter paper line the inner lateral surfaces of the end cylinder, with a 3
4 mm gap between them along the dorsal and ventral midlines of the apparatus. For each trial, one filter paper insert is saturated with 0.45% pinene solution, and the other with ethanol: H2O (95:5% v/v) solvent control solution. A lamp with a 40 W incandescent bulb is placed 10 cm from the opening of the upper end of the pipe to provide a visual stimulus for flies. Each fly introduced into the lower end of the apparatus experience 20 cm of unbiased travel space and then 8.5 cm during which it can choose to have its tarsi in contact with ethanol or pinene solution
saturated filter paper, each lining one side of the perforated section of the tube. A video of each individual trial is recorded and analyzed for time spent on each filter paper insert as flies passed through the distal filter papercontaining end of the tube. Three replicates, each consisting 10 male and 10 female flies, are conducted.10 Electroantennagram assay—filter paper—adult Whole, living female flies are positioned and restrained on a glass platform using Tackiwax. Electroantennagram (EAG) recordings are obtained by inserting a glass pipette Ag/AgCl electrode filled with 0.1 M KCl into the base of the antenna at the first antennal segment. A second recording electrode is inserted into the distal end of the funiculus of the same antenna. Signals are amplified 10 3 using a Model 3100 Electrometer and visualized and recorded using LabChart 8 software. The stimulus delivery is adapted and revised from Visser and Piron.13 An airstream of 40
50 cm3/s is continuously blown over the antenna prior to the delivery of the stimulus. Air is purified through carbon and HEPA-VENT filtration and then rehumidified by bubbling through distilled water. The stimulus is delivered for 1 s using a timed stimulus delivery system that consists of a volatile chamber connected to the main airstream. The volatile release into the purified airstream is under valve control. Each volatile chamber contained a 2.5 cm diameter filter paper disk saturated with pure volatile (100 μL). Paraffin oil is used as the blank control. Each stimulus is separated by 30 s intervals. Antennal preparations are discarded after 25 min. EAG amplitudes are normalized by dividing the magnitudes

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elicited by 1 and 2 by the mean amplitude elicited by the 95% ethanol control solution.10 Pseudofly Petri dish bioassay—adult These tests14,15 are based on the mating behavior of houseflies. Provided with the proper stimuli, males leap onto the backs of other flies or fly models and exhibit a behavioral sequence typical of that observed in their normal mating. The fly model in the pseudofly Petri dish test is a fly-sized knot cut from a black shoelace and cemented to the center of a 9-cm plastic Petri dish. The knot (pseudofly) is then treated with 0.03 mL of a hexane solution containing 30 μg of the test chemical or with 0.03 mL of hexane alone. After the solvent has evaporated, two male houseflies, 3
10 days-old, are introduced into the dish that is covered. In any single session, six dishes are used for each test material to minimize variations in the behavior of individual flies. Light (35 footcandles, incandescent) and temperature (25 C) are controlled in the observation room, and dishes are arrayed in groups and observed sequentially for six 3-min periods, with data being recorded for each period. Observations on each test material at any one session are pooled, and the numbers are converted to strikes/fly/h. The test array permitted comparison of 12 variants during any observation session. Data from three to four such sessions were fully amenable to statistical treatment. Data from each test session were converted to a ratio to the mean of all data taken to remove variation normally found between sessions. Analysis of variance and the Dunnett “t” separation of means test were used to determine the significance of differences between test materials and muscalure. Individual test means were corrected by subtracting values for the blanks and converting such corrected means to a percentage of the values for muscalure (5l00).

In vivo method(s) Parasiticide efficacy testing against nuisance flies on ruminants should be designed and conducted in accordance with the WAAVP guidelines for evaluating the efficacy of ectoparasiticides against biting and nuisance flies on ruminants (see Appendix A of Ref. [16]). The housefly is by far the most common ectoparasite pest on horses and an important intermediate host for Habronema muscae.17,18 Hundreds of houseflies may be seen feeding around the eyes, nostrils, anus, genital openings, and wounds. Feed-through larvicide products containing cyromazine, diflubenzuron, or tetrachlorvinphos provide good fly development control in isolated facilities where the primary fly breeding medium is horse manure. On-animal repellents as well as pyrethrins combined with residual pyrethroids may provide the longest efficacy following a single application. Face flies (M. autumnalis) are occasional pests of horses, particularly near pastured cattle. They feed on secretions around the eyes and the face. A study

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was conducted to estimate the effects of flying insects (Order Diptera) on the behavior of grazing horses in relation to the use of insecticide and repellent substances.19 The investigations were done between June and August in 2008 in three periods of 7 days each. An insecticide and two repellent substances were used: “Well-care emulsion” containing permethrin and “Bremsen-Frei-Plus” based on etheric oils. Both groups were compared with a nontreated control group in a crossover design. Each group (n 5 3
5) was used alternately as control and treatment groups. Several climate parameters were taken during the study. Furthermore, the number of insects per animal was estimated at certain times. Once per observation period, insects were caught using Malaise traps and differentiated by species. The proportion of horse relevant species caught of the order Diptera, especially Culex pipiens and M. autumnalis, was 9% on average. There was no correlation between the number of Tabanidae caught in the Malaise traps and the number observed near by the horses. Behavior parameters, such as tail swishing, headshaking, stamping, tail switching, snapping at the body, and moving, were observed more frequently with increasing insect infestation. When horses were infested with a high number of flying insects, feeding activity was significant lower, whereas locomotion activity was significant higher. Both substances had positive effects for B50 h PT with no apparent difference between the substances. However, a lower frequency of headshaking and tail swishing could be observed in the permethrin-treated horses.

Musca autumnalis De Geer, 1776—face fly Biology and life cycle The face fly is distributed throughout most of Europe, Central Asia, northern India, Pakistan, China, parts of North Africa, and North America (temperate parts of the United States and southern Canada). M. autumnalis transmits the nematode Thelazia spp. to the eyes of cattle. The face fly is also an important mechanical vector in the transmission of pinkeye (M. bovis) to cattle. Parafilaria spp., a filarid nematode causing hemorrhagic nodules in the skin of horses, deer, cattle, and buffalo, is also transmitted by Musca spp. Nuisance (fly worry) caused by the face fly can be considerable. The adult is very similar in size and appearance to the housefly. However, the abdomen of the female face fly is darker, and the male abdomen has a medium black band with a typically yellowish-orange along the sides. The adult fly is grayish and measures 6
9 mm long 3 2
3 mm wide. The thorax bears four longitudinal black stripes on the dorsum. Female flies lay eggs singly with the respiratory mast up in the upper surface of fresh cattle manure. The eggs are 0.5 mm wide 3 3 mm long possessing a dark respiratory mast B0.7 mm in length.

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The larva is white, maggot shaped, with a single, medium black mouthhook. The mature third instar is lemon yellow and disperses from manure to pupate in surrounding soil. The puparium is uniquely white, measuring 6
10 mm in length. The entire life cycle from egg to adult is 12
20 days at 20 C
26 C.20

Rearing method(s) Fales21 found cow manure to be the only medium that sustained larvae of M. autumnalis to maturity. Rearing methods of M. autumnalis are included herein with notable method differences with those of M. domestica.4 Laboratory culture method for Musca autumnalis De Geer (Diptera: Muscidae) The following procedure is a standardized method for rearing M. autumnalis at Inveresk Research (OECD Guidelines for Testing of Chemicals Feb 26, 2007, www.oecd.org/chemicalsafety/testing/38207891.pdf). Housing and environmental conditions Cultures are housed in plastic chambers (B50 3 50 3 50 cm) with externally mounted heater boxes. Environmental conditions are 30 6 2 C and .60% RH. All developmental timings reported in this method are based upon rearing at this temperature and must be reassessed if a lower temperature regime is implemented. The lighting regime is 16L:8D hour. In addition to fluorescent lighting, an incandescent light source (such as a tungsten filament spotlight) may be provided to create an area in which the flies can bask. Feeding Water is provided ad libitum in each cage by inverting a waterfilled beaker onto a tray lined with absorbent paper. Dried egg yolk powder, milk powder, and sucrose (1:1:1 ratio) are provided ad libitum. Honey solution soaked cotton wool (25% honey solution w/v) is provided and replenished about twice weekly. Fresh pig liver is provided weekly as an additional protein source for female flies (it is predominately the females that feed upon the protein-rich facial secretions of livestock). Strips of pig liver are hung from hooks on the walls of the cage. Oviposition Dung is collected from cattle with a known veterinary history. The cattle are not treated with any pharmaceutical products for at least 8 weeks, or with an anthelminthic for at least 5
6 months, prior to collection. Dung is frozen at 220 C upon collection and stored at this temperature until the day before use. Dung is defrosted at RT for 24 h before addition to the culture. In an unsynchronized culture, bovine dung is provided weekly. In a synchronized culture, bovine dung is provided when adult flies are between 7 and 10 days old, 3
4 days after copulation is first observed. The defrosted

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dung is homogenized using a laboratory mixer for B5
10 min before addition to the culture. The dung should be wet enough to be easily molded into a 7 cm diameter ball, but dry enough that the ball will retain its shape. This ball is dropped onto a plastic tray to produce an artificial pat. The pat is then placed into the culture. Each batch of egged manure is transferred onto 1 kg bovine dung in a plastic bucket. If egg densities are high, the eggs should be divided up to ensure that no more than 500 eggs are transferred to each 1 kg batch. Forty-eight hours after oviposition, 3 cm of sawdust is added to the dung surface, and a fine mesh or muslin cloth (nappy or diaper liner is ideal) secured over the top of the bucket. Larvae will migrate to the dung surface and into the sawdust to pupate. Life cycle and developmental timing Eggs are laid both singly and in clumps. Eggs are primarily deposited under the dung surface, and only the terminal respiratory horn is apparent. Clumps of eggs can by removed from the dung and gently teased apart for experimental use. Egg hatch occurs after 24
36 h. If larvae are required they are removed from the dung at 48 h after oviposition. There are three instars, third instars are cylindrical yellowishwhite maggots that taper anteriorly and are B12 mm long. Larvae usually “wander” at B4 days after egg hatch, migrating to the dung surface and sawdust layer to pupate. Pupae can be removed from the sawdust at 6 days after oviposition and placed in the culture or be allowed to emerge from the dung naturally. Pupae are white/gray and 5
7 mm long. Adult eclosion occurs at 4
5 days after pupal formation. Therefore egg-to-adult development takes 10
11 days at 30 C. At 25 C development takes B17 days. The female adult flies are easily distinguished from the adult males by the proximity of the eyes, the eyes of the males almost touch, while those of the females are distinct. Adult face flies can be reared on a mixed dry diet of powdered egg, powdered milk and sugar, and free-choice water.22 Larvae are reared in fresh bovine feces mixed with B80% water. The insectary is maintained on a 16L:8D hour cycle at 25 C
28 C and 50%
60% RH. Modified adult cages, shallow larval rearing pans, dry pupation pans, and specially made shelves and tables minimized handling of larval, pupal, and adult stages.

In vitro method(s) Many of the contact bioassays described for M. domestica are applicable to M. autumnalis. Contact—topical—adults Insecticides are applied topically in acetone to female adult flies of a laboratory colony of M. autumnalis with LD50 values in μg/g of fly, as calculated from 24-h mortality counts.23 Poor control of M. autumnalis in the field may

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be due to the fact that the flies do not come into contact with insecticides long enough to acquire a lethal dose and that reinfestation from untreated animals is rapid. Animals should therefore be treated more often against M. autumnalis than against some other pests. Contact—cages—adults The duration of effectiveness of test compounds is determined by placing flies 6
7-days-old adults of a laboratory colony of M. autumnalis in cylindrical wire cages that had been dipped in acetone solutions of the toxicants, allowed to dry, and then hung on lines indoors or outdoors during the winter or summer.23 The indoor cages were tested six times, 1
49 days PT, and the outdoor winter and summer cages four times, 7
97 and 10
48 days PT, respectively. Mortality counts are made after exposure for 18 h, and the toxicants ranked for duration of effectiveness.

In vivo method(s) Parasiticide efficacy testing against nuisance flies on ruminants should be designed and conducted in accordance with the WAAVP guidelines for evaluating the efficacy of ectoparasiticides against biting and nuisance flies on ruminants (see APPENDIX A of Ref. [16]). Fannia canicularis Linnaeus, 1761—lesser or little housefly Biology and life cycle F. canicularis, the lesser or little housefly, originally from Europe, is now cosmopolitan. It occurs throughout the United States, all states and territories of Australia except the Northern Territory, China, the United Kingdom, Ireland, and Scandinavia and is widely distributed throughout the world. The little housefly rarely feeds directly on animals but may be attracted to smeared feces, sweat, and mucus on an animal. They occur near excrement and can be abundant in poultry, cattle, and dairy facilities. Flies may become extremely abundant under wet conditions constituting an annoyance. They may mechanically transmit pathogenic organisms. The little housefly transmits the nematode Thelazia spp. to the eyes of dogs. It is a pest on mink farms and poultry facilities, particularly caged layer facilities. The life cycle is generally similar to the housefly, but development time from egg to adult is longer, minimum of 22
27 days. Eggs are deposited on the surface of wet, decaying animal or vegetable matter. The adult fly superficially resembles the housefly but is smaller (5
6 mm in length), grayish with three dark longitudinal strips on the dorsum of the thorax, a more yellow abdomen, and relatively larger wings with distinct venation.

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The eggs are elongate with narrow lateral wing-like processes and hatch in B25 h. Instars are 5
6 mm in length, flattened with dorsal and lateral spine-like projections that increase in size toward the base and terminate in single points. The larvae are filter feeders living in or under wet decaying vegetation and among dead decaying insects. Larval development takes B7 days. Pupae resemble the spiny larvae. Pupae develop in drier areas and in B7 days.

Rearing method(s) Adult flies are maintained at 10 C and 65% RH in a 12 in2 screened cage with plywood and front circular opening fitted with a muslin sleeve.24 A cotton pad saturated in 20% sugar solution is provided as adult food. Eggs are laid on 2-day-old pads impregnated with 2.25 g protein hydrolysate and 50 mL water. Pads with 200
300 eggs are removed from Petri dishes in the cage and placed on larval rearing medium (30 g dried Brewer’s yeast, 112 g finely ground laboratory chow, 225 g milled red wheat bran, and 12.5 mL concentrated dimalt, and 598 mL water) in a 3.78-L jar covered with muslin. Pupae develop at the top of the rearing medium that is placed into cages to allow adult flies to emerge. The complete life cycle from egg to egg is B24
29 days at 10 C and 65% RH. The larval period is 8
10 days, while the pupation period is 8
12 days with adult eclosion in 36
48 h. The little housefly can also be reared at 26.6 C and 65% RH.25 Adult flies fed on molasses and canned evaporated milk solution both diluted 1:1 with water. The solution is provided in 5 oz paper cups with toweling and replaced twice weekly. Oviposition occurs on the toweling saturated in household ammonia diluted with equal amounts of water and CSMA medium. Collected eggs are placed on 1.5 in. deep larval rearing medium (1 yeast cake/1000 mL water) and 20% molasses solution mixed together and added to dry CSMA medium mixed to near saturation in 6 in. battery jars; 100 larvae/jar. Pupae are collected on tightly rolled cardboard on the larval medium surface or by washing the larval medium through a coneshaped strainer. The complete life cycle from egg to adult is B18.5
22 days at 26.6 C and 65% RH; egg to adult is 22
27 days. The larval period is 8
10 days, while the pupation period is 9
10 days with adult eclosion in 1.5
2 days. A final rearing method includes adult flies in cages fed a diet of four parts sugar, 1 part powdered milk, and 1 part powdered egg. Water is provided via a water wick. Eggs are recovered by wrapping a small portion of chicken feces in a fine, mesh cloth and placing in a small cup. Typically, 0.1 mL of eggs equals B600 eggs. The eggs are reared in a medium consisting of 300 g dry CSMA, 26 g dry yeast, 600 mL water, and 50 g powdered milk. These ingredients are mixed, then distributed evenly in three 32 oz

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containers filled approximately half full. The cups are covered with vented lids and set aside for 48 h. After 48 h the upper portion of the contents are stirred to break up mold, then 200
300 eggs are added to each container. Adult flies emerge in B21 days at 26.6 C and 65% RH.

In vitro method(s) Many of the contact assays described for M. domestica and M. autumnalis are applicable to F. canicularis. Feeding—treated poultry manure—first instars Standard chicken laying ration is mixed with a candidate and positive control (cyromazine) compound. The formulated poultry oral larvicide feed mixture is fed to laying hens (white Leghorn hens, B23
25-weeks old) for 3 days within the treatment (candidate compound and positive control) and control groups. Fecal collection boards are placed under the cages of the hens. Fecal samples are collected from each bird at 48 and 96 h intervals. Individual fecal samples are homogenized and either moistened or allowed to dry depending on their moisture content. Eighty grams of each sample are placed in Styrofoam cups. Twenty first instars (2
4-h old) of F. canicularis are placed on the surface of the feces. The cup is covered with a double layer thickness of cheesecloth and incubated at 26.7 C
27.5 C and 60%
75% RH.

In vivo method(s) Ronnel and dimethoate produces rapid knockdown of the little housefly held in large cages.26 Fly mortality from residual insecticide was high for a 2-week period following application. Malathion toxicity was low due to resistance. Black light traps reduce the lesser housefly populations and the number of ovipositions in the dropping pits in semiopen poultry houses.27 Glossina spp. Wiedemann, 1830—tsetse flies Biology and life cycle There are B30 known species and subspecies of tsetse flies belonging to the genus Glossina. Tsetse flies are confined in tropical and subtropical Africa although two tsetse fly species have been reported in southwestern Saudi Arabia.28 Tsetse fly’s transmission of trypanosomes by both sexes cause fatal nagana disease in domestic cattle and African sleeping sickness in humans. Only nine species and subspecies, belonging to either the Glossina palpalis or the Glossina morsitans group, are known to transmit sleeping sickness. The adult fly measures 6
15 mm in length and is a honey-brown dipteran about the size of a housefly. Their wings have a characteristic venation

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(the discal or “hatchet” cell is cleaver shaped) and fold completely over one another covering the abdomen. The antennae have a long branched arista that is bristled or feathered along one edge. Both male and female flies are voracious blood feeders and are usually telmophagous (i.e., feeding from a pool of blood caused by tissue laceration). Tsetse flies spend a greater part of their life resting in cool, shady places provided by trees, holes, etc. Eggs are produced in the ovary with only one egg fertilized at a time and then pass into the uterus one at a time. The larvae hatch and feed from the secretions of the milk glands. This unique reproductive mode of insects is called adentropic viviparity meaning “gland fed, live birth.” Larval development of all three instars usually requires 9
12 days and is completed in the abdomen of the female fly. The third instar is expelled, crawls a short distance usually to a shaded site, and burrows into the soil to pupate. The pupal stage develops in B20
34 days at 23 C
24 C. Tsetse flies spend B50% of their life span under the ground as pupae. Peak female emergence occurs at B32 days after pupation. Tsetse flies typically produce B4 generations/year and may have up to 31 generations during their life span. Female flies mate only once in their life.

Rearing method(s) Laboratory rearing A comprehensive standard operating procedure for the mass rearing of tsetse flies is provided by the FAO/IAEA SOP29 for mass-rearing of tsetse flies. Briefly, pupae are field collected by sieving sand or soil from a breeding site using a coarse flour sieve. Collected pupae are stored in a vial or matchbox lined with cotton wool. Pupae are transferred to the insectary, placed in an emergence cage in a quarantine area (may be infected with trypanosomes), and maintained at 23 C
24 C and 75%
80% RH for 35
40 days. Emergence occurs around the end of the pupal maintenance period. Flies are collected using a separating tube and males and females identified. Keep female and male flies in separate cages and feed using a Trolley membrane feeding system. Switch on a heating polyvinyl chloride mat or aluminum plate with an adjustable heat source. Transfer sterile feeding trays (anodized aluminum, diamond-shaped surface with a 1-cm edge) and silicone membranes from an oven onto the heating mat or plate. Monitor the temperature of the heating mat or plate using a thermocouple surface thermometer. At a heating mat or plate temperature of 35 C
37 C, lift two-thirds of the membrane at one corner of the tray, pour 100 mL of blood into the tray, and immediately replace the membrane. Avoid the formation of bubbles while pouring blood. Distribute the blood evenly on the feeding tray using a roll of tissue paper. When the membrane surface temperature has returned to 35 C
37 C, manually put the fly cage on the membrane.

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Invert the cage so that the normally upper surface faces downward, enabling the flies to feed through clean netting. Dim the lights in the feeding room. If the temperature of the heating mat or plate is $ 39 C, remove the cage and wait until the temperature returns to 35 C
37 C before replacing the cage. If the flies are reluctant to feed, cover the cage with black and/or damp cloth. Allow the flies to feed for 10
15 min. After feeding, return the fly cages (invert so the upper side is on top) to the holding room at 23 C
25 C and 75%
85% RH, depending on the species. When flies of the appropriate age are not available, keep females and males in a 1:1 ratio in the same cage starting 3 days after emergence. Remove the males after 7
10 days. If fly ages are known, cage 3-day-old females together with $ 10-day-old males at a female:male ratio of 1:1. Remove males after 3
4 days. On day 25 after larviposition, transfer pupae to Petri dishes and place in an emergence cage.30 A silicone membrane, described by Bauer and Wetzel31 was modified and developed to meet the feeding requirements of G. palpalis.32 The thoroughly mixed ingredients, silicone paste RTV-M 539 and hardener T30, are placed on a polyvinyl chloride mold (42 3 42 cm) with a peripheral groove, 6 mm deep and wide; thus the membrane, when finished, has a marginal wall of the same size. The peripheral wall makes handling easier and prevents direct contact between the fly cages and the blood as happened formally when the fly cages protruded beyond the membrane. The ingredients are then covered with a sheet of polyethylene, and olive oil is used to prevent the membrane from sticking either to the mold or to the polyethylene sheet when it is rolled into final form. As the feeding response of G. palpalis was found to be lower than that of G. morsitans on the type of membrane developed for G. morsitans, this membrane contains only 1.6 g (2%) hardener added to 80 g paste, while the wall contains 4.8 g (4%) hardener added to 120 g paste. The thickness of the membrane is 480
560 μm. Flies (50/cage) were transferred directly to in vitro feeding on defibrinated pig blood at 36 C
38 C 6 days a week. The temperature of the breeding room was 24 C and 85%
95% RH. The flies were kept in cages 12.5 cm in diameter and 4.5 cm in height, covered with terylene netting. Bacterial infections of Escherichia coli and Staphylococcus spp. from the blood and membranes (breakdown of the heating system used to prepare the membranes, i.e., 25 C instead of 80 C) caused high adult fly mortality. Nevertheless, the method demonstrated that in vitro maintenance of G. palpalis on defibrinated pig blood is feasible. Adult female flies of Glossina longipennis were caught in the field and brought to the laboratory for rearing.33 They were kept in a climatecontrolled room at 25 6 0.5 C and 80%
85% RH. The tsetse flies were fed on the ears of lop-eared rabbits daily except on weekends. A total of 174 pupae, mean weight 79.5 6 1.8 mg, were produced by these field-caught tsetse flies. Tsetse flies that emerged from these pupae formed the parental

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stock of the colony. The number of mated females in the colony increased and reached the target of 1500 breeding females. The current rearing techniques follow.33 Adults and pupae are kept in a climate-controlled room as abovementioned. Pupae are kept in l0 3 40 3 40 cm emergence cages, up to 2000 pupae/cage. The newly emerged tsetse flies are immobilized at 2 C
3 C and sexed. The sexes are put separately into Geigy cages, 15/cage. Each cage, 15 3 8.5 3 5 cm, consists of a stainless steel wire frame having a closed end fitted with a corked hole. The cage is covered with black terylene netting, 4 3 4 mm mesh, that enables the larvae to crawl through into a tray. Teneral females, 15 in each cage, are fed on rabbits’ ears and then 16 recently fed 7
10-day-old males are introduced into each cage for mating. After B72 h, the males are removed from each cage using a 35 cm long, 1.5 cm diameter glass tube. The mating period of 72 h is used as after 48 h most tsetse flies are still mating and even after 72 h, a few pairs are still in copula. These are left to continue mating and may remain so for up to 7 days. The cages of mated females are kept on a rack.34 Each rack of 10 cages forms a unit of 150 mated females and is given a code number. The flies are offered food for 5 days each week. Pupae are also collected from the trays of all the racks on these days, and the number of pupae from each unit code is recorded; pupae from one of the codes units are weighed in bulk. Dead flies are removed on Mondays, Wednesdays, and Fridays. Female flies are maintained for 80 days, and on day 81 postemergence the surviving flies are killed. In 1989 the mean female stock of the G. longipennis colony was 1579. The insemination rate was B99%. The colony produced 30 pupae, that is, 0.37 pupae/female/week, which is lower compared with that of a colony of related Glossina brevipalpi.35 The mean pupal weight was 73.4 6 0.3 mg. Mean daily female mortality was 1.5%. Mean survival by day 80 postemergence was 17.6% in the first half and 31.7% in the second half of 1989. This improvement of the female survival resulted in better fecundity. The emergence rate from the pupae was B95%. A successful technique for feeding colonies of G. morsitans and Glossina austeni in the absence of living hosts has been described.36 Flies are fed through membranes made of silicone rubber or of agar and Parafilm, overlying blood pools poured onto grooved glass plates. The diet employed is fresh pig blood, collected at slaughter, and aseptic procedures are adopted at every stage of diet preparation and presentation. The reproductive performance of these in vitro-fed colonies in terms of adult survival, fecundity, and offspring size is the same as that of colonies fed on living hosts, when compared over a long period of time. The fact that the technique is successful when used with a diet of pig blood, but it is not successful when used with cow blood prepared in the same way, suggests that the technique per se is adequate to elicit a normal feeding response from these tsetse species.

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Rabbit/Goat on animal rearing Adapting G. morsitans submorsitans to laboratory conditions using domestic animals (rabbits, goats) as hosts has been difficult requiring a 2-year period of adaptation.37

In vitro method(s) Contact—topical—adult The susceptibility of mature and immature male Glossina tachinoides to topically applied deltamethrin was tested on successive days of their hunger cycle.38 An artificial hunger cycle was created by starving both mature and immature flies for 1
5 days (maximum) following the last blood meal. Different batches of 40 mature male tsetse flies were topically treated with deltamethrin dissolved in 2-ethoxyethanol once on one of the successive days of their hunger cycle: TD 0, 1, 2, 3, 4, or 5. Flies were immobilized with N2 gas. Topical application of 0.5 μL 2-ethoxyethanol containing 0.08 ng of deltamethrin by means of an Arnold hand applicator was applied to the dorsal surface of the thorax. TD 0 being the day of the last blood meal, treatment on TD 0 was done immediately after the blood meal uptake. Groups of at least 54 immature male tsetse flies were also topically treated on one of the days of their hunger cycle (say 0, 1, 2, 3, or 4). The test was repeated for immature flies, starved during 1 or 4 days, but fed just before treatment. Batches of mature and immature male control flies were handled similarly as described earlier receiving the solvent 2-ethoxyethanol only. Mortality was recorded 24 and 48 h PT. During this period flies were not fed. Mortality among treated flies was corrected in comparison with the mortality observed in the control group. Contact—topical aerosol—adult A technique (MAP) enables single drops (size range 10
30 μm) of insecticide aerosols to be applied to individual insects.39 Monodisperse drops are generated in a wind tunnel, collected on silk threads, and transferred to the test insects. Test compounds are diluted in highly volatile IPA so that, when atomized by a vibrating orifice drop generator40 drops with final diameter of 10, 15, 20, or 25 μm and of field composition can be produced in the working section of a low speed (0.25
1.5 m/s) wind tunnel. Silk filaments drawn from second instar Spodoptera littoralis larvae were found to be entirely suitable. These filaments, diameter B2 μm, can be wound singly between the arms of Y-shaped frames, made from 1 mm diameter brass wire, and then set up in the wind tunnel to intercept the droplets. Exposure times of around 30 s, at an atomizer feed rate of 0.14 mL/min, allow the collection of adequate numbers of discrete droplets for bioassay with low probability of coalescence. The droplets adhere to the silk filaments as near spherical globules, and the frames can be transported to the insect dosing room in a sealed

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container. The tsetse flies are anaesthetized, individually, with CO2, held by means of suction from a vacuum line through a large bore, plain ended hypodermic needle applied to the dorsal thorax, and viewed through a low-power dissecting microscope (magnification 25 3 ), under low-intensity background illumination. A frame, bearing drops, was mounted in an adjustable clamp and additional light from a long-wave UV source was used to highlight the suspended drops positioned in the center of the microscope field of view [a small quantity (0.05%) of the fluorescent tracer Uvitex OB was incorporated in the sprayed formulation for this purpose]. The silk threads proved robust, and it was possible to position a drop over the chosen body site and bring about drop contact, then withdraw the fly leaving the thread intact, capillarity having effected complete drop transfer. Following dosing, flies were transferred to individual plastic cups and retained for 48 h at constant temperature and humidity to monitor survival. By applying 1, 2, 3, or more drops, it was possible to establish the dose mortality relation and estimate the median lethal dose. The technique has been used with tsetse flies and ultra-low volume formulations of synthetic pyrethroids. Contact—topical spray/cattle—adult Contact through the pulvilli of the feet of tsetse flies for a few seconds with a sprayed surface containing DDT, either fresh with solvent or as a residual deposit, was extremely lethal to the flies. Since cattle are a favored host of Glossina pallidipes, Glossina swynnertoni, G. morsitans, and G. austeni as well as some members of the fusca group, it may be feasible to topically treat cattle (75/m2) with a highly lethal compound to exterminate a population in 8
10 weeks. Therefore the objective was to develop a suitable preparation of DDT that can be placed on cattle and then using them to attract and so destroy the tsetse fly.41 Cardboard pieces sprayed with 5% DDT in kerosene were placed in Bruce boxes (a small gauze-covered box) containing 20
30 tsetse flies for 24 h was 100% effective. Flies were allowed to feed through cotton gauze (flies kept singly in tubes) on an ox sprayed at the rate of 0.5 g DDT (active isomer 60%)/ft2. All flies died within 24 h demonstrating that the flies feed through a treated surface without their feet contacting the compound and were killed through DDT contacting their proboscis, 0.3 g active isomer of DDT/ft2 being lethal. However, it was found that proboscis contact had no effect 72 h after spraying due to the residual crystals of DDT being rubbed off the treated animal. Hence, an emulsion formulation was developed consisting of 50 parts of a 5% DDT (active isomer 77%) dissolved in kerosene emulsified with 50 parts of fresh ox serum to which 0.5% sodium arsenite was added as a preservative to particles between 1 and 12 μm (mean 7 μm) in diameter and applied at the rate of 1 g/ft2. Tsetse flies were placed in contact with the surface of the treated animal without any intervening gauze for a set time, being held

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by the wings with a pair of forceps. Feet contact for 20 s caused paralysis within 15 min and death within 2 h. The residual activity in the laboratory lasted from 6 to 12 weeks. In the field the emulsion provided 79% efficacy during the first week, 75% during the second week, and 76% during the third week. Contact—vegetation—adult Various methods have been devised for exposing tsetse flies to leaf deposits of insecticides. A bioassay, based on the principles as that of Kernaghan and Johnson,42 was developed to be suitable for bioassays of insecticide residues on both leaves and the type of small twigs commonly used as resting sites for Glossina species43. Twig pieces of 3 cm long 3 1.3 cm diameter (area of B20 cm2) were cut from shrubs and small trees inside a G. pallidipes thicket habitat. The bark was removed from the twig, flattened by means of forceps on a piece of card measuring 9 3 6 cm, and secured by staples at each end. An apparatus was used for exposing insects to insecticide residues. The plunger (a rubber or cork stopper 2.6 cm in diameter, a glass or wooden rod 13 cm long and 7 mm in diameter, and a circular metal washer 3.3 cm in diameter with a central hole 7 mm in diameter) and retaining plate (a 6 3 6 cm2 piece of 1-mm thick alloy sheet and two rubber bands) of the apparatus are removed, and the treated (aerial spray of dieldrin concentrations) test material (leaf or bark mounted on the card) is placed between the platform and the retaining plate, with the residue-bearing surface facing the exposure chamber. The retaining plate is secured by means of two rubber bands so that the test material is held flat and firmly in place. A tsetse fly is placed in the glass container (a 12-cm length of glass tubing, with an outer diameter of 3 cm and a 2.7 cm bore, and a 6 3 6-cm2 piece of 8-mm thick plywood or plastic with a central circular hole 3.3 cm in diameter) and quickly confined by introducing the plunger a short distance. The apparatus is then rotated so that the retaining plate faces the light, thereby encouraging the fly to alight on the test surface at the bottom of the tube. The fly is then exposed to the treated test material and adult fly mortality determined. Contact—net or cloth—adult Deposits of deltamethrin suspension concentrate on terylene net and cotton cloth were bioassayed in Zimbabwe by exposing them to fed, female G. pallidipes, using a 45 s contact with cloth and a brief collision with the net.44 On textured yarn net the effective life of deposits produced from an immersion in 0.1% deltamethrin was longer than on flat yarn net, apparently because the textured net held more insecticide and the insecticide was lost more slowly. The addition of an absorber for UV light (0.1% 2-hydroxy-4methoxybenzophenone-5-sulfonic acid) did not significantly extend the

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effective life of deltamethrin deposits on cloth or net. Applying deltamethrin as a 0.6% suspension to cotton cloth produced mortalities of ,90% for 300 days. The applications of 0.8% deltamethrin on cotton cloth and textured net produced mortalities of ,70% for 12
16 months and 9 months, respectively, compared to 4
10 months and 5
7 months, respectively, for the applications of 0.1% deltamethrin. Chemical analyses indicated that the longer effective life of the 0.6%
0.8% deltamethrin was due to a higher initial amount of insecticide and a lower rate of loss. It is suggested that for controlling tsetse in southern Africa, all cloth targets sprayed with 0.6% deltamethrin will have an effective life of B1 year. Contact—fly cage/cattle—adult Males and females of G. palpalis gambiensis were derived from the tsetse colonies at CIRDES, Bobo-Dioulasso, Burkina Faso, where an independent large-scale in vitro rearing facility had provided sterile males for their subsequent releases in the agropastoral zone of Side´radougou, Burkina Faso.45 Considering the stress due to recapture of the released flies and a subsequent higher mortality, fly cages were attached to the bellies of bulls allowing the flies to feed on the first, second, and sixth day after emergence 170 days PT. Following their feeding, the tsetse flies were transferred into new cages. They were then maintained45 on an in vitro feeding system for 15 days in an insectary at 24 C
26 C and 80% RH and assessed for adult mortality. Feeding—artificial membrane—adult/larvae This method is proposed based on the FAO/IAEA SOP29 on rearing tsetse flies using a Trolley membrane feeding system. The objective of the assay is to determine the efficacy of test compounds ingested by adult tsetse flies. Pupae are placed in an emergence cage and maintained at 23 C
24 C and 75%
80% RH for 35
40 days. Emergence occurs and flies are collected using a separating tube and males and females identified. Female and male flies are kept in separate cages and fed using a Trolley membrane feeding system. At a heating mat or aluminum plate temperature of 35 C
37 C, lift two-thirds of the membrane at one corner of the tray, pour 100 mL of blood treated with a known concentration of adulticide or IGR or untreated blood as a control into the trays, and immediately replace the membrane. When the membrane surface temperature has returned to 35 C
37 C, manually put the female fly cage on the membrane. Allow the flies to feed for 10
15 min and return the fly cages to the holding room at 23 C
25 C and 75%
85% RH. The mortality of the female flies feed on treated and untreated blood at various time intervals postfeeding is determined to assess the efficacy of an adulticidal compound. If an IGR is tested, determine the number of pupae from female flies feed on treated and untreated blood on

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day 25 after larviposition. Transfer pupae to Petri dishes and place in an emergence cage. Determine the number of adult flies that emerge in B32 days at 23 C
24 C. Repellency Hornby and French46 successfully used solutions of pyrethroids as a tsetse repellent. The treatment deterred the tsetse flies from biting and so infecting (with trypanosomiasis) the treated baits. In fact, actual contact was needed with the treated surface before the flies were repelled, hence a “contact repellent.”

In vivo method(s) Parasiticide efficacy testing against tsetse flies on ruminants should be designed and conducted in accordance to WAAVP guidelines for evaluating the efficacy of ectoparasiticides against biting and nuisance flies on ruminants (see Appendix of Ref. [16]). Glossina transmits Trypanosoma evansi “surra” to horses. A traditional method for controlling biting insects on horses is the use of smoke released by slow fire, wherein the smoke repels the insects. However, the confinement and stress on the animals in the exposure area reduces their food intake, and the treatment only covers a limited protected area. Individual animal use of mosquito nets can be used to protect horses, but this is rather an expensive approach. It is possible to adapt fly-proof corals or stables for groups of animals. Insecticide impregnation of mosquito nets is an alternative integrated method of control, which can help reduce biting insects/vector populations.47 Stomoxys calcitrans Linnaeus, 1758—stable fly Biology and life cycle The stable fly, also known as the barn fly or dog fly, is a bloodsucking insect distributed worldwide. The adult flies are similar to the housefly in size (6
8 mm in length), but lighter in color with checkerboard ventral abdominal markings. The flies are gray with a greenish sheen body. The wings at rest are held wide apart, and the wing tips are distinctly iridescent. It is easily recognized by its large, piercing (bayonet-like) mouthparts, which project forward from the head. Both sexes of the stable fly are persistent blood feeders causing considerable irritation to host animals. They may feed twice a day, usually feeding on the lower aspects of the animal, especially near the hocks, oriented in a head upward position. After acquiring several blood meals over the course of 3
5 days, females will mate and develop a batch of eggs. Only one mating is required during a female lifetime. Male flies usually die after mating.

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The female stable fly will seek out areas, such as soiled animal bedding, spilled animal feeds, ground-piled silage, and compost piles, to deposit her eggs. Once an area is located, the female will crawl into the loose material, laying eggs sporadically as she moves. Females usually die after laying eggs. Each female will live for 20
30 days as an adult, laying between 500 and 600 eggs during her lifetime. Adult flies emerge after 6
20 days or more from puparia. The average stable fly life cycle is 28 days and can vary from 22 to 58 days, depending on weather conditions. Stable flies act as both biological (Habronema microstoma) and mechanical (T. evansi, Besnoitia spp.) vectors of disease. They are the intermediate host of nematodes including Setaria cervi. The eggs are B1 mm in length and are deposited in irregular masses of 1
50. Each egg-laying cycle produces 80
90 eggs, but the number can range up to 140. The eggs hatch in 24
48 h. Newly hatched larvae migrate downward into medium to avoid desiccation. The larvae develop for a period of 14
26 days. Larvae overwinter by slowing their development and do not diapause as do other species. Fully developed larvae crawl to drier areas on breeding medium where they pupate. Pupae are chestnut brown, elongate-ovoid and measure 5
7 mm in length. The pupal period varies with temperature and ranges from 6 to 26 days.

Rearing method(s) Numerous rearing methods for the stable fly have been developed.4,48
55 Stable flies preferred to oviposit on substrates with plant material and not on fresh manure.56 However, pupariation was maximal in fresh manure and the fresh pine shaving substrate when eggs are added to these substrates in the laboratory. Stable fly development is less successful on the substrates with soil. McGregor and Dreiss49 devised a method that has enabled rearing of an average of 1500 adults of S. calcitrans each day for 2 years in the laboratory. The larvae develop in jars measuring 7 3 9 in., half filled with a mixture of one part by volume standard CSMA medium (without the addition of yeast or malt) and five parts wood shavings, moistened with water so that one drop can be squeezed out of a handful. The eggs are stirred into this mixture, and the jar is covered with muslin and kept at 26.7 C. The eggs hatch in 24
36 h. A dry layer of shavings forms on the medium as the larvae feed, and they eventually pupate under it. If desired, it can be lifted off and the pupae removed, but it is usually more convenient to leave the pupae undisturbed and release newly emerged adults into a holding cage. Approximately 1000 flies/jar can be reared by this method, and the life cycle is completed in B3 weeks. Excessive moisture in the jars can be corrected by the addition

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of rolled oats. The flies feed through the screen of the cage on citrated beef blood in glass tubes with rubber suction bulbs, and they oviposit on black cloth wrapped round balls of damp absorbent cotton, and wetted with a few drops of 5% ammonia solution. A general stable fly rearing method by Berkebile et al.57 maintains immatures and adults at 23 6 2 C with variable RH (30%
50%) and photoperiod of 12L:12D hour. Adults were fed daily by placing an unscented feminine napkin, saturated with citrated bovine blood, on top of the screen cages. Eggs from 7- to 10-day-old adults were collected on moist black cotton cloth and rinsed into a dish. A modified pipette was used to transfer 1 mL of eggs (B8000) to a rearing medium consisting of wheat bran (500 g), fish meal (115 g), wood chips (200 g), and water (1.6 L). Immatures were allowed to develop in the medium for 12
14 days before pupae were harvested. Aggregations of pupae were spooned from around the perimeter of the plastic rearing pan (36.7 3 31.9 3 14.4 cm) where they tended to aggregate. The Gainesville house fly diet7 with the addition of pelleted peanut hulls 1:1 by volume becomes suitable for rearing S. calcitrans. This decreases the protein to B10% and increases the fiber to B30%. The rearing medium consists of 30% alfalfa meal, 50% wheat bran, and 20% corn meal plus pelleted peanut hulls (1:1). The rearing medium is measured into standard larval rearing trays (50 3 40 3 10 cm),58 moistened with water at a ratio of 5 parts diet:6 parts H2O by volume, and seeded with B30,000 stable fly eggs ,6 h old. The egg seeded diet is held at 26.7 C and 60% RH. The rearing medium had a 50% yield of pupae and a 44.9% yield of adult stable flies.

In vitro method(s) Contact—treated membrane—Adult Compounds (parasiticides) dissolved in IPA at various dilutions are spread, using the pipette tip, over the lens tissue that has been placed on top of a No. 2 pot (containing a single piece of circular filter paper) and held in place with an elastic band. One pot with only IPA spread over the lens tissue to serve as an untreated, solvent control. The treated lens tissues are left for 2 h for the IPA to flash off prior to adding the flies. Unfed flies are collected in 50 mL Falcon tubes and knocked down on ice for B30 min. Ten flies are added/pot. A watch glass filled with citrated bovine blood sealed with Parafilm is placed over the lens tissue. The flies feed through the lens tissue coming into contact with the compound. Contact—treated cattle—spot and cage test—adult The spot test technique was developed to permit an initial adult fly efficacy evaluation with 250 mg or less of a compound in a single area application to determine the toxicity, knockdown, or repellency of the compound.59 This method consists in applying a test compound or formulation to small areas

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on a steer or cow. Areas 6 in in diameter are marked with black dye on the upper half of the body in such a manner that no area is directly below another and each is on a fairly smooth area. On a large animal, six of these areas can be grouped on a side, and on smaller animals, five areas can be adequately spaced. The hair within the areas is clipped to a 0.25
0.5 in. length, so the flies can reach the skin to feed. The test compound is dissolved in acetone (usually 5% solution) or another suitable solvent, and 5 mL (adequate volume to cover area without run off) is applied with a De Vilbliss atomizer using compressed air as a propellant. The spray is confined to each area with a metal cone. Cages are made from Mason jar lids by soldering a piece of screen wire in place of the removable top. The cages are 3.5 in. in diameter, 0.5 in. deep at the edge, and 0.75 in. at the center. Each cage is supplied with a metal tray. The cage is held to the tray by a flanged edge, which is slid into the rolled edge of the tray. Stable flies 3
5-days-old and unfed for 18 h are anesthetized with CO2, and 25 flies are placed in each cage. When the flies recover, the cage is slipped off the tray onto the treated area and the flies are exposed for 20 min (at least 15 min is required for 90% of the flies to feed to engorgement). The cage is held in position with two pairs of clips, joined with a heavy rubber band, which are anchored to hair treated with rubberized branding cement at four points around the treated area. The rubber bands, under tension, hold the cages in position regardless of the animal’s movement. After exposure the flies are moved to a holding room at 22.2 C and 75% RH. Toxicity is measured by the percent mortality of the flies 24 h after exposure and knockdown by the percent of flies unable to fly or walk immediately after they are removed from the test area. The cage test can be used to confirm a compound active in the spot test. A yearling is sprayed with 3.8 L of insecticide at a concentration indicated by the spot test. On different days after treatment the animal is placed in a screened cage and exposed to 100 unfed stable flies. Two hours after release, all flies remaining alive are collected, the number that feed are recorded, and the flies are held 24 h for mortality counts. The difference between the number released and the number collected is used as a measure of knockdown. This difference plus the number dead after 24 h gives the percent control. Contact—treated animal—calf patch test—adult On TD 1, four heifer calves (175
225 kg) are tagged with individually numbered ear tags (one/ear), vaccinated per study site requirements, and not treated with any ectoparasiticide product for at least 60 days prior to treatment. Calves are weighed and housed in individual stanchions or chutes. Each of the four calves will have a maximum of 4 areas designated on either side (L 5 left, R 5 right) of the spine as treatment sites for a total of 32 treatment sites using the designation: Calf ID 1—Patch ID 1
4 L and 1
4 R; Calf ID

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2—Patch ID 1
4 L and 1
4 R; Calf ID 3—Patch ID 1
4 L and 1
4 R; Calf ID 4—Patch ID 1
4 L and 1
4 R. The patch areas are clipped for compound application. The calf will then have a muslin sleeves attached, to delineate areas using a marker pen. On TD 0 the application of the test compound will be made using a measured volume of 640 μL to deliver fixed amounts of compound to each patch according to the study design. Dose calculations: If a test compound is at 0.1% a.i. (1 mg/mL 5 1 μg/μL), then at 10 g/cm2 the required amount is equivalent to 10 L/cm2. The feeding area is dictated by the radius of the area treated, which is B4.5 cm. The area of treatment equates to B64 cm2, so at 10 L/cm2 a volume of 640 L is required to deliver doses of 10, 30, and 90 μg/cm2 when delivered at concentrations of 0.10%, 0.30%, and 0.90% w/v, respectively. Likewise, a volume of 640 L is required to deliver doses of 5, 15, and 45 μg/cm2 when delivered at concentrations of 0.05%, 0.15%, and 0.45% w/v, respectively. The control sites will receive 640 L of IPA. Test compounds will be dissolved in IPA. Applications of the test doses will be made using a measured volume of 640 μL to deliver fixed amounts of compound to each patch according to the randomization treatment site. The formulations will be applied using a handheld pipette so that the total dose is evenly distributed across the delineated area on the calf. Treatment drying time, a minimum of 1 h, will be allowed before the feeding of flies. Unfed Stomoxys adult flies (B30) in cages will be allowed to feed on treated areas for 45 6 5 min. Two control cages of Stomoxys adult flies will also be collected. These control cages will be kept at a constant temperature of B20 C
22 C and fed bovine blood on a dental plug. All Stomoxys will have been starved of a blood meal 18
24 h prior to feeding. After feeding the caged flies will be removed from the calves, kept at a constant temperature of B20 C
22 C, and monitored at 1, 4, and 24 h PT for knockdown, toxicant effects, and fly mortality. If no knockdown effect upon the flies is observed from B24 h after exposure the compound will be deemed inactive at that dose. Fly knockdown is defined as the inability of the fly to hold onto and/or maintain position within the test cage. Mortality is defined by the absence or lack of significant movement of the body and mouthparts with the exception of characteristic twitching of the legs and mouthparts. Following the S. calcitrans feeding, B50 newly emerged, unfed H. irritans adult flies in cages can be allowed to feed on the treated areas. Refer to the horn fly in vitro testing section for details. This additional procedure maximizes the use of the treated animals. Stable fly count reduction figures (mortality and knockdown) for treated groups will be generated using geometric group mean counts and expressed as percentage efficacy relative to control group mean count. For each treatment group the minimum lethal dose that kills 100% of the flies (MLD100) will be determined by probit analysis based on the percentage efficacy by treatment concentration.

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Feeding—dental roll/blood—adult The objective of this bioassay is to determine a lethal concentration of test compounds in bovine blood against adult stable flies. Test compounds are diluted with DMSO at 3 mg/mL (equivalent to 10 μg/cm2) and sonicated to have the chemical dissolve into solution, if necessary. Serial dilutions are prepared in μg/mL concentrations, for example, 0.03, 0.1, 0.3, 1.0, and 3.0 μg/mL. Fipronil can be used as a positive control. Unfed flies are collected in 50 mL Falcon tubes. For ease of collection, place a light on the Stomoxys cage. The flies will be attached to the light and are easier to put into the tube. The flies are anesthetized or immobilized on ice for B30 min. Filter paper (15.5 cm) is placed onto each labeled plastic container. Ten anesthetized flies are added per pot (No. 3 clear plastic containers) on top of the filter paper. One piece of gauze (B4 3 4 cm) is taken, opened, placed on top of the plastic container, and secured over the gauze under the container’s rim. Then 5 μL of each compound dilution in DMSO is added to 2 mL of citrated bovine blood and gently mixed. A dental roll (No. 2 dental plug cut in half) is place in the mixture of compound and blood. The saturated dental roll (1.4 mL to saturate) is placed on top of the gauze. In addition to the test dental roll, a dental roll soaked in sucrose/distilled water solution is also place on top of the container to sustain the flies until the 24 h count. The flies recover from the anesthesia and feed on the dental roll containing the treated blood. The test is conducted at RT. Flies are examined for knockdown and mortality at 2 and 24 h. The number of dead flies out of 10 is recorded. No more than 10% mortality in the controls is acceptable for quality control. After the 24 h reading the containers of flies are placed in a freezer for 24 h and then disposed of as biological waste. Note: In some cases the test compound may be removed from the blood by a solid phase extraction mechanism where a significant percentage of the compound is absorbed to the plug material, thus yielding a false reading. The systemic activity of the test compound may need to be determined in a different feeding assay. Repellency; spot test and cage test—cattle—adult The spot test and cage test consists in applying materials to small areas on cattle and confining stable flies on these areas at various intervals after treatment.59 See the test description earlier for the details of the technique. Repellency is measured by subtracting the percent of flies that fed from 100%. Electroantennograms The method used by Balacchino et al.60 was modified from Jeanbourquin and Guerin61 to conduct EAG recordings from antennae of S. calcitrans.

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Recordings were made using electrolyte-filled (0.1 M KCl) glass capillary electrodes (diameter 1.5 mm, 40 mm L), with Ag/AgCl wire (diameter 0.5 mm, 20 mm L) making contact with the recording apparatus. The antenna was maintained in a humidified charcoal-filtered airstream delivered at 14.6 mL/s through a metal tube. Aliquots of pure lemongrass oil (from Cymbopogon citratus, citral B75%) were prepared using hexane at 0.1, 0.01, 0.001, 0.0001 mg/μL. Tested solutions (10 μL) were deposited on a strip of filter paper (20 3 5 mm) placed in a glass Pasteur pipette. The solvent was allowed to evaporate for 15 min before first use. The tip of the pipette was connected to the metal tube, and the test stimulus was delivered to the antenna using an air pulse (20 mL/s for 0.6 s). Stimuli were released successively in random order at 90-s intervals to avoid receptor saturation. Octanol was used as a positive control, and hexane was used as a negative control. Differences in EAG responses were evaluated using a Wilcoxon signed-rank test. The video records of fly movement were analyzed using EthoVison XT.62 The cage was defined as an arena (30 3 15 cm) divided into three zones (each 10 3 15 cm): untreated, intermediate, and treated. The movement was recorded at 25 video frames/s, and the fly was tracked by dynamic subtraction. In this method the program compares each sampled image with a reference image that is updated regularly. Image processing algorithms are applied to detect the fly against the background and to extract relevant image features. During data acquisition, EthoVision displays the live video image, tracking statistics (elapsed time, number of samples), and the x, y coordinates of the fly.62 Several parameters were calculated: the distance moved (cm), the total time spent in each zone (s), the time spent in movement (s), and the mean velocity (cm/s). “Moving” and “not moving” were defined with thresholds at 1 and 0.9 cm/s. A comparison between males and females was made with the nonparametric Mann
Whitney test for independent samples. Comparisons of flight parameters between the treated zone and the untreated zone were made with the nonparametric Wilcoxon signed-rank test for two samples of univariate data. Feeding—blood—adult A choice test bioassay is used for testing the feeding of compounds in blood against adult flies in access to two blood sources, one of which was treated with lemongrass oil. Citrated bovine blood (1.5 mL), previously heated at 45 C, was placed on two sanitary pads (diameter 4 cm) from which the outer layer was removed. The outer layer of one pad was impregnated with 100 μL of lemongrass oil solution at 0.1 mg/μL, and the other outer layer with 100 μL of hexane. When the solvent had evaporated, each outer layer was repositioned on top of one of the blood-soaked sanitary pads that were placed just under the cage floor, 20 cm apart. Fly movement was recorded

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using a digital video camera recorder set 1 m above the center of the cage. The behavior of the fly was then recorded during a 10-min period. Flies (4
6) were tested each day with the behavior of 24 flies included in the study. The room was ventilated for at least 30 min between each test, and a new screen cage was used for successive flies. The positions of the pad treated with lemongrass oil and the untreated pad were inversed each time. The cages were cleaned every day by soaking them in a 2% solution of Decon 90 for 12 h. Contact—single-cage olfactometer—adult A single-cage, dual-port olfactometer has been used to assess the spatial repellency of catnip oil, its ingredient compounds, and DEET against stable flies63. This system was constructed from clear glass (4 mm), which was modified from a triple-cage olfactometer described in Posey et al.64 The whole dimensions of the olfactometer were 96 cm long 3 50 cm wide 3 25 cm high and placed on top of a metal tool table painted white. The front of the test cage was connected with a pair of glass ports (30 cm long and 10 cm in diameter) that were set at 10 cm apart. The rear of the test cage was connected with a similar sized glass port covered with a 1.0 mm mesh window screen in frame that allowed the flies to acclimatize to the air being used in the tests. Two small table fans were set in front of the dual ports to generate wind speeds measured at 0.35
0.4 m/s, which were regulated by separate voltage regulators. All experiments were conducted at B27 C and 55% RH. Stable flies used for testing were B3
4-days-old and were starved for 24 h prior to testing. The flies were released individually. Each fly was given 3 min to respond, and their presence in either repellent/ attractant treated or control ports (at 10 cm inside the port) was recorded. Normally, one set of six tests (six treatments) was performed in 1 day. Within each set of tests the order of ports with either attractants/repellents or the controls was randomized. All three ports were cleaned first with acetone and then hexane before new tests began. Each experiment was replicated 3
4 times with 20 flies, and different (new) flies were used in each replication.

In vivo method(s) Parasiticide efficacy testing against stable flies on ruminants should be designed and conducted in accordance with the WAAVP guidelines for evaluating the efficacy of ectoparasiticides against biting and nuisance flies on ruminants (see Appendix A of Ref. [16]). Topical parasiticide tests The stable fly is the most common blood feeding muscoid fly pest of horses and an important intermediate host of H. microstoma.17 They usually feed on

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the lower legs, especially the front pair. The painful bite of the fly on the legs causes horses to stomp their feet. Rub-on treatments of 0.5%, 1.0%, and 1.5% permethrin emulsions provided 7
10-day control of S. calcitrans on B100 horses maintained at the Clark Air Base stables in the Philippines.65 Emulsions of 1.0% permethrin rubbed on or sprayed on half of the herd also gave 7
10-day control during the late rainy season and throughout the dry season on central Luzon. Longer control was obtained during the driest periods. Alsynite panels treated with 3.2 g a.i./cm2 of permethrin wettable powder reduced the number of stable flies/horse by B50% over a 1-week period during the dry season. Permethrin-impregnated net blankets (0.28 mg a.i/cm2) reduced stable fly populations B60% on 30 separately pastured horses wearing the net blankets. Net blankets treated with 0.56 mg/cm2 permethrin retained their insecticidal effectiveness for .7 weeks under early rainy season field conditions. Physical damage to the net blankets and difficulties in coordinating net blanket use were the primary factors limiting their usefulness for biting fly control. Tabanus megalops (Diptera: Tabanidae) and other horse flies were controlled for 3
6 days following rub-on or spray-on treatments of 1.0% permethrin to 1/2 of the horses in the experimental group. Repellent test Six adult horses were individually penned in 5 3 5 m dry lots for 2 h each day for 5 consecutive weekdays over 6 weeks; all horses were fitted with ear and face fly masks.66 In a Latin square design, horses were randomly assigned to receive one of six treatments each week (five commercial repellent products including a pyrethrin spray, citronella/vinegar spray, permethrin spray, leggings (Shoofly Leggings), leg band with citronella/phenylethyl propionate and a control with no repellent). Commercial products were applied according to manufacturers’ recommendations. At the end of each 5 day period, horses were bathed, left without protectants over the weekend, and then rotated to their next treatment. Each day, horses were observed immediately following protectant application. Stable flies on legs and bodies of horses were counted at 0, 20, 60, and 120 min, and fly aversion behaviors were counted by 6 trained observers assigned to the individual horses. Behaviors were counted in 30 min periods and included 5 min to count tail swishes; 20 min for front and back limb hoof stomps and head motions where the head was brought to a limb or flank (head backs), and 5 min for shoulder twitches. Observations were repeated for four 30 min periods (total of 120 min). Results were compared using a mixed model ANOVA with horses and week as random effects. Stable fly counts averaged 3.8/horse during the study; no other fly species were present. Average counts of fly avoidance behaviors were not different between the first and second hour, indicating efficacy did not change within 2 h of application. Treatments did

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affect fly avoidance behaviors (P , .0001) with leggings the most effective at reducing these behaviors in horses.

Horn fly—Haematobia irritans Linnaeus, 1758; buffalo fly—Haematobia irritans exigua de Meijere, 1906 Biology and life cycle The horn fly is an obligate ectoparasite of cattle and is a well-established pest of cattle through many tropical and temperate areas of the Northern and Southern Hemisphere. A subspecies, Haematobia irritans exigua, known as the buffalo fly, occurs in Australia and the Pacific region. The buffalo fly and horn fly have very similar biology and life cycles. Adult horn flies appear similar to stable flies in wing venation and about a third or half the size of the housefly measuring 3.5
5 mm long. The dorsal surface of the thorax is silvery-gray, while the lateral portions of the thorax are dark with two well-defined stripes. Horn flies typically have dark reddish-brown eyes. The life cycle of the horn fly from egg to adult varies from 8 to 12 days in hot, humid weather to as long as 20
45 days during the spring and fall seasons. Each female fly lays 14
17 eggs at a time on freshly excreted cattle feces. A female horn fly is capable of laying 400
500 eggs during its lifetime. Both male and female flies have slender, black, piercing mouthparts that project forward from the bottom of the head. Average meal size is 1.5 mg or 10 μL of blood/feeding and each fly can take 24
38 blood meals/day67 for 10
25 min at a time. Mating occurs on the host 1
2 days after emergence and egg laying 1
5 days later. Both sexes remain on or in close association with the host, day and night, leaving only to pass to another animal or when the female oviposits while the animal defecates.68 The flies aggregate around the horn region and on the shoulders and backs of cattle, each fly oriented with its head in the same direction as the tips of the hair and their wings at a 45-degree angle to their body. Adult flies often move to the belly of the host on extremely hot days or during rain storms. The emerging adult horn fly is a strong flier, capable of flying 5
7 miles to find a host. But, the adults remain on a suitable host or others in the herd for life, with the female only leaving to lay eggs. Horn flies serve as an intermediate host and mechanical vectors of Stephanofilaria stilesi. Horn flies lay their eggs in fresh cow manure with the female fly known to lay her eggs in the feces (,2 min old or within 10 min of dropping) before the cow has completed defecation. The long-ovoid eggs measure 1.3
1.5 mm in length, vary in color from dark or reddish brown to yellow brown, and have a tapering longitudinal groove on one side. Eggs hatch in one day or less, depending on temperature and humidity. Newly hatched larvae crawl within cattle feces and begin feeding on bacteria and decomposed products of the resident bacteria. They undergo two molts reaching a third instar in 3
12 days.

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Pupation occurs 3
14 days on the lower part of the droppings or in the ground immediately beneath the fecal pad. The elongate-ovoid pupae are brown in color, measuring 3
4 mm long, and possess the spiracle of the third instar. After 2
8 days the adult fly emerges from the puparium and begins feeding on cattle 2
3 h after emergence.

Rearing method(s) Laboratory colony Adult horn flies in cages (21.5 cm2 wire frames with plastic-screen covers) are fed an artificial diet of beef blood containing acid citrate dextrose, nystatin, and chloramphenicol. Alternatively, kanamycin sulfate 1000 mg and nystatin 500,000 U dissolved in 100 mL water and 7.5 g sodium citrate dissolves in 50 mL water are added to a 3-L IV bag prior to bleeding a cow using a 14 gauge needle. The bovine blood is refrigerated for storage and aliquots of bovine blood are allowed to warm to RT prior to feeding horn flies. The blood diet is offered to flies twice a day on cotton pads, feminine napkins (70 mL of blood to saturate pad), or glass Petri dish lids covered with Parafilm. Egg collection is started 4 days after adult flies emerge from pupae. Eggs are collected on cotton or gauze pads covered with wet cheesecloth or a wipe-all towel placed in a collecting tray. The pads should be thoroughly saturated with water but pour off standing water. Be sure the corners of the towels have been folded in. The egg collecting tray is placed under the adult cage that is supported on the collecting tray by two fiber boards. The horn fly eggs will fall through the wire cage onto the moistened toweling. To collect the eggs, remove the collecting tray from under the cage. Rinse the contents of the eggs with water into a No. 100 mesh (0.149 mm) sieve. Discard the old toweling and replace it with a new one in a clean collecting tray. Place the tray under the fly cage to continue collecting eggs. Egg collections from multiple trays can be pooled in the sieve. Fresh cattle feces (1200 g) are weighed into a quart bucket. Then the eggs are rinsed into the feces using as little water as possible. Cheesecloth is placed over the bucket that has been labeled with the date. The bucket is placed in a media room on a shelf and incubated at 26 C
28 C and 60%
70% RH. Pupae develop in B7 days after seeding the manure with eggs. The manure bucket with pupae is dumped into a flotation tub of warm water and the manure is gently broken up, especially any floating clumps. The manure is held in the flotation tank for 30
45 min to allow the chaff to become saturated and sink. The pupae float to the surface and are skimmed off the surface of the water. Pupae are placed in a No. 20 mesh (0.841 mm) sieve and rinsed with water. The pupae are air-dried 20
30 min in the sieve and are checked while drying to break up any chaff that might be sticking together. A pupae drier can be constructed whereby the sieve of pupae fits

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onto a platform through which warm air is passed. The collected pupae are placed in a cup in a fly emergence cage incubated at 26 C
28 C and 60%
70% RH. The average cycle from egg to adult is 12 days and from egg to egg is 17 days.69
72

On-animal reared colony A bull calf (B6
8 months, B200
225 kg) is confined in a stanchion or stall on an elevated steel platform (1.2 3 1.8 m). The platform is 30 cm above the floor at the rear and contains a 45 cm section of grating 30 cm from the rear edge. The gratings cover a gutter positioned to catch urine. Heifers should not be used as urine prevents horn fly eggs from hatching. The platform is padded with rubber matting 1
2 cm thick for the comfort and safety of the animal. The 1 m distance between the side rails of each stall allows the animal to lie down at will. Alfalfa pellets (3/4 bucket twice a day) and water are provided ad libitum. When the animal is correctly positioned on the platform, manure is deposited directly into a collecting tray (60 3 90 cm plastic tub). The room housing the animal is air conditioned or heated to maintain 26 C
28 C and 60%
70% RH. Newly emerged adult horn flies (200 adult flies of Bequal sex) are released in the animal-holding room and feed freely on the animal. The female flies leave the animal to deposit eggs in the manure as the animal defecates. Once the manure collecting trays are full (a bull this size produces B4 kg each day), they are pulled to the outer areas in the animal room. If horn flies are needed for screening purposes, the eggs are allowed to hatch, develop through the larval stages, pupate, and the emerged adults can then go directly onto the animal. Bulls are rotated in and out of the system to protect their health. Some bulls can remain in the stalls more than 3 months without noticeable detriment or discomfort, but 45
60 days is a typical holding period. Horn flies are easily vacuumed from the shoulders and back to collect adult flies for testing. The flies can be immobilized in a cold room and easily sexed and counted. Alternatively, the manure trays can be removed from the animal rearing room and held for 24 h to allow the majority of eggs to hatch. Then the manure tray is allowed to incubate and monitored for the development of pupae. Or, if the animal was fed on hay rather than alfalfa pellets, the manure can be mixed in a 0.2 m3 concrete mixer with rearing medium {2 parts feces, 1 part dry mix [parts by weight: 40 ground bagasse, 8 whole wheat flour, 6 fish meal (65% protein), 1 sodium bicarbonate], and 2.6 parts water (78.3% moisture) at a mix ratio of 10:3:10} for 2 min and placed in rearing trays (corrugated boxes 30 3 46 3 13 cm deep) on racks and incubated at 26 C
28 C and 60%
70% RH. Pupae are extracted from the manure or rearing medium by flotation.

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Alternatively, adult flies can be collected as they emerge from the rearing medium whereby flies emerge in a warm (26 C
28 C) dark chamber (1.2 3 1.2 m), are attracted into a cooler (0.5 C
2.0 C) chamber (1.2 3 1.2 m) by light (40 W fluorescent) and immediately immobilized by the cold environment.73,74

In vitro method(s) Contact—cloth—adult Using the technique described by Schmidt et al. (1985),75 unbleached 10 cm2 muslin cloths are laid on aluminum foil and treated with 1 mL of an acetone solution of each concentration of organophosphate (diazinon) with an automatic pipette. When the plunger is pressed slowly and the pipette moved from side to side or in a circular motion above the cloth, the liquid flows uniformly over the cloth. Cloths are treated with a series of dilutions, which includes a control (acetone only) and several concentrations of active ingredient, treating three cloths/concentration. The cloths are allowed to dry under the fume hood until the acetone has evaporated. Cloths are then wrapped in aluminum foil and held overnight. In the laboratory, horn flies (3
4-days old) are immobilized with CO2 and held on a refrigerated chilling table for counting. Flies are placed in test containers with an aspirator. For exposure to treatment, 25 flies are placed in a 200 cm3 clear plastic specimen cup, the cup is then covered with the treated cloth that is held in place with the use of a cardboard lid with a 1 in. hole cut in the center. A 2 cm2 piece of cotton is soaked with citrated bovine blood or Gatorade is placed on a second patch of cloth (B2.5 cm2) on top of the treated cloth. The extra cloth prevents the blood or Gatorade from saturating the treated cloth and, perhaps, diluting the treatment concentration. It also allows the fly to feed through the cloth as it would feed through the skin of a treated host. Flies are held at 24 C
27 C and a minimum of 50% RH and examined for knockdown and mortality at 2, 18, and 22 h. Both the number of alive and dead flies is recorded. Lethal concentration calculations are then determined by comparing the treated and untreated fly mortalities. Contact—filter paper—adult The objective of this bioassay is to determine the concentration of a parasiticide residue on a filter to provide the knockdown and mortality of adult horn flies. Filter papers (9 cm Whatman No. 1) placed on aluminum foil or in the lids of a Petri dish cover are treated with serial dilutions of test compound (0.0108
4.3 μg/cm2) dissolved in acetone and allowed to dry at RT in a fume hood. Control (acetone only) filter papers and three filter papers/concentration are prepared. Twenty-five horn flies (3
5-days old, anesthetized with CO2) are placed in the bottom dish cover and the treated lid cover is applied. Once the flies revive, the Petri dish is inverted

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so the flies are in contact with the treated filter paper. The dishes are incubated at 26.7 C
27.5 C and 60%
75% RH. Knockdown and mortality are recorded every 15 min for a 2 h period or daily up to 3 days of exposure. This bioassay was originally developed by Sheppard and Hinkle,76 which has been accepted as the standardized test to determine both LD50 and LD90 probits and the discriminating dose. The filter paper bioassay is not suitable for organophosphate determinations. This assay is suitable for resistance testing. Contact—glass test tube—adult This bioassay is based on the technique developed by Cilek and Knapp.77 About 20 mL glass scintillation vials are treated with 250 μL of an acetone solution of each concentration of organophosphate or pyrethroid with an automatic pipette. The vials are rolled under the hood until the acetone evaporates, allowing the chemical to coat the sides and bottom of the vials. They are allowed to sit uncovered for 1 h and then tightly capped and stored in cool temperature, out of direct light, overnight. Flies are put into the glass vails with an aspirator and tightly capped. Mortalities are recorded at 1 and 2 h intervals. Dose mortalities are expressed in μg/cm2. Contact—glass Petri dish—adult The objective of this bioassay is to determine the concentration of a parasiticide residue on a glass surface to provide the knockdown and mortality of adult horn flies. Fenvalerate concentration levels have been established on the surface of glass Petri dish lids. While other investigators have developed filter paper methods for insecticide evaluations, the full potential activity of the compound on or in treated filter paper might not be available to the target pest. For example, diazinon treated filter papers must be aged prior to testing in order for the compound to migrate out of the paper matrix. Also, solid compounds, such as tetrachlorvinphos, remain inside the filter paper as crystals and are not available to the insect. For susceptible horn flies, test dilutions recommended for fenvalerate in acetone are 2.08, 1.04, 0.26, 0.065, and 0.016 μg/mL. About 3 mL of the solution is pipetted into the lid of a glass Petri dish with a surface area of 62.2 cm2 (verify as some lids can be as large as 72.4 cm2). This will yield the following residue concentrations: 1021, 5 3 1022, 1.25 3 1022, 3.1 3 1023, and 7.7 3 1024 μg fenvalerate/cm2 or 6.24, 3.12, 1.25, 0.2, and 0.05 ppm, respectively. Regression analysis of the mortality results should yield an LD90 of B0.15 ppm. The test dilutions are prepared in acetone and all active ingredients are corrected for purity. Duplicate Petri dish lids (rim up) are prepared for each dilution. Control plates will be prepared with 3 mL of acetone/dish. The lids are placed on a level surface in a fume hood.

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A 3.0 mL aliquot of each dilution is pipetted into the duplicate Petri dishes and the acetone is allowed to evaporate for 1
2 days at RT. About 20
25 horn flies (anesthetized with CO2) are placed in the bottom dish cover and the treated lid cover is applied. Flies are allowed to recover for 5
10 min. A mortality check will be performed to assess the effect of handling and immobilization. The dish will then be inverted so that the resting flies will be in contact with the treated surface. The dishes are incubated at 26.7 C
27.5 C and 60%
75% RH. Knockdown and mortality are recorded at 2 h and daily up to 3 days of exposure. The glass treated bioassay is suitable for organophosphate determinations with diazinon as the primary test compound. The correction factor from glass to muslin is 250 3 (i.e., 250 times as much toxicant). This assay is suitable for resistance testing. For pyrethroid-resistant horn flies, test dilutions recommended for fenvalerate in acetone are: 20.8, 10.4, 4.16, 2.08, and 1.04 μg/mL. About 3 mL of each dilution pipetted into a Petri dish lid will yield 1.0, 5 3 1021, 2 3 1021, 1 3 1021, and 5 3 1022 μg/cm2 or 62.4, 31.2, 12.5, 6.2 and 3.12 ppm, respectively. Expect an LD90 of B15.5 ppm depending on the resistance level of the flies. Based on the LD90 values presented earlier, the R/S value would be 103 3 for the pyrethroidresistant horn flies. Using this assay, diazinon was found to exhibit selective activity against pyrethroid-resistant horn flies yielding R/S values of 0.6 3 in the laboratory.78 The selective activity was confirmed in field studies78,79 leading to the development of the Patriot (40% diazinon) cattle ear tag. Similarly, endosulfan exhibited an R/S value of 0.6 3 and was eventually developed as the Avenger (30% endosulfan) cattle ear tag. These methods could be used to detect resistance to some of the newer compounds in ear tags and adapted in a discovery program to identify compounds active against horn flies resistant to the ear tag compounds. Contact—treated hair—adult To correlate in vitro testing with in vivo activity, hair clipping studies from cattle tagged with fenvalerate tags for B4.5 months were analyzed showing insecticide levels of 300 ppm from inside the tagged ear, 40 ppm from outside the tagged ear, 8 ppm from the jaw (cheek area), and 3 ppm from behind the shoulder. In vivo release rate studies of fenvalerate ear tags showed 0.7
1.4 ppm/day. Based on in vitro evaluations, 0.26 ppm of fenvalerate was 100% lethal to susceptible horn flies. Thus the in vitro concentration is B12 times less than the in vivo concentration on the shoulder area. Contact—treated cattle—calf patch test—adult This assay is identical to the calf patch test for S. calcitrans earlier. Newly emerged, unfed H. irritans adult flies (B50) in cages are allowed to feed on treated areas for 34 h ( 6 30 min). Two control cages of

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H. irritans adult flies will also be collected. These control cages will be kept at a constant temperature of B20 C
22 C and fed bovine blood on a dental plug. All H. irritans flies will have been starved of a blood meal 18
24 h prior to feeding. After feeding the caged flies will be removed from the calves, kept at a constant temperature of B20 C
22 C and monitored at 2, 6, and 24 h PT for knockdown, toxicant effects and fly mortality. If no knockdown effect upon the flies is observed from B24 h after exposure, the compound will be deemed inactive at that dose. Fly knockdown is defined as the inability of the fly to hold onto and/or maintain position within the test cage. Mortality is defined by the absence or lack of significant movement of the body and mouthparts with the exception of characteristic twitching of the legs and mouthparts. Horn fly count reduction figures (mortality and knockdown) for treated groups are generated using geometric group mean counts and expressed as percentage efficacy relative to control group mean count. For each treatment group the minimum lethal dose that kills 100% of the flies (MLD100) is determined by probit analysis based on the % efficacy by treatment concentration. Feeding—artificial membrane—adult Citrated bovine blood is treated with concentrations of test compounds and 1.0 mL is added to a plastic cell culture dish (60 3 15 mm, 5 mL) and covered with Parafilm membrane. The dish is placed in a modified wire screen cage along with 25 adult horn flies. Flies are held at 24 C
27 C and a minimum of 50% RH and examined for knockdown and mortality at 2, 18, and 22 h. Both the number of alive and dead flies is recorded. Lethal concentration calculations are then determined by comparing the treated and untreated fly mortalities. Repellency No specific repellency tests were found, although stable fly bioassays would seem appropriate.

In vivo method(s) Parasiticide efficacy testing against horn flies on ruminants should be designed and conducted in accordance with the WAAVP guidelines for evaluating the efficacy of ectoparasiticides against biting and nuisance flies on ruminants (see Appendix of Ref. [16]). A screening study design on cattle to evaluate the formulations of pourons or ear tags against susceptible or resistant horn flies prior to advancing a formulation to field evaluation consists of selecting heifers, 250
300 kg, individually identified with numbered ear tags (one/ear), vaccinated per site

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requirements, and not have been treated with an ectoparasiticide for at least 45 days prior to treatment. Depending on the number of formulations to be evaluated (this example is described for five formulations), at least 65 cattle will have been acclimated for at least 2 weeks prior to treatment. Horn fly counts are conducted on TD 7 and 4 and the 60 animals with the highest counts selected for study. The 60 animals are randomly allocated to 6 groups of 10 animals. The groups of 10 animals each are randomly assigned to separate pastures and to the six treatment groups. One group is assigned as the control group and is treated with a chemical vehicle, if the study is for a pour-on trial, or a placebo ear tag. All groups are treated on TD 0. Horn fly counts are conducted on TD 1, 4, 7 and weekly thereafter until week 4 (TD 28). If persistent efficacy of $ 90% in any treatment group at 4 weeks, the study may be extended accordingly with weekly counts until the efficacy drops below 90% in all treatment groups. Counts may be postponed 48 h in the case of inclement weather. Horn fly counts will be made visually on all animals by an experienced observer using either binoculars or a spotting scope or by closely approaching the animal. Counts will be made on one side of an animal in the area from the top of the back to the belly and from the point of the shoulder to the back leg when the flies are most active (between 8:00 and 14:00). The whole animal will be counted if counts are low. Fly counts will be made within a specific sized area when populations increase so to be unable to determine actual fly numbers. An estimate of the total number of flies present for one side of the animal is determined based on that fraction of the body area counted. Horn fly counts are log transformed and the percentage efficacy using geometric mean treatment group counts is determined relative to control group mean counts. The horn fly is a facultative pest of horses, generally around pastured cattle. Five insecticides were applied as sprays to horses in large-cage tests.80 Crotoxyphos, coumaphos, and carbaryl gave satisfactory control of horn flies for 2
3 weeks, while malathion control of horn flies (90% mortality) for 7 days and pyrethrins 1 piperonyl butoxide controlled horn flies for 10 days.

Melophagus ovinus Linnaeus, 1758—Sheep Ked Biology and life cycle The sheep ked is distributed worldwide with domestic sheep and can occasionally infest goats. Sheep keds are biting flies that feed on blood and are permanent ectoparasites on their host. They transmit Trypanosoma melophagium. Adults are wingless (even halters are absent), brownish-red in color, leathery, dorsoventrally flattened, unsegmented, and measure 5
8 mm long, superficially resembling ticks. The saclike abdomen is soft and leathery

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covered with short spiny hairs. They possess strong claws on their feet, which aid in clinging to wool and hair. The female sheep ked produces only one offspring at a time. Female keds become sexually mature in 14
30 days and can produce young every 7
8 days. Sheep keds typically live 4
6 months and can produce 12
15 larvae during her lifetime. Detached sheep keds can only survive for up to 4 days. Transmission to another host is largely through contact, especially from ewes to lambs. Adults crawl over the skin of its host and feed by inserting their sharp mouthparts into capillaries to suck blood. This causes considerable irritation to the animal that tries to relieve the suffering by rubbing, biting, and scratches at the wool, thus reducing the amount and quality of the fleece. The feeding puncture causes a condition known as “cockle” in tanned skins which downgrades them as weak and discolored hides. After mating a single egg is fertilized, hatches within the body of the female, and the larva develops through all three instars within the uterine pouch in the abdomen of the female fly. When the third instar is ready to pupate, the female extrudes the larva and glues it to the host’s hair. The larval integument hardens to form a brown, oval puparium. Depending on temperature and humidity, an adult ked emerges within 18
40 days from the time it was deposited.81,82

Rearing method(s) No laboratory rearing methods were found in the literature or known to the authors for the sheep ked. In vitro method(s) Contact—immersion—adult Sheep keds were collected from naturally infested sheep and identified. The parasites were maintained in plastic cups into which soaked cotton was placed to increase the humidity. The cups were covered with gauze to allow the free circulation of air into the cups. Insecticidal activity of test compounds (plant extracts) was tested using the adult immersion test by Drummond et al.83 Twofold serial dilutions of plant extracts (essential oils) and aqueous plant extracts were prepared in 2% aqueous solution of Tween 80 and distilled water, respectively. Tween 80 and distilled water served as negative controls, while ivermectin was used as a positive control. Activity of each dilution was tested by immersing a group of 10 M. ovinus in a Petri dish containing 3
5 mL of the test dilution for 1 min. The bioassay was performed in triplicate and the Petri dishes were incubated at 27 C
28 C and 80% RH. The sheep keds were studied under a stereomicroscope and corrected mortality was determined at 3 and 24 h.84

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In vivo method(s) Parasiticide efficacy studies against the sheep ked should be designed and conducted in accordance with the WAAVP guidelines for evaluating the efficacy of ectoparasiticides against biting lice, sucking lice and sheep keds on ruminants (see Appendix A of Ref. [85]). Diazinon and cypermethrin-based “pour-on” insecticides were tested against sheep biting louse and keds on naturally infested sheep.86 Mixed-sex Perendale sheep with moderate
heavy biting louse and sheep ked infestations were purchased. Animals were randomly assigned to treatment and control groups. Eight animals in diazinon treatment and control groups were assigned. The diazinon group was treated with 6.5 mL of diazinon by an applicator gun in a swathe down the middle of the back. Another treatment and control group of eight animals each were formed and the treatment group received cypermethrin (1 mL/5 kg bw) along the middle of the back. Ectoparasite assessments were done on TD 8, 48, 55, and 62 PT. Any insecticide-treated animal found with live ectoparasites was examined again 24 h later. Each insecticide was deemed to have lost its ability to reduce the ectoparasite populations when the mean reduction was ,90% compared to controls. The insecticides protected some sheep against reinfestation by the louse for B8
10 weeks PT. The cypermethrin pour-on prevented the establishment of the sheep ked for B9 weeks PT.

Sarcophagidae: flesh flies Wohlfahrtia magnifica Schiner, 1862—screwworm fly Biology and life cycle Wohlfahrtia magnifica, the spotted flesh fly, is a species of flesh fly belonging to the family Sarcophagidae, whose larvae attack mainly sheep and goats, but sometimes horses.86,87 It is the most important species of this genus throughout Europe (Mediterranean basin, eastern and central regions), Central Asia, the Middle East, North Africa, and China. It infests livestock, particularly sheep, camels, and poultry as well as cattle, horses, pigs, dogs, and rarely humans. Wohlfahrtia vigil occurs in North America infesting dogs, cats, mink and foxes. Both species cause myiasis, but W. magnifica is cutaneous, while W. vigil is furuncular. The body of adult W. magnifica is light gray with black contrasting spots on the abdomen (hence another common name, the spotted flesh fly) and measures 8
14 cm long. The female can produce up to six generations/year. The larviporous female flies deposit groups of 120
170 first instars (each 1 mm long) into wounds, mucus membrane of the eyes, the ear helix, and nasal cavity of a host. The larvae feed on tissue and develop to the third instar measuring 5
7 mm in length in 5
7 days, before leaving the wound of the host and pupae, burrowing into the ground.88,89

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Rearing method(s) Laboratory rearing Parental colonies of W. magnifica were established from larvae collected from natural infestations of sheep and cattle in central Hungary. First instars were harvested from gravid females and reared in groups of 5
20 on 1 of 6 artificial diets. The diets were based on various combinations of 5
7 of 8 ingredients: water, agar, blood (heparinized or dried), ground meat, egg yolk, low-fat milk powder, yeast and 10% formal. The larvae were incubated on the diets at 37 C. There was no mortality of first instars, which appeared to feed together in foci, in a natural manner. However, during the second phase of rearing, especially after renewal of the diet associated with disturbance of the larvae, many instars began to disperse, crawling over the surface of the media and feeding less intensively. Mortality of instars during all larval stadia was 64%
98%, compared to 33% in batches of third instars collected from natural infestations. The mean weights of puparia from artificial diets ranged from 38.7 to 59.3 mg, compared to 92.2 mg of puparia from larvae collected from natural infestations. There was a high mortality in the pupal stage ranging from 61% to 100%. Only a maximum of 6% of first instars were successfully reared to the adult stage. Further studies are needed to identify factors present or absent in the diets that contributed to the present poor development of W. magnifica in vitro.90 Flesh flies (Sarcophaga bullata) are readily maintained in the laboratory.91 Adults may be housed in commercial fly cages or in inexpensively constructed containers made of doweling and cheesecloth. Adult flies are fed sugar cubes and have water present at all times. For instar development, adult flies are provided a small piece of liver that is removed from the cage the following day. Another small piece of liver is placed in the cage B10 days later and removed 5
6 h later. This piece of liver will contain the first instar and is placed in a tinfoil cup that contains more liver. The tinfoil cup is placed in a large container of sawdust. Within B10 days the third instars will emerge from the liver and pupate in the sawdust. The pupae may be sieved from the sawdust and placed in a cage where the flies will emerge in B10 days. When a culture is no longer required, remove the source of water and within a few days the flies will expire. On-animal tissue rearing W. magnifica has a seasonal cycle from May to September. The free and obligate parasite phases were reared in the laboratory during 7 months, from September 16 to March 26. Between 97.49% and 98.27% of the third instars reached the pupal stage at 24 C and under a natural light cycle. Between 73.98% and 92.24% of the pupae developed into adults at 70% RH and 22 C. The ratio of females to males was between 1.87:1.00 and 1.40:1.00 at 70% RH and 24 C. In the trials involving the diapausd pupae, 35.29%

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emerged as adults during a period of 53 days (from January 23 to March 20), when held at an average storage temperature of 18 C. Eighty-five percent emerged during 2 days (February 23 and 24) when the storage temperature was 4 C.92 Larvae from adults of W. magnifica emerging from pupal cold storage (4 C for 80
100 days) were reared in wound and dead tissues. Of 2150 first instars placed on a mixture of muscle plus liver in a climate-controlled room, 47.1% molted to second instars, 6.1% to third instars, and 4.6% pupated. Two females emerged from these pupae after 14 days. To synchronize adult emergence, 191 pupae that were reared in living or dead tissues and when 2-, 8-, and 11-days-old, they were cold stored in lots according to age. Adult emergence was greatest in pupae of 2-days-old (57.1%) and pupae developing in living (22.5%) and dead tissues (8.7%). At 11-day-old cold storage in living tissues, the highest emergence was again in 2-day-old pupae (55.0%).93

In vitro method(s) No in vitro tests were found for W. magnifica or other flesh flies, the availability of the life cycle stages reared in vitro would allow for the development of parasiticide testing in vitro. In vivo method(s) Parasiticide efficacy testing against W. magnifica on ruminants should be designed and conducted in accordance with the WAAVP guidelines for evaluating the efficacy of ectoparasiticides against myiasis causing parasites of ruminants (see Appendix A of Ref. [94]).

Calliphoridae: blowflies Cochliomyia hominivorax: Coquerel, 1858—New World or American Screwworm Biology and life cycle Cochliomyia hominivorax belongs to the family Calliphoridae: blow flies. It is an obligate ectoparasite that infests a wide range of warm-blooded animals (cattle, horses, sheep, pigs, and dogs) including humans, and occasionally birds. The fly is distributed throughout the American tropics and subtropics from the southern United States to Central America, the Caribbean Islands, northern Chile, Brazil, Argentina, and Uruguay. The adult fly, like most calliphorid species, has a deep greenish-blue metallic color with a yellow, orange, or reddish face and three dark stripes on the dorsal surface of the thorax. It is B3 times the size of a housefly, measuring 10
13 mm in length. The female fly is oviparous laying batches of 200
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2
3 days during their 7
10 day life in compact masses on the skin around fresh or necrotic wounds or body orifices. Eggs hatch in 12
24 h and the gregarious larvae burrow head down or “screw” into the living tissue. Larvae develop for 4
10 days, growing to a length of B17 mm. Larvae exhibit a dark pigmentation of the tracheal trunks leading from the posterior spiracles to the 9th or 10th segment. They leave the wound, fall to the ground, and pupate in the top 2.5 cm of soil. The pupa last from 1 week in the summer up to 3 months in the winter. The entire life cycle from egg to adult is B24 days.88,89

Rearing method(s) Four larval diet and two adult diet formulations have been used in the mass production of C. hominivorax.95 Many formulations of artificial diets have been used in the screwworm mass rearing, evolving from the original lean meat
based diet used in Florida, to a liquid diet made from blood, milk, and egg powder prepared in Texas and Mexico, followed by a gelled diet with the same nutrient powders applied in Mexico and Panama, and to a cellulose fiber-based diet with the nutrient powders that was used later in the rearing plant in Mexico.95 Adult diets were also improved for the screwworm mass production.95 Artificial diets used in mass rearing Lean meat larval diet The first artificial medium used for laboratory level rearing was composed of 500 g ground lean beef, 750 mL milk, 250 mL citrated calf blood, and 0.5 mL formaldehyde.96 Melvin and Bushland96 were encouraged by the observation that larvae could finish their development in guinea pig carcasses kept at high temperature and by tests showing that neonates fed on raw lean beef or hard-boiled chicken eggs maintained at 37 C could reach the adult stage although in a small body size. During their tests of artificial diets, they found that the decomposition of the diets caused the death of young larvae. Although the aseptic technique could solve the issue, the method was not practical for mass rearing. Therefore formaldehyde was added to prevent the putrefaction of media to an acceptable degree. For mass production, milk was removed because of the volume and required cold storage.97 The formulation was changed to 1000 mL water, 6 mL formalin, 500 mL citrated beef blood, and 1000 g ground animal meats.95 Liquid larval diet The development of liquid media for mass rearing was initialized because of unsustainable supply, high price and required cold storage of meats.95 With a 3-cm layer of acetate fibers to hold the liquid and larvae, a standard optimal liquid medium (6% blood, 5% egg, and 1.3% milk) was perfected that maximized the screwworm life history parameters of biomass, survival, emergence, and fecundity.95 The liquid diet system

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encountered problems of small larval size, intensive labor requirements, waste contamination, premature or delayed larval crawl-off, and poor quality control.95 A polyester pad can be used as solid support for rearing C. hominivorax larvae in liquid diets.98 Gelled larval diet A gelled diet containing the polyacrylamide polyacrylate copolymers, 6%
7% spray-dried blood or defibrinated bovine blood, 3% dried egg, 3%
5% nonfat dried milk, 0.1% formalin, and 1.0%
1.2% sodium polyacrylamide gelling agent was developed to eliminate the issues with the liquid larval diet in the hydroponic rearing system.95 This diet did not reduce labor, the amount of diet used, was not biodegradable in the environment, and may accumulate and clog the pipes of the waste disposal system.95 Fiber-based larval diet A recycled newsprint product used as a bulking agent and cellulose fiber replaced the gelling materials.95 The fiber-based diet of 6% spray-dried blood, 4% spray-dried egg, 4% dried milk, 0.1% formalin, and 6% recycled newspaper product (Terra-Mulch) or 5% cellulose fiber (CF-100) was low priced, easier to handle and use, environmental friendly, and may be disposed in a landfill.95 However, more metabolic waste ammonia is released from the fibers than the gels.95

Adult diets A formulation with spray-dried egg powder, honey and molasses diets for flies showed promising results for both small-scale and mass-rearing application.95 A granulated sugar and milk powder (1:1) and a dry adult diet with sucrose and dried egg powder in various ratios have been adopted for mass rearing of adult flies.95 These diets in dry powder eliminate the mold issue in the fly cages.95

In vitro method(s) No in vitro tests were found in the literature, However, all the life cycle stages are available for the development of parasiticide in vitro tests should they be deem necessary. In vivo method(s) Parasiticide efficacy testing against C. hominivorax on ruminants should be designed and conducted in accordance with the WAAVP guidelines for evaluating the efficacy of ectoparasiticides against myiasis causing parasites of ruminants (see Appendix A of Ref. [94]).

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Phormia regina Meigen, 1826—surgical maggots or black blow fly Biology and life cycle Phormia regina is distributed in the northern Holarctic and is more common in livestock myiasis, particularly sheep, in northern United States and Canada. P. regina is important in forensic entomology to aid in the determination of postmortem interval in criminal investigations and in medicine for maggot therapy to clean out only necrotic tissue within a wound. The adult fly is dark metallic blue-black in color and the anterior spiracle is yellow or orange clearly standing out on the dark background of the thorax. The fly is about the size of a housefly or a little larger. The wings are specialized having a sharp bend halfway through the wing. They have a well-developed mostly white calypter. The adult fly feeds by sponging with dung constituting a major nutritional source for sexual development of both males and females. Female produce eggs that are completely mature. The female fly arrives within min to oviposit B250 eggs on orifices of the body, open wounds, and dead flesh. Flies are strong fliers capable of traveling up to 20 km in a day. Fully developed eggs hatch within 24 h at # 25 C. The larval stages feed and develop with the third instars producing heat that can raise the surrounding temperature of the developing site by .10 C. The third instars move away from the body and burrow into the ground to pupate with emergence of adult flies 7
14 days later. The life cycle of P. regina from egg to pupae at B20 C takes 6
11 days. Rearing method(s) Laboratories have strived to rear P. regina under conditions to avoid the stench and other undesirable features associated with the putrefying meat on which the larvae ordinarily feed. Aseptic cultures of the larvae used to treat osteomyelitis99,100 have been developed with meat and synthetic diets under sterile conditions.101
103 Adult flies are maintained in cages in a rearing room at 27.5 6 3 C, with a 16L:8D hour cycle. Multiple generations are maintained in a single cage, and B1000 adult flies are introduced every 1
2 weeks (B1 month adult life span in colony). Cages are 0.61 3 0.61 3 0.61 m in size (collapsible cage No. 1450D) to avoid any potential selection on adult size. Adults are provided sugar water as a carbohydrate source, and raw beef liver for protein and as an ovipositional substrate. After egg laying, eggs and liver are maintained in plastic boxes (15 3 10 3 5 cm) in a Percival growth chamber set at 26 6 1.5 C. Within the plastic container, liver and feeding maggots are placed on a plate that rested on pine shavings. The pine shavings provide an area for larval migration at the end of the third instar and a substrate for pupation (larvae bury themselves within the pine shavings after migratory

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movement). Because larvae of P. regina are more likely to move off the substrate as compared to other species, moist sand is used instead of pine shavings to avoid desiccation as larvae move on and off the liver, as well as providing an acceptable pupation medium. Twenty eggs (collected within 30 min of oviposition) are counted onto a moist black filter paper triangle and placed in direct contact with 10 g of beef liver in a 29.5 mL plastic cup. The cup is placed in a 7 3 7 3 10 cm plastic container that had 2.5 cm of wood shavings in the bottom. The container is then placed randomly in an incubator.104 A chunk of ground horse meat (roughly cubical, 3
4 in on a side) is placed for 24 h into a breeding cage of adult flies, on which oviposition will occur. Eggs are removed from the surface of the meat using sterile forceps or needles and placed in 2% NaOH for 15
20 min. The eggs are rinsed several times in 70% ethanol and allowed to sit in 70% ethanol for 15 min at the end of which time the eggs are sufficiently sterile for introduction into the culture bottles. Eggs (B500) are added to a culture bottle containing powdered casein (630 g), Brewer’s yeast (63 g) and powdered agar (21 g) in the proportions of 30:3:1 to which is added lanolin (3
4 g) and modified Belar solution containing phosphate (60 mL) to 30 g aliquots of the three constituents in half-pint milk bottle with a nonabsorbent cotton plug. The larval rearing medium is autoclaved for 15 min at 15
17 lbs pressure. The culture bottles are incubated at 25 C and 70% RH. Eggs hatch in ,24 h and by about the eighth day, the larvae cease to feed and pass into a prepupal stage. Then the cotton plug is removed, and autoclaved sawdust is poured to a loosely packed level of 2 cm below the top of the bottle. The bottle is covered with a disk of fine meshed wire (40
80 mesh). At 25 C and 70% RH, the duration of the pupal stage is 5
6 days, with fly emergence 15
16 days after the eggs are laid. Pupae are placed in a small cup that has been securely mounted on the end of a piece of plywood (1.5 3 6 in.). A small cup (20 mL) on the other end contains sugar and water. A piece of paper toweling is laid on the water surface. The cup and plywood assembly is placed in a 1-quart Mason jar laid on its side. A short taper cork (No. 7) is fastened to the underside of one end of the plywood support; the other end rests in the neck of the jar. The mouth of the jar is covered with a wire screendisk held in place by the jacket of an ordinary two-piece Mason jar lid. The screen is pierced with a small hole, fitted with a cork, through which a pipette may be inserted to replenish water in the cup containing the sugar solution. The pupae should be toward the rear of the jar. On emergence the young flies must be supplied with sugar and water along with fresh ground horse meat if they are used for breeding. On a meat diet, flies will oviposit by the end of the first week after emergence. A halved orange can also be added in the cage. Flies should not be overcrowded, with B300 flies/ft3 of cage space being satisfactory.105

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In vitro method(s) No in vitro tests of P. regina were found in the literature. However, those tests described later for Lucilia sericata might be applicable. In vivo method(s) Although P. regina in vivo study designs are not specifically covered, one might consider consulting the WAAVP guidelines for evaluating the efficacy of ectoparasiticides against myiasis causing parasites on ruminants (see Appendix A of Ref. [94]). A mouse/blowfly assay using first instars of P. regina or L. sericata allows for the determination of systemically active parasiticides against developing blowflies. The assay also provides information on the absorption, distribution, metabolism, and excretion of the compound under test. Mice (four/test group 3 two dosage rates 3 two exposure times) are administered 0.5 mL PO (gastric gavage) of compound concentrations of 0.5 and 5.0 mg/ kg bw. Test compounds are prepared in corn oil or corn oil:DMSO (10:1). Control mice are treated with corn oil:DMSO (10:1) in the same manner and volume as the treated mice. Half of the mice in each group are euthanized 8 h PT and the remainder at 120 h PT. The hair is removed from the hind legs of the dead mice and the muscle/bone detached from each mouse. Four hind legs from two mice in the same treatment group are placed into 7 dram vials (two legs per vial). The tissue will cool to RT and then 10 first instars are added to each vial. The samples remain at RT and the assay scored for adult fly emergence 2
3 weeks later. Efficacy is determined by the number of emerged adult flies in the treated groups by dosage as compared to the control group. Lucilia (5Phaenicia) sericata Meigen, 1826—sheep blowfly or wool strike fly or common green bottle fly Biology and life cycle L. sericata and Lucilia cuprina are members of the family Calliphoridae: blowflies. The larvae of both species infest sheep and infestation is commonly known as blowfly strike. While both species are found worldwide, generally L. sericata is more common in cool-temperate zones, while L. cuprina is found in warm-temperate and subtropical climates. L. sericata is the most important species of sheep myiasis throughout the northern Europe, while L. cuprina is absent from most of Europe. L. sericata is present in Australia, but generally confined to urban habitats. It is the primary myiasis fly in New Zealand and South Africa. L. sericata plays an important role in forensics, where the larvae or maggots help to determine the period of insect colonization as it relates to the time of death, aiding law

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enforcement in their investigations. Medical treatment using maggot therapy can help to heal infections that are otherwise incurable.106 Sheep blowflies are important ectoparasites of sheep causing sheep strike or sheep blowfly strike that affects an estimated 1 million sheep, as well as 80% of sheep farms each year in the United Kingdom. Larvae cause cutaneous myiasis on sheep, initially feeding on serous exudate at the skin surface before penetrating the skin and feeding on the underlying tissue. This causes a huge economic impact on sheep and wool production. The adult flies are metallic green in color, 10
14 mm long, and are characterized by the presence of a bare stem vein and the presence of three pairs of bristles on the mesothorax. Both species require a meal especially of protein to produce eggs (anautogenous). Female flies deposit batches of 225
250 light yellow eggs at 3-day interval during their B7 days life. Adults are diurnal. The life cycle from egg to adult can be as short as 7 days but is typically B4
6 weeks depending on temperature. Flies can live a month or longer and can also hibernate.88,89,107 The eggs of L. sericata are often laid in batches or masses in a wound, carcass or corpse, or in necrotic or decaying tissue. The eggs are usually white but can be a pale yellow. The eggs are elongated with one end tapered slightly and measure B1.5 mm long. They hatch in B9 h in warm, moist weather, but can take up to 3 days in cooler weather. In the northern Europe the fly lays its eggs in sheep wool. The larvae then migrate down the wool where they feed directly on the skin surface. This can cause massive lesions and secondary bacterial infections. Hatched larvae feed superficially on the epidermis and lymphatic exudate or necrotic tissue. The larvae develop through three instar stages in B2
19 days. The third instars are B10
14 mm long, grayish-white or pale yellow in color, sometimes with a pink tinge. The anterior extremity bears a pair of oral hooks and stigmatic plates are situated on the broad, flattened posterior end. The second segment bears a pair of anterior spiracles. The mature smooth larvae leave the wound or carcass and drop to the ground. They may wander fair distances over or through soil before pupating. The pupal stage lasts 3
7 days in the summer to much longer (hibernating) in the winter.

Rearing method(s) Adult L. sericata are held in a fly cage. Adult flies need to be fed protein before they will lay eggs. Slices of liver are provided in a Petri dish for 2 days inside the cages after flies have emerged from pupae and then remove the liver. The flies will be ready to oviposit in B2
3 days and every 3
5 days thereafter. To collect fly eggs, place a Petri dish bottom with a few slices of liver and place the dish in the cage. Allow the flies to oviposit for B2 h. The liver

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should be dotted with numerous clumps of eggs. After removing the dish, replace the Petri dish lid and wrap the dish in a moist paper towel to prevent desiccation. Incubate the eggs at 25 C, 50% RH, and a 16L:8D hour cycle. The eggs will hatch in B16 h as newly emerged first instars. Set up a larval culture tray using B0.5 kg of meat. Cover the bottom of a larval tray with 1
2 cm of vermiculite, and place the meat on top. Add 2000
3000 larvae (B0.5 g, a clump about the size of a pea) to the meat. The larvae will feed for 5
7 days at 25 C and 50% RH and then leave the meat and pupate in the vermiculite. The meat should be removed as soon as most of the larvae have crawled into the vermiculite to pupate. Pupae should be sieved from the vermiculite once the majority has hardened. Place pupae in a Solo cup B500 pupae (20 g) in a small cage, or B1000 (40 g) in a large cage. Incubate the pupae at 25 C, 50% RH, and a 16L:8D hour cycle. Place a water cup and sugar cup in the cage. Adult flies will begin emerging in B2
5 days. Check the flies a couple of times a week for water and food. Waterers should always be at least half full. Flies will die quickly within an hour without water and food. As the flies get older, freeze them overnight and then discard them. Be sure to also freeze all leftover medium from larval pans, eggs cups and pupae overnight to prevent a major laboratory outbreak. If any red areas appear on the water wicks or any dead flies have fuzz on them, then add antibiotics to the water. Fleece-rot has been experimentally induced in vitro by wetting and incubating Merino wool samples embedded in serum agar.108 Gravid females of L. cuprina are readily attached to the culture plates to oviposit. Fly strike and larval development occurred with freely available serum.

In vitro method(s) Contact—filter paper or cloth square—larvae Circles (9 cm in diameter) of white polyester cloth are cut and weighed. The test compounds are initially dissolved in DMSO to ensure the dissolution of the compounds. A series of five dilutions of the DMSO solutions of the test compounds and β-cyfluthrin are prepared in acetone. These serial dilutions will be prepared at a rate to provide concentrations of active ingredient on the cloth of between 0.001 and 10.0 mg/cm2. Cloth circles (three/dilution/ exposure interval) are dipped in the series of dilutions of the three compounds, allowed to dry, and then reweighed. Thus all test material and cloth/ chemical concentrations will be prepared in triplicate. The amount of active ingredient deposited on the cloth will then be calculated by comparison of the two weights. For both negative controls, cloth circles will be dipped in acetone and circles will be dipped in water. First instar and feeding third instars of L. sericata are placed individually onto the cloth in a Petri dish and allowed to crawl over the material for 0.5,

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1, and 2 min. Each dosed cloth is used for the exposure of 20 larvae only. Following contact, first instars are placed onto diced lamb liver, in pots of 20 larvae/10 g of liver, to complete development at 25 C, 50% RH, and a 16L:8D hour cycle. Under these conditions, larval development will require B48
60 h. At the end of this period the number of larvae entering the larval wandering stage and the number of wandering larvae that pupate subsequently are recorded. For third instars, exposed larvae are placed directly into sawdust, incubated at 25 C, low % RH, and a 16L:8D hour cycle, and then the number pupating will be recorded after B3
5 days. Following the initial assay at the concentrations specified earlier, the trial may be repeated with a more focused range of test and positive control chemical concentrations. The percentage mortality of all treatment groups are determined and the LC50 and LC90 of the test substances and β-cyfluthrin determined by probit or regression analysis. Contact—chromatography paper—larvae Test compounds are dissolved and diluted to the testing concentration in acetone (for random screening the initial test concentration of 100 ppm is recommended). About 1 mL of the test solution is added to a strip of chromatography paper (No. 3 paper) and allowed to dry. Individual strips (3 3 12 cm) are then rolled lengthwise and flattened laterally to allow air spaces between the layers of rolled paper being placed in the vials (8 3 25 mm glass or polystyrene shell vials). About 1 mL of bovine serum fortified with yeast extract (2% w/v) and buffered with monobasic potassium orthophosphate (0.5% w/v) is then added to each tube.109 Newly hatched L. sericata larvae (B50, a clump B 1/2 the size of a match head) are added to each tube using a tooth pick, the tube is then plugged with cotton wool. Assay tubes are held at B27 C and 70% RH. Mortality is determined at 24 or 48 h. The numbers of dead and alive larvae are counted. Since this bioassay is based on a serum-soaked, treated filter paper substrate, it is suitable for testing/detecting both contact and systemic test compounds. Contact—wool—larvae This assay provides a slightly more realistic form of contact than the cloth contact assay. However, it can only be used for first instars and for relatively short periods of contact. For larger second or third instars, individual larvae can be topically treated with known concentrations of active ingredient from a Hamilton syringe. Aliquots of clean sheep wool (collected and stored frozen) are weighed. The test compounds are initially solubilized in DMSO. A series of five dilutions of the test compounds and β-cyfluthrin are prepared in acetone. These serial dilutions are prepared at a rate to provide

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concentrations of active ingredient on the cloth of between 0.001 and 10.0 mg/cm2. The wool (three aliquots/dilution/exposure interval) are dipped in the series of dilutions of the test compounds, allowed to dry, and then reweighed. Thus all test material and wool/chemical concentrations will be prepared in triplicate. The amount of active ingredient deposited on the wool is then calculated by comparison of the two weights. For both negative controls, wool aliquots are dipped in acetone or wool aliquots will be dipped in water. First instars of L. sericata are then be placed onto bundles of individual wool fibers in a Petri dish and allowed to crawl over the material for 0.5, 1, and 2 min. Following contact exposure at each concentration and time interval, the batches of 20 first instars are placed onto diced lamb liver, in pots of 20 larvae/10 g of liver, to complete development at 25 C, 50% RH, and a 16L:8D hour cycle. Under these conditions, larval development requires B48
60 h. At the end of this period, the number of larvae entering the larval wandering stage and the number of wandering larvae that pupate subsequently are recorded. Following the initial assay at the concentrations specified earlier, the trial may be repeated with a more focused range of test and positive control chemical concentrations. The percentage mortality of all treatment groups are determined and the LC50 and LC90 of the test compounds and β-cyfluthrin determined by probit or regression analysis. Contact—96-well plates—larvae Mother plates of test compounds are subsampled into labeled 96-well plates. The subsample plate is covered with Mylar film and stored frozen at 220 C until assayed. About 1 μL of test sample (diluted to 10 mg/mL in DMSO) is pipetted into each test well. A solution of 500 μL 80:20 acetone:water with crystal violet is added to each well to make a 20 ppm solution, and the plate is placed on a shaker for 30
40 s. A 200 μL sample of the 20 ppm test solution is added to a well in the assay tray, containing three filter papers measuring 0.5 in. diameter, and allowed to dry. Fly eggs collected from the breeding colony are stored at 27 C overnight to hatch. On the assay day, 1- or 2-day-old blowfly larvae are mixed in a crystallization dish with a serum preparation (500 mL serum, 10 g yeast extract, 2.5 g sodium orthophosphate monobasic, and stored frozen in 50 mL aliquots). Two-day-old larvae will have been stored in a refrigerator overnight in liver plates, covered with a damp towel to prevent desiccation. A sample of the 200 μL of this mixture containing B20 larvae is then added to each well. Assay trays are covered and held at B27 C and 70% RH. Mortality is determined at 24 and 48 h. The percentage of dead and/or abnormally small larvae is recorded. Active test compounds are defined as .80% mortality after 48 h. Active test compounds are further tested at 2.0

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and 0.2 ppm concentrations as described earlier. Since this bioassay is based on a serum-soaked, treated filter paper substrate, it is suitable for testing/ detecting both contact and systemic test compounds. Contact/Feeding—liver—larvae Batches of neonate blowfly larvae of L. cuprina are transferred onto homogenized bovine liver containing varying concentrations of cyromazine or dicyclanil (mg/kg), and the numbers of larvae pupating and completing development is recorded.110 Homogenized liver containing 1 mg/kg cyromazine (susceptible discriminating concentration, i.e., concentration lethal to susceptible individuals) has been reported to kill all susceptible larvae and has been used for more than 25 years to monitor blowfly populations for resistance. Contact—topical/fly motor activity and wing beat frequency—adult Blowflies are briefly anesthetized using CO2. Copper recording electrodes (50 μm) are inserted into the left and right dorsolongitudinal muscles from the anterior of the pterothorax, and the common reference ground electrode was inserted into the left and dorsolongitudinal muscle using micromanipulators. Muscle insertions were located by referencing their locations to prominent dorsal setae. Once positioned correctly, the electrodes themselves were sufficient to hold the blowflies and allow flight to occur. Electrical activity from the flight muscles was recorded. At the same time, acoustic signals from wing beats were recorded. Ten minutes after recovering from CO2 anesthesia, 50 μg of test compounds were topically applied to the tip of the abdomen, or 50 μg of octopamine or GABA were injected into the abdomen using a microsyringe. Control insects were topically treated or injected with 5 μL of DMSO or water. Recordings began immediately and firing rates (impulse/s) were calculated by counting the number of action potentials that occurred after the application of the test compound. The mean responses and their standard errors were determined, and amplitudes were calculated with their standard errors. Each compound was tested on a minimum of three blowflies. The tethering apparatus did not restrict flight, and control flies could be induced to fly at any time throughout the experiment by touching the wings or tarsi.111

In vivo method(s) Parasiticide efficacy testing against L. sericata and L. cuprina on ruminants should be designed and conducted in accordance with the WAAVP guidelines for evaluating the efficacy of ectoparasiticides against myiasis causing parasites of ruminants (see Appendix A of Ref. [94]).

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Oestridae—bot flies Oestrus ovis Linnaeus, 1758—sheep nasal bot fly Biology and life cycle Oestrus ovis is a member of the family Oestridae, commonly known as bots and warbles. The nasal bots are placed in the subfamily Oestrinae and develop exclusively in the nasopharyngeal cavities of mammals. O. ovis is widely distributed worldwide wherever sheep, deer, and goats are found (North America, Central America, Brazil, Mediterranean Sea area, the Middle East, northern Europe, Australia, and South Africa). The adult fly is light to dark grayish in color B10
12 mm in length. It has small black spots that are especially prominent on the thorax, and it is covered with short, light brown hair, superficially resembling a bee-like insect. The mouthparts are atrophied being reduced to small knobs, hence they do not feed living only 15
30 days to mate and reproduce. Adult female flies are oviviparous with fertilized eggs hatching to 1 mm long larvae within the body of the female (larviparous). During flight the female deposits up to 25 live first instars at a time, in or near the nostrils of the host. The white or slightly yellow first instars crawl up the nasal passages by using their mouth hooks. They attached and feed on the secretions and desquamated cells of the mucous membranes and then enter the frontal sinus before molting to the second instar. This stage measures 4
12 mm in length developing to third instars in the frontal sinuses. The third instar is B20 mm long, tapering anteriorly and ending with flat surface posteriorly. There are large, black, anterior oral hooks, and the ventral surface bears rows of small spines with conspicuous black stigma plates on the posterior surface. The third instars reenter the nasal cavities where they crawl or are sneezed out onto the ground. The length of time the larvae take to mature may take 25
35 days in warm weather with up to three generations/year or it may be delayed up to 10 months in cold weather during the winter. They cease development in autumn with the first instars overwintering within the head of the host, not migrating to the frontal sinuses until the following spring. Pupae take 3
9 weeks to mature depending on climatic conditions. Adult flies emerge from the puparia in the ground and burrow up to the surface. Adults have a life span of 2
4 weeks.88,89,107 Rearing method(s) No in vitro rearing methods were found in the literature or known to the authors.

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In vitro method(s) Contact—Petri dish—larvae O. ovis first, second, and third instars were collected from the heads of goats slaughtered in the municipal slaughterhouse. Larvae were pooled according to larval stage and cultured in Petri dishes at 20 C
25 C (5 mL) in three randomly assigned groups: 0 (Control), 5% and 10% neem extract in PSS. Each individual larva was observed for movement under a stereoscope. When a larva ceased to move it was considered as dead, and withdrawn from the culture. Time to larval mortality was recorded by checking the cultures at 0, 0.5, 1, 1.5, 2, 3, 6, 9, and 12 h. Further observations were carried out every 12 h up to 144 h postculture. A total of 1193 first instars, 45 second instars, and 62 third instars were included in this in vitro test.112 In vivo method(s) Parasiticide efficacy testing against O. ovis on ruminants should be designed and conducted in accordance with the WAAVP guidelines for evaluating the efficacy of ectoparasiticides against myiasis causing parasites of ruminants (see Appendix A of Ref. [94]). Hypoderma bovis Linnaeus, 1758—warble fly or northern cattle grub; Hypoderma lineatum Viller, 1789—heel fly or common cattle grub Biology and life cycle Hypoderma bovis and Hypoderma lineatum are members of the family Oestridae, subfamily Hypodermatidae: cattle warbles or bot flies. These species are widespread throughout the Northern Hemisphere. H. lineatum is prevalent in Canada and the United States. Two species of Rhinoestrus (i.e., Rhinoestrus purpureus and Rhinoestrus usbekistanicus) cause nasal myiasis in horses, donkeys, and zebras. In the past 15 years myiasis caused by R. purpureus has been reported in Egypt and by R. usbekistanicus in Senegal and Niger, both in horses and in donkeys. The adults (15 mm long) are only active on warm, sunny days from April to June. H. bovis is found principally in the northern United States and Canada with adult flies (13 mm long) active during mid-June to early September. The adult flies of both species are bee like in appearance with dense hair in a characteristic light-dark color pattern, but H. bovis has a more profusely hairy thorax than H. lineatum. The mouthparts of both species are nonfunctional and lack palps. The nonfeeding adult flies usually live only B2
3 days with only one generation/year. However, female flies emerge with all their eggs fully developed and can begin to oviposit within 20 min after copulation.

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Eggs of H. lineatum are typically attached in rows of 3
15 on a single hair, whereas H. bovis generally attached eggs singly. Eggs are most commonly deposited on the lower regions of the legs and lower body with a single female laying as many as 300
600 eggs during short adult life span. Eggs hatch in B4
7 days and the first instars (,1 mm in length) crawl down the hairs and either burrows directly into the skin or into the hair follicles. The larvae then burrow beneath the skin, but the mortality is estimated to be .50%. The precise larval pathway and migration pattern depends on the species. During a period of B4 months, usually by autumn, the larvae of H. bovis migrate below the skin along nerves to the spinal cord reaching the epidural fat of the spine in the region of the thoracic and lumbar vertebrae. H. lineatum migrates between the fascial planes of the muscle and along connective tissue to the submucosa of the esophagus. The larvae about the size of a rice grain overwinter in these specific locations. About 9 months after oviposition, larval migration resumes in the spring until they reach the skin of the back of the animal B25 cm either side of the midline. The larvae cut a small hole to the hide surface, reverse their position with its posterior spiracles close to the opening to breath, and form a characteristic small swelling, the “warble,” an Anglo-Saxon word for boil. The larvae molt twice, growing rapidly, more than doubling in length. The warbles of H. lineatum appear between January and February up to April, while those of H. bovis appear between March and June. After 50
60 days the third instar exits the warble pouch and drops to the ground where pupation occurs. The duration of the pupal stage ranges from 2 weeks to 2 months depending on the climate conditions. The entire life cycle takes B1 year.88,89

Rearing method(s) Blagoveshchenskii and Pavlovskii113 devised a means of collecting the larvae when they left the host. The body of the infested animal is covered with a linen case, which consists of two parts. The top part is stretched over a frame of wire covered with felt. The frame is shaped to fit close to the sides of the animal and leave a space over its back. A slit can be opened at the side to allow observations. The belly of the animal is covered with a piece of linen that is buttoned to the frame so that it sags a little. The whole is kept in position by straps. The larvae emerging from the warbles either remain in the top part of the case or make their way into its lower sagging part. Larvae from cases on cattle kept in stalls should be collected at least three times a day. If the infestation is slight, individual larvae can be caught in small muslin bags sown on flat rings that are glued on the skin of the animal over the warbles. The larvae collected were allowed to pupate in flower pots filled with earth or moss and covered with muslin over a bell-shaped

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frame. To obtain natural conditions, pots without bottoms were sunk in the soil and covered with a portable cage. First instars of H. lineatum and H. bovis are dissected from the esophagus and spines of cattle at an abattoir. Larvae are placed in penicillin saline for 30 min before being transferred to 2 oz medicine flats that served as culture bottles. A mixture (EYL medium) of 5 g lactalbumin hydrolysate, 0.1 g yeastolate, and 50 mL of horse or bovine serum/L of ESS at pH 7.4 is prepared also containing phenol red as an indicator and antibiotics: penicillin, streptomycin, mycostatin, and polymyxin. The EYL medium (9 mL with a depth of 2
3 mL) is added to cultures and incubated at 37 C. Larvae are transferred to fresh bottles of medium very 3
4 days. A mass-culture technique was also devised along the lines of that used for the culture of the cestode, Hymenolepis diminuta.114 About 10 first instars were arranged on filter paper placed inside a sloping glass tube measuring 14 3 1.5 cm. The EYL medium is run in slowly from a reservoir Marriott bottle and the tubes incubated at 37 C. First instars of H. lineatum and H. bovis survived an average of 26.1
62.5 days with individual survival of up to 11 days.115 A new method116 was successfully used in collecting the mature larvae of the cattle warble fly by means of woven wire flooring. In laboratory colonies the influence of temperature on pupal development, percentage of hatched adults and sex ratio, weight of pupae and adults, its changes during their life cycle, copulation and survival of adults were studied. Field experiments involved studies on the influence of humidity on the survival of pupae, percentage of hatched adults, duration of the pupal stage in the spring and summer periods. Stability of the total sum of temperatures above the temperature threshold necessary for the development of adult in pupa, facilitating to determine the term of hatching, was demonstrated. Laboratory rearing made it possible to obtain gravid females of the cattle warble fly for experimental purposes. Transplantation—mammalian hosts First instar larvae of H. lineatum from the gullets of commercially slaughtered cattle were transferred to goats, rabbits, and white mice.117 Larvae were transferred directly into the SC tissue of the goats on the thigh or shoulder. Up to 20% of the larvae implanted in a goat were recovered at necropsy, and all were found in areolar connective tissue. None reached full growth, though some survived in the goats for 4 months. One larva that hatched from eggs laid on the hind leg of a goat by a captive fly reached the gullet in ,2 months. Rabbits were more satisfactory experimental hosts and as many as 85%, of the larvae implanted in the thigh of a rabbit were recovered after 26 days. Larvae transplanted into mice remained in connective tissue for more than a

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week, but many died during the third and fourth week. Mice are especially useful for testing rare and costly chemicals because of their small size.118

In vitro method(s) Contact—blood serum—larvae In vitro bioassays on larvae of H. lineatum were attempted in physiological saline and in normal bovine blood serum.117 Blood was collected in sterile containers, defibrinated, chilled, and the serum separated by centrifugation. The serum was held refrigerated until use. First instars (found in SC connective tissue of the esophagus, or gullet, within 2
3 months after egg hatch and obtained within 24 h after slaughter of cattle) were washed in several changes of PSS in sterile beakers, before being placed in the blood serum. Larvae (5
20) were placed in a sterilized beaker containing 10 mL of normal serum. The test chemical was dissolved in acetone or other suitable solvent depending on the compound and then pipetted into the serum containing the larvae. The beakers were placed in a closed glass jar containing water to prevent contamination and evaporation, and incubated at 30 C for 24
48 h. Some 200 chemicals were tested on first instars of H. lineatum in blood serum, which proved to be a better medium than PSS at the concentrations of chemicals tested. However, it is doubtful whether the results obtained in tests in vitro are reliable indices of the possible chemotherapeutic action of the compound in vivo.117 Contact—culture medium—larvae This is apparently the first historical record115 of the in vitro exposure of insecticides of first instars of H. bovis. Systemically active organophosphate insecticides were added to a glass tube containing B10 first instars arranged on filter paper inside a sloping glass tube containing EYL medium. Crystals of pure insecticide (fenchlorphos, trichlorphon, and crufomate) were dissolved in 80% ethanol to yield a stock concentration of 125 ppm and further serially diluted with EYL medium to 0.0625 ppm. A control solution of EYL containing 0.1 mL 80% ethanol/50 mL total was used. Larvae were rapidly killed by exposure to 32 ppm of trichlorphon and 8
16 ppm of fenchlorphos.

In vivo method(s) Parasiticide efficacy testing against Hypoderma species and Przhevalskiana silenus on ruminants should be designed and conducted in accordance with the WAAVP guidelines for evaluating the efficacy of ectoparasiticides against myiasis causing parasites of ruminants (see Appendix A of Ref. [94]).

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Gasterophilus nasalis Linnaeus, 1758—horse nasal bot fly; Gasterophilus intestinalis De Geer, 1776—common horse bot or horse stomach bots; Gasterophilus haemorrhoidalis—Linnaeus, 1758—nose bot fly Biology and life cycle The Gasterophilus species belong to the family Oestridae, subfamily Gasterophilinae: horse bots. Gasterophilus intestinalis and Gasterophilus nasalis are common in most parts of the world where horses are raised, whereas Gasterophilus haemorrhoidalis is uncommon. The adults of the three species possess nonfunctional mouthparts, so they cannot feed. Consequently they are short lived with an effective life span of only a few days to B2 weeks. Horse gastrointestinal myiasis caused by larvae of Gasterophilus spp. flies has a worldwide distribution and, where present, it is primarily caused by larvae of G. intestinalis and G. nasalis.119 Other species, that is, Gasterophilus inermis, Gasterophilus pecorum, and G. haemorrhoidalis, present in different or in the same regions of the gastrointestinal tract.120 Adult flies of Gasterophilus species are brown with densely covered yellowish hairs and superficially resemble honey bees in appearance possessing on pair of clear wings with brown patches. They are medium to large flies measuring 10
20 mm in length. Adults mate shortly after emergence (3
10 weeks from pupal formation) and females begin oviposition almost immediately. Female flies have a strong and protuberant ovipositor. Gasterophilus spp. eggs (B1
2 mm in length) differ in color and placement on the host. G. intestinalis lays pale yellow eggs especially on the forelegs and shoulders, up to 1000/female fly. Moisture and friction from the horse licking itself cause the eggs to hatch by stimulation in B7 days. Gastrophilus nasalis lays B500 yellow eggs around the submaxillary region, which hatch in B7 days without stimulation. The female G. haemorrhoidalis lays 150 black eggs around the lips of the horse, which hatch in 2
5 days. After hatching, G. intestinalis larvae are licked into the host’s mouth, but G. nasalis and G. haemorrhoidalis larvae burrow under the skin to the mouth. The first instars are extremely small and motile and begin development in the mouth, tongue, and gums of the host. The first instars develop to the second instar on the pharynx and attaches to the base of the tongue. After B1 month wandering in the mucosa of the tongue and mucosa of the mouth, the larvae of all three species migrate to the stomach. The third instars are reddish and measure 2 cm in length. Stomach attachment location is species specific with G. intestinalis attaching to the squamous gastric mucosa along the margo plicatus, G. nasalis attaching to the dorsoproximal part of the duodenum, and G. haemorrhoidalis attaching to the gastric epithelium within the stomach. The bots have a narrow hooked anterior end that tapers from a broad, rounded body. There is a single row of

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spines/segment on G. nasalis larvae, and two rows/segment on G. intestinalis. Gastrophilus intestinalis have mouthparts that are not uniformly curved dorsally, and the body spines present have blunt ended tips. Larvae over winter attached to the mucosa of the stomach of the host. Larvae then remain and develop within the stomach for around 10
12 months. They mature in the following spring or early summer (generally between May and September), detach from their surface, and are passed out in the host feces. G. haemorrhoidalis also attaches to the rectal mucosa before being passed out in the feces. The larvae burrow into the ground and pupate on pasture. The pupae are known to be sensitive to frost, moisture, and flooding, so the environmental conditions present play a significant role in the success of the parasite.88,121

Rearing method(s) The method of collecting larvae of Hypoderma from cattle by Blagoveshchenskii and Pavlovskii113 is also suitable for collecting larvae of Gasterophilus from horses. Larvae of Gastrophilus about to leave horses are easily obtained by introducing the arm into the rectum of the animal and extracting them. They are reared by allowing them to pupate in flower pots containing horse dung with a layer of soil at the bottom. Holding temperatures and maturity of third instar G. intestinalis affected the time required for pupation and subsequent eclosion.122 As temperature increased, the duration of the larval stage decreased. Larvae removed from horses in June developed slower than those removed from horses in July and August. Changes in temperatures and humidity affected larval pupation and adult emergence. The most favorable environment tested for pupation and adult emergence was 29 C and 80%
92% RH. Larvae held at 50 C failed to pupate, whereas those held at 38 C pupated but did not eclose. Adults emerging from pupae subjected to low humidity during the prepupal stage were malformed. When third instars were removed from the stomachs of horses during March through August, the rate of development and the percent pupation and eclosion increased with the progression date of removal. In vitro method(s) Bello123 described a method by which G. intestinalis eggs are incubated, disinfected externally, and artificially stimulated to hatch in vitro. The percentage of larvae hatched varied directly with increasing differences between incubation and stimulation temperatures. Immersion for 6
7 h in a 0.5% aqueous solution of 1
100 commercial trypsin at a pH of 7.2
8.0 and at a temperature of 38 C hydrolyzed the cement that attaches the eggs without impairing the natural qualities of the hair.124 These methods may be applicable to testing the ovicidal and larvicidal activity of compounds in vitro against the eggs of Gasterophilus spp.

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In vivo method(s) Drug efficacy testing guidelines against larvae of horse bot flies, Gasterophilus spp., is provided in the WAAVP guidelines for evaluating the efficacy of equine anthelmintics (see Appendix A of Ref. [125]). An account is given by Drummond et al.126 in the first paper of tests carried out in 1959 on the effectiveness of coumaphos, dimethoate, fenchlorphos, trichlorphon, dichlorvos, and other compounds for the systemic control of larvae of Gasterophilus in horses in Texas. The insecticides were administered by stomach tube, in the feed, or by IM injection. The dung of the horses was examined for larvae of Gasterophilus 1 day before and daily for 1 week PT, after which the horses were slaughtered and their gastrointestinal tracts examined for the presence of larvae. All larvae found were G. nasalis or G. intestinalis. No larvae were expelled when the horses were treated with coumaphos, fenchlorphos, or dimethoate administered by stomach tube. Dichlorvos at 50 mg/kg and trichlorphon at 75 and 100 mg/kg were 100% effective. When added to the feed, dichlorvos at 25 and 50 mg/kg and trichlorphon at 25
75 mg/kg provided effective control. Trichlorphon at 25 mg/kg was fairly effective when injected IM. Most of the larvae expelled during the first 3 days were alive, but very few pupated. Cuterebra jellisoni Curran, 1942—rodent or rabbit bot fly Biology and life cycle Rodent or rabbit bots are members of the family Oestridae, subfamily Cuterebrinae. They are exclusively distributed in the Nearctic and Neotropical regions. Larval species of the genus Cuterebra are largely dermal parasites found in the furuncular lesions in the skin of rodents and rabbits but may occasionally infest dogs and cats. Adult flies measure up to 20 mm in length and are covered with dense, short hair with a blue-black colored abdomen. They have small, nonfunctional vestigial mouthparts and do not feed. Female flies do not lay eggs directly on the host but instead oviposit on the ground near entrances of rodent burrows or on grass near host runs where the moist, sticky eggs are picked up to the fur coat. Female flies can lay one to several thousand eggs, in groups of around 5
15/site. The first instars hatch instantaneously as the host comes into contact with the egg, and the larvae crawl immediately into the fur of the host. The larvae enter the host through natural body openings.127 The first instars remain within the nasopharyngeal region for 6
8 days.128,129 Then they migrate to the trachea through the tracheal wall into the thoracic cavity, through the diaphragm into the abdominal cavity, and finally to the SC sites in the inguinal and thoracic areas.130 The larvae molts to the second instar and to the third

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instar in B17
19 days. The larva enlarges an exit pore during the last 4
5 days of development and backs out of the warble. The development time from infection to when the larva exits the warble varies from 3 to 8 weeks. The third instar burrow into the soil and pupates. The adult fly emerges anywhere from a month to several years.

Rearing method(s) On host—Peromyscus spp.—Cuterebra fontinella A laboratory colony of the rodent botfly, Cuterebra fontinella, can be maintained on its natural hosts, Peromyscus lucopis, the white-footed mouse or Peromyscus maniculatus, the deer mouse. Peromyscus spp. are reared in the laboratory. One male and one female mouse are mated in a colony cage. The gestation period is B21 days and mouse pups are weaned at 30 days. A male mouse is left in the cage with the female for several days after birth and then removed. The same male is placed back with the female after weaning is completed. Only B10% of the Peromyscus spp. colony end up as breeding pairs. To begin a Cuterebra spp. colony, B15
20 mice (20 g) are used per week. Five Cuterebra spp. larvae are placed in the eye, with a brush, of the mouse. Pupation occurs B25 days later. Consequently, 18 days PI with newly hatched larvae, the mice are placed in collection cages (0.5 in. hail screen) so pupae will fall through. Mice are used with 2-week rest intervals between pupae collection. The ocular inoculation method usually yields an infection rate of 80%
90% with the first infection, but the percentage of infections plateaus at B60% after the 2
3 infections. Pupae are collected and placed in paper cups (half-pint ice cream carton with the top of lid cut off) with a nylon mesh over the lid. Pupae (B20) are placed in the carton lined with wood shavings. Pupae are place in an alternating temperature
photoperiod box (dark for 9 h at 15 C and light for 15 h at 27 C) that is important for emergence of flies. A two-stage bulb thermostat is used to control the temperature and light cycle. Adult flies will emerge from pupae B30 days after pupae are collected. Flies are sexed (females have a wider space between the eyes than males and males have claspers), separated, and placed into Petri dishes lined with filter papers. The sexed adult flies are left for 24 h at RT, which allows the fly to void emergence materials, before placing two of the same sex/dish into an incubator at 16 C. This temperature keeps the flies inactive and minimizes wing damage. In vitro method(s) No in vitro bioassays were found in the literature.

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In vivo method(s) A model has been developed to screen systemic compounds in mice experimentally infested with a rodent botfly.131 Basically, laboratory white mice are experimentally infested with first instars of Cuterebra spp. on TD 0. The infested mice in treatment and control groups are treated PO with the compound or placebo solutions on TD 2. Control and treated mice are examined on TD 7 for larvae that formed breathing holes in the skin and oral LD50 values of the test compounds are determined. The systemic activities of the compounds tested in the Cuterebra mouse model were similar to the activities of the same compounds tested against larval Hypoderma in cattle.

Dermatobia hominis Linnaeus Jr, 1781—human bot fly Biology and life cycle The human botfly is a member of the family Oestridae, subfamily Cuterebrinae: human bots, exclusively distributed in the Nearctic and Neotropical regions. This species is native to the Americas from Southeastern Mexico (beginning in central Veracruz) to northern Argentina, Chile, and Uruguay. This botfly occurs in Mexico, Central America, and South America132 where it is economically damaging to the cattle industry. Countries with the highest infection rates in travelers are Belize, Bolivia, and Brazil.133 This dipteran fly has a most unusual life cycle. The adult fly inhabits the forests of Mexico, Central America, and South America. Adult D. hominis are 12
15 mm long, about the size of a bumblebee, and has a short life span (1
9 days). It has a blue-gray thorax, a metallic blue abdomen, and yellow-orange legs.134 The adult has no functional mouthparts (as is true of other oestrid flies) and takes no nourishment.135 Food stored during the larval stage provides the adults with nourishment. The principal stimuli leading to the meeting of the sexes probably are visual. The flies remained in copula an average of 9 min (range 4
21 min), and both sexes mated repeatedly. The flies reached sexual maturity at B1.5
4 h after emergence.136 The female fly does not deposit her eggs directly on the host. Instead, she captures another dipteran fly, usually a bloodfeeder (e.g., mosquitos). Dermatobia flies have been shown to be vectored by over 40 species of mosquitoes and muscoid flies, as well as one species of tick.137 The female captures the mosquito and attaches its eggs to its body, then releases it. Either the eggs hatch while the mosquito is feeding and the larvae use the mosquito bite area as the entry point, or the eggs simply drop off the muscoid fly when it lands on the skin. The eggs are attached to the carrier in such a manner that when contact is made with the prospective definitive host, the anterior end of the egg is

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directed downward. An operculum forms on this end of the egg, through which the larva emerges while the insect feeds on warm-blooded hosts. The larvae penetrate the skin of the animal within a few min of hatching and remain in the SC tissue for 4
18 weeks. During this period the larvae grow within warbles with breathing holes. D. hominis does not meander through the subcutis. The mature larva at 2
3 months of age is 1.8
2.4 cm in length. It has a definite club shape and can be identified by rows of anteriorly directed spines on its anterior segments. The larval stage also possesses the caudal spiracles that protrude through the host’s skin to the exterior to guarantee an adequate air supply. The larva of D. hominis is narrow and tubular at its posterior extremity and somewhat flask-shaped anteriorly.138 The presence of the superficially positioned swelling with a central opening may lead to a tentative diagnosis of myiasis due to D. hominis. A definitive diagnosis can be made only after extraction and identification of the typical larva. The to´rsalo matures to its most advanced larval stage in the body of the host.139 Development in the host requires 35
70 days. The larva then drops to the ground where it enters the soil for pupation. After the pupal period, which lasts 4
11 weeks, the flies emerge as adults. The entire life cycle takes 90
120 days.139,140

Rearing method(s) The laboratory rat is a suitable model for studies of D. hominis infestation.141 Two days after rats, Rattus norvegicus, were experimentally infested with D. hominis, the first instars were located deep in the dermis. A small warble was formed after the fourth day, increasing in size until the seventh day, when the first instar molted to the second instar. After 18
20 days the second instar molted to the third instar. The third instar usually left the host after 30 days. D. hominis larvae were cultured in a semidefined liquid medium.142 First instars grew well up to 44 days; 29.1% molted in a mean period of 8.62 days. Two larvae reached the third instar but lived only 1 and 18 days, respectively, after the second molt. The increase in size, measured in 4 larvae, was B10-fold. Second and third instars, obtained from the skin of cattle, survived and grew in the medium for up to 2 months; 39.0% of the second instars molted, while 77.3% of the third instars pupated, and some produced flies when transferred to sand after 14.84 6 10.08 days in the culture medium. Some maturation factor, obtained from the skin, may be necessary for the larvae to grow satisfactorily and to complete the full parasitic cycle in vitro. Heavy damage to the cattle industry in tropical Latin America by D. hominis provided economic justification for research on the sterile-male approach to eradication of this species. A method of colonizing D. hominis

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by the use of a natural host for rearing the larvae was developed, along with techniques for collecting the full-grown larvae, holding the pupae and adults, inducing mating and oviposition in small cages, and infesting the hosts.143 Naturally infested cattle from the field were stabled and larvae dropped from cysts in the early morning hours. Larvae were placed on humid soil (sandy clay loam with 20%
30% water w/w) in cylindrical cardboard box 9 cm in diameter and 9 cm high. Pupae were left undisturbed for 10 days and then removed from the soil, washed, air-dried, and transferred into cardboard boxes held at 24 C
25 C and 70% RH. The pupal period averaged B34 days. Cardboard boxes of pupae were placed in in 30 3 30 3 30 cm cages in an incubator at 24 C
25 C and 70% RH. Emerged flies were collected, separated by sex and transferred to smaller cages. Both male and females sometimes reached sexual maturity as early as 3 h after emergence. When adult flies were 1 day old, equal numbers of females and males (4
10 of each sex) were confined to cages incubated at 28 C and 80% RH with a 100 W daylight bulb suspended over the top of the cage. Oviposition occurred when female flies were 2 days old. Carrier flies (M. domestica), 20 houseflies/female D. hominis were placed in the cage. No water or food was provided and the houseflies were dead after 24 h. Female D. hominis laid as many as 600 eggs with an average of 250 eggs. The eggs were glued to the ventrolateral parts of the abdomen of the carrier flies in masses of 10
80 eggs, averaging 35 eggs. The eggs masses were removed with forceps and placed in Petri dishes with a pad of cotton soaked with water, which yield nearly 100% RH. Eggs hatched between 5 and 15 days after oviposition at 28 C. Hatching could be induced by placing the eggs directly on the host, by incubating them for 3 min at 36 C, or simply by blowing over them. Newly hatched larvae were placed on the skin of rabbit, goat and cattle (1500 larvae) hosts. The duration of the larval stage ranged from 35 to 128 days. The host was tabled when the larvae appeared full grown and the entire process repeated.

In vitro method(s) While no in vitro tests were found in the literature, the availability of the life cycle stages of D. hominis by laboratory rearing methods provides suitable material for the development of in vitro parasiticide assays. In vivo method(s) Parasiticide efficacy testing against D. hominis on ruminants should be designed and conducted in accordance with the WAAVP guidelines for evaluating the efficacy of ectoparasiticides against myiasis causing parasites of ruminants (see Appendix A of Ref. [94]). In a therapeutic trial, 12 calves were infested along the dorsal line with 25 first instars of recent field isolates of D. hominis, but in one calf nodules

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did not develop.144 Twenty-four days later animals were allocated to two groups on the basis of the number of parasite nodules present. Six calves were treated with doramectin, and five received saline solution. Animals were examined daily for 11 days PT, and the number of nodules mapped and recorded. Larvae that completed development were collected and incubated to evaluate viability. In the persistent efficacy study, 24 calves were allocated to six groups (T1
T6) of four animals each. On the day of treatment, three groups (T1, T3, and T5) were treated with saline and three groups (T2, T4, and T6) with doramectin. At 21, 28, and 35 days PT, 25 first instar D. hominis were seeded along the dorsal line of each calf of T1 and T2, T3 and T4, and T5 and T6, respectively. Animals were examined daily for 18 days, and the number of nodules mapped and recorded 6, 12, and 18 days PI. In the therapeutic efficacy study, parasitic nodules in the doramectin-treated animals were reduced by 74% (P , .05) at 48 h PT, and efficacy reached 100% at 6 days PT. In the saline-treated calves, parasitic larvae remained inside the nodules and completed their normal larval development. Sixty-five percent of the larvae that emerged from the nodules of control animals developed into adult flies. The persistent efficacy of a single injection of doramectin extended beyond 35 days, and no parasitic nodules developed in the treated calves at any time. By contrast, all saline-treated calves developed nodules with presence of viable larvae recorded at 6, 12, and 18 days PI.

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10. Haslelton AT, Acevedo A, Kuruvilla J, Werner E, Kieman J, Dhar P. Repellency of α-pinene against the house fly, Musca domestica. Phytochemistry 2015;117:469
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Chapter 3d

Arthropoda, Phthiraptera, Anoplura Larry R. Cruthers, MS, PhD1 and Douglas D. Colwell, PhD, FRES, Assoc. EVPC2 1 LCruthers Consulting, Chesapeake, VA, United States, 2Agriculture and Agri-Food Canada, Lethbridge, AB, Canada

Arthropoda Phthiraptera Anoplura: sucking lice

Biology and life cycle Sucking lice, like all lice species, are wingless, dorsoventrally flattened and permanent ectoparasites of mammals. Sucking lice have piercing mouthparts adapted for sucking the tissue fluids and the blood of the host. They infest a wide range of domestic livestock as well as humans. They are a source of constant irritation, itching, rubbing, and biting of the skin or hair coat and are closely adapted to their hosts and are completely dependent upon them for survival. The entire life cycle of lice occurs on the coat of the host animal; no stage of the lice life cycle occurs off the host in the host’s environment. In general, there are three developmental stages in the life cycle (incomplete metamorphosis) of the louse. The lice egg (nit) stage of the life cycle hatches in 7
14 days, depending on the species of lice and on environmental conditions. The three nymph stages of the lice life cycle grow and undergo their molts over B9
22 days, depending on the lice species. The adult lice usually live for 2
3 weeks on the fur or hair of the host animal. They will lay their eggs during this time period. Specifically, the life cycles of several important genera and species are described next.

Linognathus setosus Olfers, 1816—sucking louse of dogs This dog louse is distributed in tropical and subtropical regions across most countries of the world. These lice have pincer-like tarsal claws for clinging to hairs of the host. The adult lice reach up to 2 mm in length with the first pair of legs smaller than the other two pairs. The body is covered with hairs and lacks pigmented plates on the abdomen. Eggs can be seen as white specks attached to the hairs of the host. The average time from egg to egg is 3
4 weeks. The eggs hatch and develop into nymphs, which are tiny, increasing in size as they go through three further molts before reaching the

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adult stage. These lice are frequently found under mats of hair and around the ears and body openings. In heavy infestations, they cover the entire body. They are spread by direct body contact with infested animals. They can be a vector of the dog tapeworm, Dipylidium caninum.

Linognathus vituli Linnaeus, 1758—long-nosed or blue cattle louse This cattle louse, which occurs worldwide, gets its common name from the narrow, long head in front of the large, five-segmented antennae. This louse, B2.5 mm long, is morphologically similar to Linognathus setosus and can be found all over the body, preferably around the neck, but also feeding on the head, brisket, back, and inner thighs. In heavy infestations, they cover the entire body. The heaviest infestations occur in late winter and early spring with very light infestations in the summer. They are spread by direct body contact with infested animals and the life cycle is 2
3 weeks. Linognathus vituli can serve as a mechanical vector for Anaplasma marginale, the causative agent of bovine anaplasmosis.

Linognathus ovillus Neurnann, 1907—face louse of sheep This sheep louse occurs worldwide and can be found all over the body of sheep, except the extremities, such as the lower parts of the limbs. When their numbers are low, they are found principally on the hair-covered parts, particularly, the face. The eggs are glued to wool fibers or hairs. Various breeds of sheep are reported to be more, or less, susceptible to lice infestation. This sheep louse is not known to be a vector of any disease agents.

Linognathus pedalis Osborn, 1896—foot louse of sheep This sheep louse is worldwide in distribution and is found on the legs of sheep and also the scrotum area of rams. This louse is not considered very injurious because feeding occurs on the hairier parts of the sheep’s body, and the animal exhibits little discomfort. Younger animals are more heavily infested and rams seem to be preferred. Light infestations commonly occur as clusters of lice around the accessory digits. The life cycle takes B30 days at a minimum. They are spread by direct body contact with infested sheep or from an infested environment. This louse is not known to be a vector of any disease agents.

Haematopinus eurysternus Nitzsch, 1818—short-nosed cattle louse This cattle louse species is worldwide in distribution, gets its common name from the narrow, short portion of the head in front of the large, fivesegmented antennae. Each segment of the abdomen is distinct, narrowing to a point laterally. When engorged with blood, the lice appear as oval,

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grayish-to-bluish bodies moving sluggishly in the coat of the host. It is the largest among cattle lice (3
5 mm long). It is found not only on the head, especially around the horns, the eyes, and the ears, but also on the neck, shoulders, and the base of the tail. It is considered by some to be the most damaging louse for cattle. The average time from egg to egg is 3
4 weeks. They are spread by direct body contact with infested animals. These cattle lice are not known vectors of any disease agents.

Haematopinus quadripertusus Fahrenhotz, 1916—cattle tail or tail switch louse This cattle louse is worldwide in distribution but occurs primarily in the tropics and subtropics. Adult lice are normally found on the last 7 cm or so of the tail and deposit eggs on the hairs of the brush, occasionally when heavy infestations occur, adults and eggs may also be found in the ears. Under optimal conditions, the cattle tail lice will complete its life cycle (egg to egg) in as few as 25 days. They are spread by direct body contact with infested animals or by the contact of areas where animals have scratched or rubbed. It has been reported that this louse can survive up to 40 days off the host, although most individuals will die in a few days. These cattle lice are not known vectors of any disease agents.

Haematopinus suis Linnaeus, 1758—sucking louse of swine Swine lice are the largest of the Anopluran sucking lice, measuring up to 0.5 cm from the tip of the head to the end of the abdomen. It is distributed worldwide wherever domesticated swine are found. It can be seen with the naked eye, and the only species of lice found on swine. Their very large tarsi and claws are used to grasp the host’s hair. Each segment of the abdomen as dark-pigmented lateral plates. The average time from egg to egg is 3
4 weeks. The lice move sluggishly and cover the entire body in heavy infestations. They are spread by direct body contact with infested animals. Swine lice can transmit small pox, eperythrozoonosis, a rickettsial disease caused by Eperythrozoon suis, and hog cholera.

Haematopinus asini Linnaeus, 1755—sucking louse of equines This louse is the largest species among horse lice (3
4 mm long) and is a blood-sucking lice found worldwide infesting horses, mules, and donkeys. It is found mainly on the head, neck, back, brisket, and between the legs. The average time from egg to egg is 3
4 weeks. The heaviest infestations occur in late winter and early spring. Transmission occurs by direct body contact with infested animals and occasionally through contaminated equipment. Horse lice are vectors of equine infectious anemia.

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Solenopotes capillatus Enderlein, 1904—little blue cattle louse or tubercle-bearing louse This louse of cattle is a small louse (1
1.5 mm in length) which gets its latter common name from the prominent tubercles that project from the sides of each abdominal segment. It is found throughout the world but generally restricted to areas with domesticated livestock. The average time from egg to egg is 3
4 weeks. The female louse lays 1
2 eggs/day, each attached to a hair. Often the hair is bent, a feature not observed with other cattle lice. They feed on the head, primarily the face and jaw regions, with sporadic occurrences on other body regions. Clusters of lice around the eyes cause heavily infested animals to look as if they were wearing glasses. The lice are spread through direct contact between cattle during mating, nursing, or other interactions, such as herding. These cattle lice are not vectors of any known disease agents.

Polyplax serratus Burmeister, 1839—mouse louse; Polyplax spinulosa Burmeister, 1839—rat louse The spiny rat louse is worldwide in distribution and commonly infests its type host, the brown rat (Rattus norvegicus) as well as other related rat species. It is occasionally found in other rodents, such as the marsh rice rat (Oryzomys palustris). The louse causes hair loss and itching. They spend their entire life cycle of B14
21 days, from egg to nymph to adult, on the host. These lice can cause anemia, but even more importantly for rats, they may transmit the blood parasite Hemobartonella muris, which is a rickettsial blood parasite similar to tick fever. They may also transmit Rickettsia typhi between rats and are known vectors of Mycoplasma spp.

Pediculus humanus humanus Linnaeus, 1758—human body louse This louse, which is sometimes called Pediculus humanus corporis, is commonly called the clothing louse, which during WWI became known as the “cootie” and during WWII was properly called “mechanized dandruff.” It infests humans throughout the world with a condition known as pediculosis. The skin of persons who continuously harbor lice becomes hardened and deeply pigmented, a condition designated as vagabond’s disease or morbus errorum. Body lice are vectors for the transmission of epidemic typhus, trench fever, and relapsing fever. This louse is indistinguishable from the head louse but will interbreed only under laboratory conditions. In their natural state, they infest different body regions and do not usually meet on the host. Body lice attach their eggs to clothes whereas head lice attach their eggs to the base of hairs. The eggs are generally easy to see in the seams of an infested person’s clothing, particularly around the waistline, under

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armpits, or even in body hair. The eggs hatch in 1
2 weeks, nymphs mature into adults about 9
12 days after hatching, and the adult louse lives for about 1 month, in which time the female lays upward of 300 eggs. The lice are spread through direct contact with infested persons.

Pediculus humanus capitis De Geer, 1767—human head louse This louse, worldwide in distribution, resides close to the scalp of humans to maintain its body temperature. The eggs are difficult to see and are often confused for dandruff or hair spray droplets. The nits are cemented by the females at the base of the hair shaft nearest the scalp, and usually take about 1 week to hatch. Viable eggs are usually located within 6 mm of the scalp. After hatching, the nymphs become adults about 1 week after hatching. The adult louse is about the size of a sesame seed and live up to 30 days, feeding several times daily. The head louse is considered to be a potential vector of louse-borne diseases in both laboratory studies and field investigations. However, it is considered to be nondisease transmitting louse. The majority of infestations are asymptomatic but when symptoms occur, it is usually itching caused by an allergic reaction to louse saliva. Secondary bacterial infections may be a complication.

Pthirus pubis Linnaeus, 1758—human pubic or crab louse This louse is easily recognized by its crab-like appearance. It primarily infests the pubic or genital areas of humans but also the armpits, mustache, beard, eyebrows, or eyelashes. Eggs take B6
10 days to hatch, and hatched nymphs develop to adults B2
3 weeks later. As with all lice nymphs, they must feed on blood. Pubic lice are spread through sexual contact and are most common in adults. Occasionally, pubic lice may be spread by close personal contact or contact with articles, such as clothing, bed linens, or towels, that have been used by an infested person. A common misconception is that pubic lice are spread easily by sitting on a toilet seat. This would be extremely rare because lice cannot live long away from a warm human body and their legs are not designed to hold onto or walk on smooth surfaces, such as toilet seats. Pubic lice are not known to transmit any disease agents.

Rearing method(s) Pediculus humanus humanus (corporis) de Geer, 1778 In many countries, body lice, although vectors of several disease-causing organisms, are of only minor importance. Unlike its relative, human head lice are still an important public pest. Although human lice can be treated effectively with topical pediculicides, resistance has developed and become a

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concern. Therefore the evaluation of products and active substances against human lice is of public health importance. Rearing of both the body and head louse had for many years been practical only on humans1,2 until the body louse had adapted itself to feeding on the only substitute host—the rabbit.3,4 During feeding, rabbits must lie on their backs and have their legs fixed to a special table for about 1 h since lice were only able to suck blood on the shaved rabbit abdomen. This feeding technique did not correspond to the natural feeding performance of lice and was not compatible with animal welfare. Thus the need arose for establishing in vitro feeding techniques for maintenance and rearing of P. h. corporis. Human head lice (Pediculus humanus capitis) have been shown to be difficult to rear and maintain the laboratory in large numbers,5 but the human body lice have been successfully adapted to rabbit hosts as previously cited. Since differences in the physiology of body and head lice are minimal, the rearing laboratory strains of rabbit-adapted body lice are an important way for efficacy testing of pediculicides, which is of increasing importance because of the developing resistance problem. Fuller et al.6 and Haddon7,8 fed P. h. corporis through membranes during a 14-day and 48-day period, using chicken skin or Gutta membranes (similar to natural rubber) and defibrinated human blood or defibrinated hemolyzed human blood, respectively. A study by Lauer and Sonenshine9 involving defibrinated rabbit blood revealed that the blood uptake by the body louse was suboptimal. Mumcuoglu and Galus10 observed sucking rates of the body louse after offering different blood fractions and their components through silicone-Parafilm membranes. Despite these promising feeding studies, a continuous breeding of the human body louse in the laboratory using a membrane feeding technique had yet to be established. The objectives of a study by Habenank et al.11 were threefold: (1) confirm the membrane feeding method for P. h. corporis (feeding system, membrane, necessity of olfactory stimulation), (2) evaluate feeding media continuously available for nutrition of a lice colony, and (3) establish a longterm in vitro breeding of P. h. corporis. The body lice used for their feeding experiments were derived from a body louse strain fed on rabbits 4
5 times/ week for 15
20 min. Between daily feeding times, the parasites were stored at 32 C and 60% RH, and at 25 C and 60% RH on nonfeeding days. Adult and first instar lice were used for the in vitro feeding experiments. Lice of the same age were randomly divided into equal experimental groups. The last feeding of adult lice on rabbits had been about 24 h before, with first instars never having been fed (5teneral instars). After the daily feedings, lice were stored in darkness in an incubator at 31 C and 70% RH. On days without nutrition, lice were kept at 19 C
22 C and B70%
80% RH.

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Feeding system11 for Pediculus humanus corporis Parafilm M was used as a membrane over the nutrition medium. The figured glass rings were small (42 mm diameter, 40 mm height; made by a glass blower for this purpose) to give Parafilm M more stability when stretched over the ring. Glass dishes had a diameter of 50 mm. The feeding devices were placed on a hot plate in a sterile flow (Gelaire Flow Laboratories, model HF 48) to prevent nutrition media from being contaminated, thus avoiding the addition of antibiotics or fungicides. A Sartorius laboratory balance, precision 0.1 mg, was used to determine louse weight. Nutrition media: Fresh human blood, stabilized with sodium heparin, was stored at 4 C until used. Porcine blood was taken from the jugular vein of slaughter animals and treated in the same way as fresh human blood. Preserved human blood was obtained from a blood bank as three separate, stabilized units: DFP, EcC, and BC. All units were obtained for experimental lice feedings only after their transfusion medical expiration date. For experimental use, the DFP, EcC, and BC were reunited into PHB stored at 4 C or 227 C. For feedings, 2
3 mL of nutrition media was filled into each dish. Feeding experiments: The feeding system was restricted to the required minimum: thin membrane (Parafilm M), nutrition medium, temperature at the hot plate similar to the body temperature of the host body (38 C), no olfactory stimulation, no magnetic blood mixing, and daylight. Four samples of adults were exposed to the in vitro system for 6 h. Fresh human blood was offered as optimal nutrition medium. In following trials, other promising nutritional sources (porcine blood, preserved whole blood) were examined in comparison to fresh human blood controls (exposure 1
2 h). The behavior of larvae in the in vitro system was also tested. First instars were exposed to porcine blood and preserved whole blood for 1 h. To establish the nutritive quality of short-time stored PHB, blood of the same sample was offered to adults and larvae on TD 1 of preparation and on TD 14 of storage at 4 C or 227 C. Each day, live lice were counted and dead specimens removed. The weight before and after nutrition, measured per lice sample (including lice specimens without blood uptake), served to calculate the mean individual louse weight. To establish a membrane fed colony, P. h. corporis were regularly exposed to superimposed human preserved blood 4
7 times/weeks for 60
90 min and only in exception (i.e., after storing at 19 C
22 C) up to 120 min. Colony feeding started with groups of 200 first instars without previous blood uptake. PHB was used immediately after preparation or storage 1
14 days at 4 C or at 227 C. Two samples were offered PHB, stored at 4 C and two samples PHB, frozen at 227 C. The initial feeding experiment with P. h. corporis confirmed the earlier investigations previously cited, that lice can be fed well through membranes.

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The artificial membrane Parafilm M may be used for feeding all stages of P. h. corporis. Attraction of the lice to temperature was sufficient to induce piercing of the membranes. Lice tasted and engorged the offered nutrition media. This study showed that in vitro feedings of body lice are possible in a feeding system reduced to a required minimum. In addition, the use of a sterile flow avoided contamination of the blood media and avoided the use of antibiotics often practiced during in vitro feeding of other hematophagic arthropods. Continued in vitro maintenance of lice cannot depend on frequent blood donations by human volunteers. Daily lice feedings require a stock of nutrition media. Blood of animals (donor or slaughter animals) and units of human blood are continuously available. The pig seemed to be the most promising species among animals. Porcine blood induced high sucking rates in all observed lice stages but caused high mortality rates. In fact, all lice died during a course of repeated feedings within 1 week; possibly, the hostspecific body lice are not able to digest some porcine blood components. Therefore porcine blood cannot serve as a nutrition medium for a continuous breeding of human lice. The conclusion of this study was that PHB offered a return to human blood as natural nutrition medium of the body louse. Despite increased developmental times, a continuous maintenance of P. h. corporis by membrane feeding on PHB is possible, as demonstrated over nine generations. These preliminary results indicate that in vitro feeding could be an alternative method of breeding and, possibly, mass rearing of the human body louse. Further studies are required to optimize the composition of the PHB. Membrane feeding saves laboratory animals and is considered to be a contribution to animal welfare. Artificial feedings using standardized nutrition media are recommended for medical or biological studies on the vector role of lice, for the examination of the parasite itself, and for qualitative and quantitative insecticide testing. Schrader et al.12 reported on the survival and reproduction of a laboratory strain of body lice at different ambient temperatures. These authors used the only laboratory strain in Germany, one initially established from a strain at the USDA Research Laboratory in Orlando, FL, United States. Even though this strain could be maintained in the laboratory, data on temperature physiology were scarce. Body lice complete their entire life cycle on or near their endothermic hosts and are likely adapted to specific temperature criteria crucial for their survival and reproduction. In order to optimize their lice rearing conditions and understanding of body louse temperature physiology, the authors evaluated the influence of different temperature regimens on body lice. The emphasis of their study was on the influence of temperature on adult louse survival after their last blood meal and additionally, on oviposition and egg hatch rates.

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For routine lice rearing, the lice were fed on rabbits, as substitute hosts, 4
5 times/week for 15 min. Between feedings, the lice were held in glass Petri dishes on a 5 3 5 cm piece of corduroy cloth at 32 C and 45%
55% RH on feeding days and at an ambient temperature of 25 C and 45%
55% RH on nonfeeding days. When fed 4 times/week, the lice developed to adults after 11 (10
12) feeding episodes 20
21 days after hatching from eggs. The lice deposit their eggs on the corduroy cloth and remain in the incubator at 32 C until larval hatch after 8 (6
9) days. The routine rearing conditions were then modified according to the experimental objectives: the main parameter that was changed being ambient temperature. These studies showed that unfed adult survival was negatively correlated with increasing temperature, with the longest survival (15 days) at an ambient temperature of 16 C and the shortest survival at 32 C (4
5 days). In contrast to this, oviposition rates and egg hatch rates positively correlated with increase of temperature (1.15 eggs/day/female at 22 C and 3.52 eggs/day/female at 32 C). Experiments evaluating the influence of different temperature regimens revealed that eggs must first be incubated for 5 days after oviposition at .25 C. Eggs which were first incubated at 25 C and then transferred to 32 C had high hatch rates, indicating that the early development of eggs after oviposition is less temperature sensitive than later development. The lower RH conditions in these studies (45%
55%) may account for the longer adult survivability times than studies conducted by others at 90% RH, as well as the use of different strains of body lice. It is thought that high humidity rates can be disadvantageous for body lice since their copious feces, which is normally dry, becomes damp and sticky. Lice can become coated with their own excrement and get stuck on the clothes where they live.

Pediculus humanus capitis de Geer, 1778 Takano-Lee et al.5 were the first to demonstrate that head lice could be reared successfully in vitro through a complete life cycle. Filter paper assays carried out in Petri dishes are the most common method to determine levels of topical insecticide resistance in head lice.13 These authors reasoned that although such bioassays are informative and provide comparable, standardized results, filter paper bioassay data may not accurately represent in vivo pediculicide treatment and are inappropriate for systemic pesticide testing for the following reasons: insecticide-treated lice are likely to continue blood feeding, and blood ingestion may alter the availability of the insecticide or the physiology of the louse, thereby modifying subsequent mortality. The performance of similar bioassays on human hosts was impossible and unethical because mortality effects may not be observed for 24 h and more and the observation of lice on human hosts is extremely difficult. They concluded that the best alternative was to observe

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insecticide-treated head lice within an in vitro setting that allowed lice the opportunity to blood feed. There currently was no such in vitro setting or rearing mechanism available. Maintenance of lice on a human host is irritating and time consuming because head lice feed 4
5 times daily. The rearing methods developed to date were for many different hematophagous insects, but most of these insects were temporary parasites and only feed on blood periodically. The ability to maintain blood-sucking arthropods without a live host would be a great convenience and would eliminate the need for human or animal use protocols and approval by institutional animal use review boards. The system developed by Lee et al.13 (later identified as the “manual” in vitro rearing system) only used newly hatched first instars (teneral instars) in their feeding experiments, because the older stages were unlikely to feed in vitro unless “entrained” from a teneral state. Teneral first instar lice were placed on human hair tufts on the upper side of a Parafilm-silicone membrane-covered feeders, which were immersed bottom-side down within a vessel containing warmed human blood, purchased from a local blood bank as separate units of RBCs and plasma from the same donor. These blood products were recombined at a ratio of 1:1 before use. Relative to lice reared on a human host, in vitro reared lice required a significantly longer time (10%
20%) to molt and survived a significantly shorter time as adults (30%
50%). In vitro reared lice may have required more time to develop, because of smaller ingested blood meals through artificial membranes, or reduced nutritive value of the stored blood bank blood that was used in the study. Lice spent a significantly greater portion of time searching in the in vitro apparatus than on a host, but the proportion of time spent feeding did not differ. These behaviors were separated into three major categories: (1) feeding or ingestion of blood, (2) resting or lack of feeding and walking activity, and (3) searching or ambulation. Time spent in each behavioral category was determined. With the success of an “automated” system described next, the specifics of design and detailed study results of the manual in vitro system are not described. Interested readers can refer to the published article for details regarding this manual rearing system. Since the “manual” in vitro rearing system required daily human maintenance, a follow-up study was conducted by Takano-Lee et al.14 to develop an “automated” in vitro system that minimized human labor. Once again, only teneral first instars were initially placed in the “automated” in vitro rearing system, because, as these authors had previously observed, this was the stage that fed most successfully through a siliconereinforced Parafilm membrane. The “automatic” feeding system, fabricated to maintain large-scale rearing of head lice, consisted of three distinct parts: the fluid-release system, the feeder system, and the fluid-drainage system (Fig. 3d.1). This system was large in size and contamination was of concern;

FIGURE 3D.1 Automated in vitro rearing system by Takano-Lee et al.14 (A) fluid-release system, (B) feeder system, (C) drainage system. Reprinted with permission from publisher.

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therefore an “improved” automated system was developed, several years later by these authors, and it is that system that will be discussed in more detail. Readers are recommended to review the 2003 publication by Takano-Lee et al.14 for the construction details of their initial automated feeding apparatus. Yoon et al.15 described an “improved” in vitro rearing system that reduced the size and contamination concerns of the “automated” in vitro feeding system14, using modifications mainly to the “manual” in vitro rearing system previously referenced by Lee et al.13 This practical and easily maintained in vitro rearing system, in addition to allowing for the large-scale rearing of both pediculicide-susceptible and resistant strains of human head lice, also allowed for the development of a mortality bioassay using over-the-counter pediculicidal formulation-treated hair tufts in conjunction with the stretched, silicone-reinforced Parafilm M membrane for “semiclinical validation” of product efficacy and resistance status of maintained louse strains.

In vitro rearing system As mentioned, an improved in vitro rearing system was developed based on modifications to the manual prototype and maintained under semisterile conditions (Fig. 3d.2). The lid of a 50 mL polypropylene conical tube served as the blood feeding reservoir. Human blood (4 mL), which was reconstituted from fresh

FIGURE 3D.2 Assembly of the in vitro rearing system by Yoon et al.15 Reprinted with permission from publisher.

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RBCs (A1, one part), plasma (A1 one part), and a penicillin plus streptomycin antibiotic mixture (0.01 part of a 1000 U penicillin plus 1 mg streptomycin plus 0.9 mg NaCl/mL stock solution in distilled DI water), was added to the blood feeding reservoir and mixed with a small magnetic stir bar. Each blood feeding reservoir was placed on a water-circulation plate that was situated on the top of a multipoint magnetic stirrer. Water was cycled through the circulation plate at 31 6 1 C. An inverted and truncated (30 3 40
45 mm, diameter 3 height) 50 mL polypropylene conical tube was used as a rearing vessel. The screw top end of each rearing vessel was covered by a silicone-reinforced Parafilm M membrane sandwich. The membrane sandwich was formed by adding B55 mg of silicone (aquarium sealant) between two Parafilm M squares (25 3 25 mm). Silicone was evenly distributed by gently rolling a wooden applicator stick over the Parafilm M squares that formed the sandwich. The membrane sandwich was stretched (B100 3 100 mm final dimensions) over warm air, laid firmly over the screw top end of a rearing vessel, and permitted to dry overnight. In order to assemble a rearing unit, the rearing vessel (membrane side down) is slowly immersed into the blood feeding reservoir at an angle and gently rotated around its axis to avoid air bubble formation underneath the membrane. Because the cap used as the blood feeding tube is manufactured to fit the tube serving as the rearing vessel, a tight fit is ensured. The interface where the blood feeding reservoir and rearing vessel contacted each other was wrapped with Parafilm M to prevent water evaporation from the blood and to protect from bacterial and fungal contamination. A semicircle piece (B26 mm in diameter) of wide gauze, which still allowed the lice to feed, was placed on the membrane surface to reduce accidental leakage caused by trafficking of lice. Hair tufts (200
300 hair strands, B23 mm in length) were prepared using human hair and situated on the wide-mesh gauze to mimic a human scalp. Blood, rearing vessels, and blood feeding reservoirs were changed Bevery 72 h. An incubator (31 6 1 C, 70%
80% RH) maintained the eggs harvested from the rearing vessel until hatched. Disposable sterile plastic wares, examination gloves, laboratory coat, safety goggles, and a face mask were used to maintain laboratory biosafety level 2. Surface areas of blood feeding reservoirs and rearing vessels were cleaned with 10% (v/v) bleach and 70% (v/v) ethanol to reduce bacterial and fungal contamination. Developmental characteristics for the colonized louse reared on the improved in vitro rearing system were recorded daily. Values for development time (days) and percent survival at the first, second, and third ecdysis were determined. Adult longevity (days) between the final ecdysis and mortality of male and female lice were also determined. Mating pairs were selected and transferred to a new in vitro unit to record female fecundity. Laid eggs were counted and transferred daily to an incubator and mean daily

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number of eggs produced per female was calculated until all the females died. Egg hatch was determined for the eggs collected. The improved in vitro rearing system provided an increased membrane surface area (706.5 mm2) in each rearing vessel compared to that (122.7 mm2) of the previous manual and automated rearing system, maintained 100
150 first or second instars per rearing vessel, and B50 third instars or adults per rearing vessel. This allowed 1125
1800 eggs/day/stirrer to be produced in 15 rearing units per multipoint magnetic stirrer with 25 females and males in each rearing unit at the fecundity level of 3.0
4.8 eggs/day. Other specific improvements were as follows: (1) multiple hair tufts (3
4) arranged on a semicircle patch of wide-mesh gauze were added to the rearing vessel to mimic a human scalp, resulting in widespread distribution of lice on the feeding membrane surface, (2) the use of a circulation bath and water-circulation plate to regulate rearing temperature greatly reduced the amount of time and effort required to change blood and the disposable rearing units since the water-circulating tubing around each rearing unit was eliminated, and (3) fabrication of the blood feeding reservoir form the lid of a 50 mL polypropylene conical tube insured a tight fit of the reservoir to the rearing vessel, which virtually eliminated evaporation of water from the blood, and utilizing a semisterile environment along with sterile and disposable materials, substantially reduced bacterial and fungal contaminations. The frequency of blood changes decreased with the improved system from every 12
24 h to every B72 h with these modifications. No laboratory rearing methods of sucking lice of other than human species were found in the literature or not known to the authors.

In vitro method(s) Filter (chromatography) paper or cloth disk assay Pediculus capitis adults and nymphs collected from the heads of children were placed within 2 h of collection into 6-cm stainless steel baskets with nylon net bottoms into separate glass Petri dishes lined with Whatman No. 1 filter paper (7.0 cm day) on the bottom and were used for the bioassay of pediculicidal activity.16 The baskets containing adults and nymphs that were distributed in Petri dishes lined with the Whatman No. 1 filter paper on the bottom were treated with 600 μL of a test product at different concentrations and combinations, for 15 min at 65 6 5% RH in darkness and incubated at 35 6 2 C. To simulate the treatment of an infested host, the baskets were taken out, and the lice were washed with tap water until they were completely free from residual products and placed in Petri dishes with untreated filter papers and incubated in the dark at the same temperatures and RH as above. Two control tests were performed: one with lice placed on unimpregnated filter paper, and another with lice exposed to solvent

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ethylhexyl stearate impregnated filter paper dried for 5 min under a fume hood. The total number of lice ranged from 2 to 10 for each product and for each concentration. The plates with adults and nymphs were observed by stereomicroscope at 10, 15, 20, 30, 60, 120, 240 min, and 24 h PT. Depending on the insects recovering from the treatment procedure, each basket for each experiment could contain one, two, or three lice. Criteria of death were defined as the absence of movement of limbs and gut, with or without stimulation using forceps. Experiments were repeated at least twice. P. capitis eggs, attached ,1 cm from the scalp, were collected by cutting the hair and placing samples into glass jars with screw caps in a polystyrene container to minimize temperature variation. The samples were processed within 2 h after collection. Microscopically homogeneous similar eggs, with closed operculum and in the same phase of embryonic development were selected for use. These eggs were placed in separate glass Petri dishes lined with Whatman No. 1 filter paper (7.0 cm diameter) on the bottom that had been treated with 600 μL of each product at different concentrations and combinations. The two control tests performed were the same as for the adulticidal test. In each dish, the total number of louse eggs ranged from 8 to 12, for each product and for each concentration. The treated samples and controls were incubated at 37 6 2 C and 65 6 5% RH in darkness for 15 days. Depending on the louse eggs recovering from treatment procedure, each Petri dish for each experiment could contain 4
5 eggs. The louse eggs hatching was monitored daily under microscopic inspection. Mortality data of treated eggs were recorded 5 days after the hatching of controls. Louse eggs with closed operculum and nymphs inside were the criterion for embryo mortality (abortive eggs). Experiments were repeated at least twice. The results of this study offered new potential application of natural compounds for the treatment of pediculosis resistant cases and the development of novel pediculicides containing essential oils could be an important tool to control parasitic infestation. Lee et al.13 conducted similar studies using insecticide-impregnated Whatman No. 1 filter paper disks that had been dipped into acetone solutions of insecticides (0.5%, 1.0%, and 10%). Newly hatched first instars blood fed on human volunteers were transferred to a Petri dish fitted with a treated disk and the KT50 and the LT50 were calculated by logit analysis. After logit analysis, the maximum log-likelihood ratio test was used to test the hypothesis of parallelism and equality (P 5 .05) of the regression line. All bioassays were conducted at 30 C and 70% RH. The WHO protocol 198117 for determining resistance in body lice, Pediculus humanus humanus, requires holding lice for long periods, that is, up to 2 days without causing excessive control mortality. In addition, the test required placing the lice on rectangular test paper and covering them with a Petri dish weighted down to prevent escape. The lice had to be removed from the treated paper and placed in a separate clean container. This method

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proved awkward and difficult under certain field conditions and some lice would attempt to escape and be injured by the Petri dish when it was placed over them. The purpose of a study reported by Zeichner18 was to modify the WHO test procedure to make the holding period of lice shorter and the handling of lice easier. Zeichner18 placed lice on insecticide-treated chromatography paper cut into circles (9 cm diameter) and laid flat on stainless steel trays. A pipette was used to evenly distribute 0.82 mL of an insecticide-carrier dilution onto the paper. Insecticide-treated papers were placed in the lid (or larger half) of a standard plastic Petri dish (100 3 15 mm). The Petri dish was used upside down with the top of the lid sitting on the table. Ten to 20 adult lice were placed on the smaller half of the Petri dish, which was then tilted so the lice accumulated on the bottom edge. They were then dropped onto the center of the insecticide-treated paper and the smaller half of the Petri dish was quickly placed into the larger half before the lice could run to the sides of the dish. The lice were unable to climb the sides of the Petri dish, hence were forced to stay in contact with the insecticide-treated paper. Because the paper was slightly larger than the smaller half of the Petri dish, the lice could not crawl off it. Testing was conducted in an illuminated environmental chamber at 28 6 1 C and 60 6 10% RH. At least five different doses, three replicates each, were tested on three different days for each insecticide. Only one insecticide was tested on any given day. After 6 h, knockdown readings were taken. The test time of 6 h was chosen because it allowed conduct of the test within a normal 8-h workday. The Petri dish was tuned on its side and lightly tapped on the table. Those lice that were unable to cling to the chromatography paper were counted as knocked down. To enhance consistency, the Petri dish was tapped while holding it between the thumb and first two fingers, with the heel of the hand resting on the table. The Petri dish was then raised B4 cm and swung down onto the table. Healthy lice clung to the paper even if the Petri dish was tapped forcefully, whereas insecticide-intoxicated lice readily became dislodged. Probit analysis and calculation of lethal dose ratios were performed. Khater et al.19 reported on a study using the buffalo louse, Haematopinus tuberculatus (Nitzsch) and the filter paper contact bioassay previously described. Burkhart and Burkhart20 noted that a standardized method for determining efficacy of various head lice therapies for public assessment had not been adopted by the WHO or research community, because all attempts to colonize this louse species in the laboratory had failed. However, reasonable numbers for testing could be obtained by finger grasping individual head lice while foraging through patients’ scalps, or by raking a comb through sections of the scalp and removing adult lice captured between the metal teeth of the comb. Lice eggs (unhatched nits) from patients’ scalps were easily removed for testing as well, by sniping individual hairs with nits attached from infested individuals.

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The modifications to the WHO 1981 protocol by Zeichner18 offer a standardizable test but were not optimum for head lice analysis. Because head lice are treated usually with medicated shampoos, placing insects on insecticide-impregnated paper for several hours was not as appropriate as immersing insects for the clinical duration of insecticide exposure. After such a dip method of insecticidal exposure, the lice are then rinsed, dried, and observed for several hours. Moreover, by only exposing lice to treated paper, one neglected possible penetration of insecticides via the alimentary tract, as well as distortion of Nirmala diffusion kinetics through the cuticle. In addition, in assays with impregnated disks, lice dwell in a fully enclosed environment with a Petri dish upside down, seemingly giving an advantage to more volatile agents that could be absorbed through the spiracles. A perfect protocol would not offset the normal pharmacokinetics of adsorption, distribution, metabolism, and excretion observed in the clinical arena. Therefore these authors concluded that in vitro testing, which emulated clinical usage with the therapeutic agent for head lice, would produce results that mirror full-scale clinical trials. Returning to the modifications offered to WHO body lice testing by Zeichner,18 after 6 h “the Petri dish is turned on its side and lightly tapped on the table. Those lice that are unable to cling to the chromatography paper are counted as knocked down.” Given that head lice claws are designed to grab hair, this method may be inappropriate because they may be unable to sufficiently grab the paper. Therefore Zeichner18 offers for consideration an in vitro pediculicidal test in which head lice are exposed to insecticides for similar duration, concentration, and exposure imitating routine clinical usage, since such a test would be reproducible and its results would reflect results obtained in clinical trials. In short, their standardized model of insecticidal killing was merely a modification of the detailed Meinking et al.21 original protocol. For example, with testing of a specific medicated shampoo, lice would be immersed into the same dilution and for the same time as clinical protocol indicated. Lice would then be transferred into a container of water simulating the rinsing of medication from the hair, before being placed on a cloth disk dampened with filtered water. Lice activity was monitored continuously both visually and by light microscopy for the first hour, followed by 30 min observations thereafter. Even if all motor and respiratory function ceased, monitoring continued optimally for 24 h. It was noted that lice mortality after 24
36 h due to physiologic desiccation was normal. Ovicidal activity, which may be more relevant than pediculicidal function, would also be tested by this modified standardized protocol. Nits were examined under a microscope to prove an intact larva existed. The features that point to a viable egg are existence of tan to brown oval egg capsule often with a visible eye spot on the developing larvae, with an intact operculum. Hatched eggs were clear to white clinically, and by light microscopy, an empty shell was noted with a ruptured

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operculum. Because eggs are laid close to the base of hair follicles, the distal ends of the hairs were attached to small adhesive labels to facilitate transport of eggs to test solutions of insecticides for designated time intervals, followed by rinsing in tap water, and then air dried at ambient temperature. The eggs were subsequently incubated in the dark for hatching observed after 2 weeks for hatching to determine ovicidal activity. A later paper by Meinking et al.22 extensively described procedures for preparing and impregnating cotton disks with pediculicidal and ovicidal products and the rationale for using this methodology for head lice exposure.

Immersion assays Heukelbach et al.23 reported on the in vitro efficacy of over-the-counter botanical pediculicides against the head louse. In this publication, the authors listed all the 49 papers in the literature of which they were aware, which described in vitro studies of the efficacy of pediculicides. This listing is a review of publications, beginning in 1945
2008, that reported on the various exposure techniques against various stages of head, body, and pubic lice and the criteria used for mortality. Since the mortality criteria reported in these 49 studies varied in stringency, these authors conducted a study of six commercial products (and two control groups: one positive and the other negative) with an application of stringent mortality criteria. For each of the eight groups (six treated/two controls), 25 6 3 lice were tested. Lice clasping hair strands were immersed completely in the product for 1 min and then placed with hairs onto Whatman filter paper in Petri dishes, pools of the products were wiped from the lice with a jeweler’s forceps directed under a dissecting microscope. The lice in the negative control group were placed directly on moistened filter paper without any treatment. They were not dipped in water or wiped with forceps, as these actions have been shown in previous studies to have no effect on lice activity levels. To prevent lice from desiccation, the filter paper had been previously moistened with 200 μL tap water. To simulate treatment on an infested host, head lice were washed in tap water after 20 min and placed into a new Petri dish with unused filter paper. Lice on the filter paper were examined under a dissecting microscope by a single observer in all cases to prevent interobserver variation. The standard criteria for evaluating mortality were compared with a more stringent method to see if standards require revision. The standard criteria for mortality, as originally used by the WHO to measure sublethal effects as opposed to mortality in basic toxicological studies and to test for insecticide resistance, were the inability to walk in a progressive fashion and the absence of a righting reflex when rolled onto the back. In their study, the authors defined these criteria as “some vital signs.” Although the WHO criteria were meant to compare sublethal effects, the criteria have been used by many to test the overall efficacy of lice control products.

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The stringent criteria used in this study defined death of a louse as the complete absence of any vital signs such as gut movement and movement of antennae or legs, with or without stimulation using forceps. This status was defined as “no vital signs.” Lice were defined as “active” if no changes in their levels of activity or behavior were observed PT. An evaluation of these categories was conducted at 30, 60, and 180 min PT. All lice were maintained at 27 C during the tests. Statistical analysis was performed using the exact version of the chisquare test (to compare differences between experimental groups) and the McNemar
Bowker test to compare paired data within each experimental setting. This study demonstrated that the commonly used less stringent criteria overestimated mortality of head lice as a result of the protective phenomenon of stasis (physically shut down) or sham death observed in exposed lice that might recover after some time. A variation of the direct immersion technique24 has been reported where head lice were given squares of nylon gauze as a substrate and these gauze squares (15 3 15 mm) containing 20 lice each were immersed for 10 s in a test solution. The use of the gauze squares also provided a substrate for oviposition so that lice eggs could be handled without risk of damaging them and the effects if treatments could be easily observed. The squares containing lice eggs were immersed in a similar fashion as were the adult lice. Two reports of the in vitro evaluation of pediculicides against Linognathus species were found. A 2016 Master’s thesis by Amante25 described the use of the FAOmodified protocol for the immersion of adult face louse (Linognathus ovillus). The test substances (plant extracts) were diluted in distilled water and 2% DMSO at various concentrations. The in vitro tests were conducted within 1 h after lice collection. Ten active lice were placed in Petri dishes in three replicates and 2 mL of each concentration was directly added to the three replicate Petri dishes and incubated at 27 C
28 C and 75%
85% RH for 24 h. The test solutions were removed, using Whatman No. 1 filter paper, after a 1
2 min contact time. The lice in the Petri dishes were closely observed for death (no signs of movement at all) under a stereomicroscope at 30 min, 1, 2, 3, 6, 12, and 24 h PT. Mortality was determined using the Abbots formula. A research paper by Kumar et al.26 reported on an in vitro immersion study of crude plant extracts against a goat louse Linognathus africanus. Lice were randomly divided into groups containing 20 lice each and the lice were immersed (pour-on method) into a serial distilled water dilution of each extract. A distilled water control group was maintained for each experiment. After immersion, the lice were placed into Petri dishes that were kept at 25 6 2 C and 80 6 5% RH in a desiccator within an incubator. The mortality in all groups was recorded after 30 min, 1, 2, 4, and 6 h PT. Probit analysis was done on mortality data to calculate the LC50 value.

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Similar in vitro methods have also been used for the detection of ectoparasiticide resistance in species of sucking lice and the screening of test compounds against resistant species.27 Australian sheep have long been treated to remove lice. Dipping, either by swimming shorn sheep through a long bath of insecticide solution or by topical spraying in sheep showers have been in use since the insecticides that could be applied like this were available from the 1950s. Organochlorines were commonly used until banned in 1957. The organophosphates then became available as well as arsenic until it was deregistered in 1987. By that time, however, pyrethroid products had captured a considerable part of the sheep lice treatment market (70% market share), many being applied by the “pour-on” method. Pyrethroid resistance was suspected after about only 4 year postregistration. An investigation of the causes of pour-on failures was initiated in 1986 and a laboratory bioassay was quickly developed by Levot and Hughes.28 The bioassay is a self-dosing
treated surface test utilizing a fiber substrate which allows sheep lice to move freely. One milliliter aliquots of serial dilutions of acetonic solutions of the synthetic pyrethroid cypermethrin were pipetted onto 6 cm squares of pure cotton cloth (25 6 5 thread/cm). This size square absorbed 1 mL acetone evenly without access. Cypermethrin was chosen as the initial test insecticide because this chemical was implicated in most reports of control failure. Approximately 8
10 concentrations plus controls were set up in duplicate in each bioassay. Adult lice collected from sheep by vacuum into a sample tube were transferred by camel’s hair brush onto each cloth and confined within a stainless steel ring (50 mm diameter, 15 mm high). Bioassays were conducted at 34 C and 80% RH in darkness for 16 h. Although sheep skin is B37 C, early experience suggested that control mortality was less at 34 C. At this time, lice were categorized under 10 3 magnification as alive (active) or moribund (unable to move away if touched) and the responses analyzed using the probit method. This bioassay was adopted by State Departments of Agriculture throughout Australia to test for pyrethroid resistance. The bioassay is a simple test and is suitable for any fast-acting contact insecticide, which encompassed, at the time of this publication, all the compounds currently registered for lice control in Australia. The bioassay is not, however, suitable for chemicals that must be ingested by the insect. Although some lice survived for several days under the test conditions, no recovery period was allowed for moribund lice because there was unsatisfactorily high mortality among the control lice.

In vivo method(s) Efficacy studies of sucking lice of ruminants should be conducted in accordance with the WAAVP guideline for the evaluation of ectoparasiticides against biting lice, sucking lice, and sheep keds on ruminants (see Appendix A of Ref. [29]). Sucking lice are vulnerable to systemic insecticides, whereas

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skin-chewing (biting) lice are better controlled with parasiticides that act on the surface of the externally treated animal.30 Clinical field trials to eradicate sucking lice (L. vituli) on cattle using pyrethroid pour-on products were successful in 28 of 33 herds, but sucking lice were present in 3 herds 3
6 months PT.31 Two herds were reinfected with sucking lice 12 months PT. Ectoparasiticides are highly effective against susceptible species of sucking lice. However, the development of ectoparasiticide resistance in sucking lice of ruminants may reduce the effectiveness of the treatment.32

References 1. Culpepper GH. The rearing and maintenance of a laboratory strain of the body louse. Am J Trop Med 1944;24:327
9. 2. Lang JD, Roan CC. An improved method for rearing head lice. J Med Entomol 1974;11(1):112. 3. Culpepper GH. Rearing and maintaining a laboratory colony of body lice on rabbits. Am J Trop Med 1948;28:499
504. 4. Gilbert IH. Laboratory rearing of cockroaches, bed-bugs, human lice and fleas. Bull WHO 1964;31:561
3. 5. Takano-Lee M, Yoon KS, Edman JD, Mullens BA, Clark JM. In vivo and in vitro rearing of Pediculus humanus capitis (Anoplura: Pediculidae). J Med Entomol 2003;40(5):628
35. 6. Fuller HS, Murray ES, Snyder JC. Studies of human body lice, Pediculus humanus corporis. I. A method for feeding lice through a membrane and experimental infection with Rickettsia prowazekii, R. mooseri and Borrelia novyi. Pub Health Rep 1949;64(41):1287
91. 7. Haddon Jr. W. An artificial membrane and apparatus for the feeding of the human body louse Pediculus humanus corporis. Am J Trop Med Hyg 1956;5:315
25. 8. Haddon Jr. W. The maintenance of the human body louse Pediculus humanus corporis through complete cycles of growth by serial feeding through artificial membranes. Am J Trop Med Hyg 1956;5:326
30. 9. Lauer DM, Sonenshine DE. Adaptations of membrane feeding techniques for feeding the squirrel flea, Orchopeas howardi, and the squirrel louse, Neohaematopinus sciuropteri, with notes on the feeding of the human body louse, Pediculus humanus var. corporis. J Med Entomol 1978;14(5):595
6. 10. Mumcuoglu YK, Galus R. Engorgement response of human body lice Pediculus humanus (Insecta: Anoplura) to blood fractions and their components. Physiol Entomol 1987;12:171
4. 11. Habenank B, Schrader G, Scheurer S, Schein E. Investigations on the in vitro feeding and in vitro breeding of the human body louse Pediculus humanus corporis (Anoplura: Pediculidae). In: Robison WH, Rettich F, Rambo GW, editors. Proceedings of the 3rd international conference on urban pests; 1999. 12. Schrader G, Schmolz E, Konning M, Dahl R. Survival and reproduction of a laboratory strain of body lice (Phthiraptera: Pediculidae) at different ambient temperatures. In: Robinson WH, Bajomi D, editors. Proceedings of the 6th international conference on urban pests; 2008. 13. Lee SH, Yoon KS, Williamson MS, Goodson SJ, Takano-Lee M, Edman JD, et al. Molecular analysis of kdr-like resistance in permethrin- resistant strains of head lice, Pediculus capitis. Pestic Biochem Physiol 2000;66:130
43. 14. Takano-Lee M, Velten RK, Edman JD, Mullens BA, Clark JM. An automated feeding apparatus for in vitro maintenance of the human head louse, Pediculus capitis (Anoplura: Pediculidae). J Med Entomol 2003;40(6):795
9.

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15. Yoon KS, Strycharz JP, Gao J-R, Takano-Lee M, Edman JD, Clark JM. An improved in vitro rearing system for human head louse, allows the determination of resistance to formulated pediculicides. Pestic Biochem Physiol 2006;86:195
202. 16. Di Campli E, Bartolomeo S, Pizzi PD, Di Giulio M, Grande R, Nostro A, et al. Activity of tea tree oil and nerolidol alone or in combination against Pediculus capitis (head lice) and its eggs. Parasitol Res 2012;111(5):1985
92. 17. World Health Organization [WHO]. Instructions for determining the susceptibility or resistance of body lice and head lice to insecticides. 1981; WHO/VBC/81.808. 18. Zeichner C. Baseline susceptibility of a laboratory strain of Pediculus humanus humanus (Anoplura: Pediculidae) using a modified World Health Organization testing protocol. J Med Entomol 1999;36(6):903
5. 19. Khater HF, Ramadan MY, El-Madawy RS. Lousicidal, ovicidal and repellent efficacy of some essential oils against lice and flies infesting water buffaloes in Egypt. Vet Parasitol 2009;164:257
66. 20. Burkhart CN, Burkhart CG. Recommendation to standardize pediculicidal and ovicidal testing for head lice (Anoplura: Pediculidae). J Med Entomol 2001;38(2):127
9. 21. Meinking TL, Taplin D, Kalter DC, Eberle MW. Comparative efficacy of treatments for pediculosis capitis infestations. Arch Dermatol 1986;122:267
71. 22. Meinking TL, Entzel P, Villar ME, Vicaria M, Glendene AL, Porcelain SL. Comparative efficacy of treatments for pediculosis capitis infestations. Arch Dermatol 2001;137:287
92. 23. Heukelbach J, Canyon DV, Oliveira FA, Muller R, Speare R. In vitro efficacy of over-thecounter botanical pediculicides against the head louse Pediculus humanus var capitis based in a stringent standard for mortality assessment. Med Vet Entomol 2008;22:264
72. 24. Burgess IF, Brunton ER, Brown CM. Laboratory and clinical trials of cocamide diethanolamine lotion against head lice. Peer J 2015;3:e1368 Available from: https://doi.org/10.7717/ peerj.1368. 25. Amante M. In vitro lousicidal and acaricidal activities of alkaloid of Calpurnia aurea and fractions of Ricinus communis extracts against Linognathus ovillus and Amblyomma variegatum [Master thesis]. Bishoftu, Ethiopia: Addis Ababa University, College of Veterinary Medicine and Agriculture, Department of Pathology and Parasitology; 2016. 26. Kumar A, Mahour K, Verma S, Vihan VS. Phytochemical investigation and in vitro licicidal activity of some indigenous plants extract under organized and farmer flocks of goat. Natl Conf Rect Innovat Appl Sci Humanit 2015;4(10). Special Issue Oct. 2015, ISSN No. 2277 8179. 27. Levot G. Resistance and control of lice on humans and production animals. Int J Parasitol 2000;30:291
7. 28. Levot GW, Hughes PB. Laboratory studies on resistance to cypermethrin in Damalinia ovis (Schrank) (Phthiraptera: Trichodectidae). J Aust Entomol Soc 1990;29:257
9. 29. Holdsworth PA, Vercruysse J, Rehbein S, Peter RJ, Letonja T, Green P. World Association for the Advancement of Veterinary Parasitology (W.A.A.V.P.) guidelines for evaluating the efficacy of ectoparasiticides against biting lice, sucking lice and sheep keds on ruminants. Vet Parasitol 2006;136:45
54. 30. Stromberg BE, Moon RD. Parasite control in calves and growing heifers. Vet Clin: Food Anim Pract 2008;24(1):105
16. 31. Nafstad O, Grønstøl H. Eradication of lice in cattle. Acta Vet Scand 2001;42:81 Available from: https://doi.org/10.1186/1751-0147-42-81. 32. Taylor M. Applied clinical parasitology for cattle. Chapter 21. In: Cockcroft PD, editor. Bovine Medicine. 3rd ed. Oxford, UK: Wiley & Sons, Ltd; 2015. p. 198
210.

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Chapter 3e

Arthropoda, Phthiraptera, Mallophaga Larry R. Cruthers, MS, PhD1 and Douglas D. Colwell, PhD, FRES, Assoc. EVPC2 1 LCruthers Consulting, Chesapeake, VA, United States, 2Agriculture and Agri-Food Canada, Lethbridge, AB, Canada

Arthropoda Phthiraptera Mallophaga: biting (chewing) lice

Biology and life cycles Biting lice, like all lice species, are wingless, dorsoventrally flattened and permanent ectoparasites of mammals and birds. Biting lice have mouthparts adapted for chewing up epithelial debris on the skin of the mammalian host or on the feathers of birds. These lice infest a wide range of domestic livestock and poultry. They are a source of constant irritation, itching, rubbing, and biting of the skin or hair coat and plumage and are closely adapted to their hosts completely dependent upon them for survival. The entire life cycle of lice occurs on the coat of the host animal; no stage of the lice life cycle occurs off the host or in the host’s environment In general, there are three developmental stages in the life cycle (incomplete metamorphosis) of the biting louse. The lice egg (nit) stage of the life cycle hatches in 7
14 days, depending on the species of lice and on environmental conditions. The three nymph stages of the lice life cycle grow and undergo their molts over B9
22 days, depending on the lice species. The adult lice usually live for 2
3 weeks on the fur or hair of the host animal. They will lay their eggs during this time period. Specifically, the life cycles of several important genera and species are described next.

Trichodectes canis de Geer, 1778—the canine chewing louse This louse of canids is found on domesticated dogs and wild canids throughout the world. The head is broad and rounded anteriorly. The brownpigmented mandibles are located in the center of the head and the abdomen is covered with hairs and lacks pigmented plates. The average time from egg to egg is 3
4 weeks. They prefer to inhabit the back, neck, or head of the host, particularly under mats of hair and around the ears and body openings, whereas in heavy infestations, they cover the entire body. Spread of

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infestations is usually by direct animal contact. This louse is a well-known vector of the dog tapeworm, Dipylidium caninum.

Felicola subrostratus Nitzsch, 1838—the feline biting louse This felid biting louse is found on domesticated cats throughout the world. This louse is readily identified by the shape of the head. The portion of the head anterior to the antennae is triangular. The straight sides (lines) do not quite meet at the point because of a hair groove that extends back to the mandibles. The biology of this louse is very poorly known. The life cycle from egg to egg is B3
4 weeks. It is presumed that their only significant source of food is epidermal debris. Although few clinical signs have been described in infested cats, debilitated cats can develop large numbers of lice if they lose their ability to groom. Lice are typically found under mats of hair and around the ears and body openings. Spread of infestations is usually by direct animal contact. This louse is not known to be a vector of infectious disease agents.

Bovicola (Damalinia) bovis Linnaeus, 1758—the cattle biting louse The cattle biting louse is geographically cosmopolitan in distribution, found on cattle worldwide, but is more frequent in regions with cold winters. It is easily recognized by its large, bluntly triangular head and a yellow-white body with dark bands. Development to adults takes about 4 weeks. The eggs are easily distinguished from those of other cattle lice by their smaller size and their transparent shells. They are most commonly found in colonies or “patches” at the base of the tail, shoulders, and top line of the back. Skin around colonies of this louse can have the appearance of mange lesions. These lice are primarily spread by direct animal contact. This louse has not been linked to any disease transmission.

Bovicola (Damalinia) ovis Schrank, 1781—the sheep biting louse The sheep louse is distributed worldwide. Females cement their eggs to wool fibers about 6
12 mm from the sheep’s skin, but eggs are also placed on hair of legs and other body parts of the sheep. Under normal conditions, the length of a complete life cycle from egg to egg is 34
36 days. Transfer between animals occurs when sheep are in close contact. Lice move to the surface of the fleece when it is shaded and warm and transfer occurs more quickly when sheep have short wool than when the wool is longer. Sheep lice are not known to vector any pathogenic diseases.

Bovicola (Damalinia) caprae Gurlt, 1843—the goat biting louse or short-haired goat louse This goat louse has a brownish-red head and thorax and the yellowish abdomen has brown cross bands that are lighter than and not quite as wide as

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those of Bovicola (Damalinia) limbatus, Gervais, 1844 (Angora goat biting louse) which also occurs on goats. Distribution of the goat-biting lice is cosmopolitan. In the laboratory, an entire life cycle from egg to egg required B37 days. Transmission of lice is generally by close contact of one host to another, although it is possible for goats to become infested by occupying facilities that have been previously occupied by other goats. Goat-biting lice feed on or near the surface of the skin. Goat lice are not known to be a vector to any disease agents.

Bovicola (Damalinia) crassipes Rudow, 1866—the Angora goat biting louse The distribution of this goat louse is limited to that of Angora goats. These lice are covered with setae, giving them a hairy appearance. Their life history and transmission are similar to other species of Bovicola.

Bovicola (Werneckiella) equi Linnaeus, 1758—the horse biting louse This horse louse is worldwide in distribution. The eggs are deposited on the fine hairs of the horse’s coat with the attached end near the skin. Apparently, the coarse hairs, such as those in the mane, are too large for the female to use, because the eggs are usually never found on them. It localizes on the sides of the neck, in the flanks, and at the base of the tail, but when heavily infested, the lice may be found over most of the body except the lower legs, tail, mane, and ears. The egg to egg life cycle is usually 30
45 days. They are readily passed from one horse to another by physical contact, especially when confined together, but can also be spread mechanically by brushes and equipment or from the environment like a horse trailer. A second biting louse also infests horse, Trichodectes pilosus Giebel, 1874. No known equine pathogens are transmitted by these lice.

Werneckiella ocellatus Piaget, 1880—the donkey chewing louse This donkey louse along with the subspecies, Werneckiella equi asini, infests donkeys and mules worldwide. They occur mainly on the neck and around the base of the tail but may occur over the entire body in heavy infestations. Adult lice are $ 2 mm in length and feed on the skin and hair debris. Adult female lice lay eggs attached to the hairs of the animal. The life cycle from egg to adult takes about 3 weeks with all three nymphal stages taking place on the same animal. The adult louse life span is limited to 2
3 months. Werneckiella species can act as vectors of equine infectious anemia.

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Menacanthus stramineus Nitzsch, 1818—the chicken body louse This chicken biting louse is the most common and most destructive louse found on chickens and has a worldwide distribution. The complete life cycle requires about 2 weeks. The eggs have characteristic filaments on the anterior half of the shell and on the operculum. The eggs are mostly deposited in white clusters at the base of the feathers, especially around the vents. The lice also occur in other sparsely feathered areas, such as the breast and thighs, causing a marked reddening of the skin. The lice have chewing mouthparts and feed on blood, feathers, skin, or scales of the bird. They are transferred between birds when in close contact. The chicken body louse is not a vector of any known disease agents.

Menapon gallinae Linnaeus, 1758—the poultry shaft or feather louse This rather small poultry louse is cosmopolitan in distribution. It feeds mainly on skin and feather debris but may also suck blood from the wounds it produces. It is found in groups on shafts of new quill feathers on the chest, shoulders, and the back of birds. The whitish eggs are laid in strings or clusters at the base of the feathers. The complete life cycle takes about 3
5 weeks. Chickens are also infested with other lice, such as the brown chicken louse (Goniodes dissimilis), fluff louse (Goniocotes gallinae), head louse (Cuclogaster heterographus), large chicken louse (Goniodes gigas), and the wing louse (Lipeurus caponis).

Rearing method(s) No reports of in vitro rearing of any members of Trichodectes canis or Felicola subrostratus were found. Bovicola crassipes and Bovicola limbata are parasites of Angora-type goats. A study by Hopkins and Chamberlain1 describes techniques that were successfully used to colonize these two goat lice in the laboratory. Food was prepared by cutting 20 3 20 cm sections from a fresh, closely sheared Angora goatskin, placing them flesh side down on glass plates, and freezing them. Then the flaky outer portion of the frozen skin was scraped off, the scrapings dried at 35 C for 24 h, and the fragments were cut into particles small enough to pass through a 14 3 18-mesh screen. For long-term storage, the screened material was placed in moisture-proof containers at 25 C; however, sufficient food for the colonies for 2
3 weeks could be removed from the frozen stock and held at 5 C
8 C. The lice used for starting the colony were obtained from infected Angora goats. The nymphs and adults were separated and the nymphs were discarded. The adult lice were separated by species and placed in glass shell

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vials. (0.5 dram for ,50 and 1 dram vials for 50
200 lice). Food was placed in the vials at the rate of 175 mg/100 lice. These vials were placed in 30- and 50-mL glass beakers which, in turn, were placed in round polystyrene containers (12 cm wide at top, 11 cm wide at bottom, and 14 cm tall) that had tight-fitting lids. A 95 3 35 mm polystyrene Petri dish with holes in the bottom was inverted and placed in each container to make a platform on which to set the beakers. Several trial runs resulted in the selection of 72% RH and a temperature of 35 C 6 1.5 C for rearing B. crassipes and 76% RH and 35 C 6 1.5 C for the rearing of B. limbata. The eggs were never exposed to temperatures below 27 C. The photoperiodic regimen was 15 h dark, 8 h dim light, and 1 h of room light daily. Adults were maintained in the vials at a ratio of one male to three females. Since the lice naturally cement their eggs to mohair on the host animal, several unwashed goat hairs 4
8 cm long were made into loose coils and placed in the vials. Portions of the mohair coils were covered with food because the lice in vials appeared to prefer to oviposit below the surface of the food. Eggs were collected every 1
4 days. The eggs on the pieces of mohair were then counted and placed in new vials (0.5 dram for ,50 and 1 dram for 50
200 eggs) to incubate after the loops bearing the eggs were cut into short lengths so emerging nymphs would have other hair or food particles to grasp. Food was added at the same time (35 mg/100 eggs) or when hatching began. Also, after 22 days (9 days for the eggs to hatch and 13 days for nymphal growth), a new vial and fresh food (140 mg/100 B. crassipes and 70 mg/100 B. limbata) were supplied. If ample food was provided, cultures could be reared successfully without making this change, but a provision of a new container and fresh food at 22 days minimized the exposure to disease organisms. Thus goat-biting lice fed a diet prepared from goatskin can be reared apart from the animal. After Hopkins and Chamberlain1 developed procedures for colonizing the goat-biting lice, B. crassipes and B. limbata, apart from the natural host, they attempted to determine whether the sheep louse, Bovicola ovis, could similarly be colonized.2 These techniques were similar to those used to colonize B. crassipes and B. limbata. After an experimental colony of B. ovis was established, it was used to study the survival and fecundity of the lice when they were provided a sheepskin diet and held at various combinations of RH and temperature. The combination of 68% RH and 37 C was selected as the most suitable for laboratory colonization. At these conditions and with the sheepskin diet, 61% of the eggs hatched and developed into adults. These procedures for colonizing sheep-biting lice were similar to those of the goat lice with the following exceptions: (1) sheepskin was substituted for goatskin as a diet and wool was substituted for mohair as an egging medium, (2) the eggs were collected every 7 days instead of every 1
4 days, (3) the diet and diet vial were not changed at 22 days; instead, the lice were

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removed from the vial after they became adults, and (4) the colony was held at 68% RH and 37 C 6 1.5 C instead of at 72% or 76% RH and 35 C. Hopkins and Chamberlain3 also reported on an in vitro colony of the cattle biting louse, Bovicola bovis. The colonization techniques were the same as those used to colonize goat lice1 except that (1) cow skin scrapings were substituted for goatskin scrapings as a diet, (2) eggs of B. bovis were collected on mohair every 1
6 days instead of every 1
4 days (mohair was used in lieu of cow hair because it was generally longer and thus more convenient to use and B. bovis oviposited readily on it), (3) 30-mL beakers were used for rearing large cultures (200
1000 lice) though, as before, 0.5 dram and 1 dram glass shell vials were used for smaller numbers of lice, (4) the rearing vessels were held at 70% RH and 37 C 6 1.5 C instead of at 72% or 76% RH and 35 C, and (5) the diet and rearing vessels were not changed at 22 days; instead, the mohair was removed after the eggs hatched (8 days) and the contents (diet and lice) of each vessel were sifted through a screen (0.37 3 0.37 mm openings) to remove old diet and feces when most of the lice became adults (15
20 days posthatch). Then new diet and mohair were added. As demonstrated, Hopkins and Chamberlain1,2 reared goat and sheepbiting lice in vitro, on preparations of skin from the natural host. However, since such skins are not always obtainable, and diets from them can vary in nutrient value, an attempt was made to develop diets composed of commonplace ingredients likely to be consistent in quality. Various tissues and oils were mixed together and their effect upon fecundity and longevity of lice was evaluated by Hopkins et al.4 On the basis of performance and availability of ingredients, specific artificial diets were chosen as standards for in vitro colonies of the three species of lice: dehydrated veal and dehydrated lanum, prepared in a ratio of 3:1 for B. limbata; dehydrated veal and wool extract, prepared in a ratio of 4:1 for B. crassipes; and dehydrated veal and wool extract, 3:1 for B. ovis. Occasionally, in vitro investigations are undertaken to verify life-cycle information for which little is known. For example, although the chicken body louse, Menacanthus stramineus, is a common parasite of domestic chickens and turkeys throughout the world, little was known of its biology. Stockdale and Raun5 conducted in vitro investigations to elucidate the temperature, humidity, and nutritional requirements of M. stramineus and then conducted in vivo studies to verify the life-cycle information obtained from the in vitro studies. Note: similar studies may be useful for the in vitro rearing of other bird lice of interest to an investigator. Louse eggs were obtained by plucking a feather, containing the egg mass from a heavily lice-infested bird. The feather was placed in a water bath incubator at 35 C and 95% RH. After a sufficient number of nymphs had hatched, 6
10 first instars of a known age were placed in plastic rearing cages. The cages in all the in vitro investigations were round plastic zipper boxes of two sizes, the larger 5 3 2.6 cm deep and the smaller 3.1 3 2.6 cm

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deep. Ten to 20 holes, B2 mm in diameter, were made in the top and bottom of each cage by plunging a heated probe through the plastic. Silk bolting cloth was used to line the tops and bottoms of the containers to prevent the lice from escaping through the ventilation holes. The tight-fitting friction cover prevented louse escape and was easily removed. A water bath was used as an improvised incubator. An aluminum cake pan, 29.2 3 21.6 cm, was floated on the water surface, and the plastic cages containing the lice were placed in this pan. This constant temperature source was most successful and was used for most of the tests. Commercial incubators were also used as a heat and humidity source for some of the studies. Also, a slide drier was also used as a heat source for rearing and as an aid in observation. When cages of lice were removed from the incubators for feeding, counting, and data recording, they were placed on a slide drier to keep them at the desired temperature. Freshly plucked pinfeathers, taken from young broilers, were placed in the cages as food. These fresh feathers, the shafts of which were filled with a pulpy, liquid material (lymph and blood), were placed in the containers at 3
4 h intervals, seven times during a 24 h period. Once during each 24 h period, the lice were examined, and the number surviving and their progress toward maturity were recorded. At this time, all accumulated feathers from the preceding 24 h were removed. Later experiments were successfully conducted when fresh pinfeathers were provided every 6 h (four times a day), but once daily feedings were not successful. The highest percentages of lice reared from the egg stage to maturity in the shortest time were achieved by using the water bath set at 35 C and 95% RH; 9
13 days were required, with 59% of the lice reaching maturity. A 3-day interval was the usual period required for each instar. Mated females produced 0
4 eggs/day and averaged 1.6 eggs a day during a 12.4-day adult life. Peak oviposition occurred when females were 5
6 days old. A study by Maturano and Daemon6 demonstrated that the inclusion of turkey skin to a feather diet resulted in the greatest number of adult large turkey louse Chelopistes meleagridis. The addition of skin in the diet was a determining factor for development to the adult stage, since 48% of the lice fed this diet reached maturity versus 1.3% that reached maturity fed only with feathers from the host turkey (Meleagris gallopavo). The development time of the males and females was similar (mean of 29.38 days), without any differences in the sexual proportion of the adults.

In vitro method(s) Filter paper contact assay An in vitro assay, which utilizes biting lice of cattle (B. bovis), was established as a primary screen for the evaluation of compounds against this target

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ectoparasite (personal communication). The relative ease of obtaining these ectoparasitic species from infested animals and the potential of screening large numbers of compounds provided an efficient experimental model for determining the potential activities of new parasiticides. Test substances were weighed into 10 mL glass tubes and solubilized in DMSO. Then an appropriate volume of IPA was added. The test substances were thoroughly mixed to yield a solution. Serial dilutions were prepared at a rate to provide concentrations of active ingredient on the filter paper disks of between 0.001 and 10 mg/cm2. Untreated controls consisted of IPA alone. The serial dilutions were poured into 50 mm Petri dishes. The filter paper disks were dipped in the solution for 1 min and then removed allowing the excess solution to drain into the Petri dish. The treated filter paper disk was laid on a sheet of aluminum foil to dry at RT in a fume hood. Five filter paper disks were prepared for each test substance and dilution rate. Once the filter paper disks had dried (B24 h), the treated filter paper disks were wrapped in aluminum foil and stored at 4 C until ready for use. If the evaluation is conducted the following day of filter paper preparation, then no storage was required. For evaluation, the treated filter paper disks were placed in 50 mm Petri dishes. Ten biting lice (stage identified) were added onto the surface of the disks. The Petri dishes were incubated at 30 C and 80% RH for 2 h. Following the 2-h incubation period, the lice were examined. The effect of the test substance on the specimens was classified by activity as follows: Inactive: normal mobility or mobile but some effect. Active: movement of body or legs but not of insect or no movement visible. The number of live/dead insects in the untreated control, reference standards (e.g., permethrin and flumethrin), and experimental insecticides were recorded. In order to determine the concentration activities of the test substances, LD50 and LD90 values were determined by probit or regression analysis. A high mortality rate ( . 10%) in the untreated controls required a repeat test with new insects and/or repeated at a shorter incubation period (1 h exposure). The in vitro residual and ovicidal efficacy of tea tree and lavender essential oil suspensions were tested7 against the donkey chewing louse Bovicola ocellatus using contact assays. Filter papers (90 mm diameter, Whatman No. 1) were placed in 90 mm diameter polystyrene Petri dishes and 800 μL of each test suspension was pipetted evenly onto separate filter papers. This volume was enough to fully saturate the filter paper. The Petri dishes were then placed in a fume hood for 5 min to allow the filter paper to absorb the liquid. Ten female lice were placed onto the filter paper, lids were fitted, and the Petri dishes were maintained in darkness in an incubator at 35 C and 75% RH. Lice were checked for mortality at 15 min and at 1, 2, 3, 4, and

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24 h after exposure to the filter paper. During these checks, the Petri dishes were removed from the incubator and mortality was assessed by observing the lice under a dissecting microscope (30 3 magnification); absence of movement of the legs, mouthparts, antennae, or abdomen even when stroked with a dissecting needle was taken as indicating death. The Petri dishes were immediately returned to the incubator and the assays were repeated three times with adult lice and also three times with small nymphs that were considered to be first or second instars. The residual activity of these essential oils was observed in open (free evaporation) and closed (reduced evaporation) Petri dishes. Contact assays were conducted as previously described, except that treated filter papers were left for 0, 1, 2, and 5 h with the Petri lids off, or for 0, 20, or 40 h with the Petri dish lids on, before lice were exposed. Lice were examined for mortality after 1, 4, and 24 h PT. The assays were repeated three times with both adult and nymphs, as described earlier. The ovicidal activities of these essential oils were also determined using the filter paper contact assay. There was also an untreated (dry filter paper) control. Filter papers were treated in the same way as described and left in a fume hood for 5 min before 20 B. ocellatus eggs were removed from the hair on which they were attached and placed onto each filter paper using a fine paint brush and forceps. Lids were placed on the Petri dishes, which were maintained at 35 C and 75% RH. The number of eggs that hatched were recorded on days 1, 4, 8, and 12 after their incubation, after which no further eggs hatched in any treatment. For observations of hatch, Petri dishes were removed from the incubator and the eggs were inspected under 10 3 magnification; lice were considered to have hatched if the head and thorax had emerged from the chorion. The ovicidal trials were triplicated. The toxicity of the essential oils and excipient formulations to B. ocellatus were examined at 2 h postexposure. Louse mortality was the dependent variable in a three-way ANOVA with live stage (adult or nymph), essential oil, and excipients as factors. Tukey multiple range tests were used for post hoc analysis. The median time (h) required to kill 50% of lice after exposure to the formulations (LT50) was calculated for each time period for which the filter papers were left before lice were introduced, for adults and nymphs in open and closed containers. The LT50 was used as the response variable in a general linear model with preexposure time as a continuous variable and excipient type as a factor. Egg hatch data were analyzed using chi-square analysis.

Cloth square contact assay An in vitro assay, which utilizes biting lice of sheep (B. ovis), and previously described by Levot and Hughes8 was used as an evaluation assay against susceptible and pyrethroid-resistant sheep body lice.

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An insecticide susceptible strain (Peak Hill) and a pyrethroid-resistant strain (Rowena) of adult B. ovis were used for the evaluation. Serial dilutions of the test compound and cypermethrin were prepared in acetone. These serial dilutions were prepared at a rate to provide concentrations of active ingredient on filter paper disks to produce 0%
100% mortalities. Untreated controls consisted of acetone alone. One mL aliquot of the acetonic serial dilutions was pipetted onto cotton squares (6 cm2, 25 threads/cm). Duplicate samples were prepared for each serial dilution concentration. The treated cotton squares were air dried at RT for B24 h in a fume hood. For evaluations, the treated cotton squares were placed into Petri dishes (9 cm diameter). Adult body lice from the susceptible Peak Hill or pyrethroid-resistant Rowena strains were collected by suction from naturally infested sheep. Adult lice (B10) were added to the surface of the treated cotton square and confined using a stainless-steel ring (50 3 15 mm). The Petri dishes were incubated at B34 C
35 C and 75% RH. Each concentration (replicated) was assessed after 24 and 30 h (test compound) or 16 h (cypermethrin) PT. Following the incubation periods, the lice were examined and the effect of the test substance on the specimens was scored as live or responding (dead or knockdown). The number of live/dead insects in the untreated control, reference standards, and experimental insecticides was recorded. In order to determine the concentration activities of the test substances, LD50 and LD95 values were determined by probit regression analysis.

Wool assay Tea tree oil (TTO) from the Australian native plant Melaleuca alternifolia had been shown to have a wide range of insecticidal and repellent activity against arthropods. A paper by James and Callander9 reported the results of in vitro laboratory experiments conducted to test the effect of TTO against sheep lice and eggs. Lice and eggs used in this study originated from sheep from an experimental infested flock held since 2001 without chemical treatment. The lice were freshly collected on the day each study was established using an aspirator connected to a vacuum pump and held in an incubator at 35 C and 65% RH until used. Since the initial two treated surface methods that were used have been previously described for assessing contact insecticides (cotton cloth squares and filter papers), the details of these bioassays will not be elucidated. However, wool assays were also used to detect bioactivity and this procedure will be explained in details to follow. Dilutions of TTO were emulsified in water with a 1% blend of ethoxylated castor oil and ethoxylated oleic acid (ALK). Amounts of 400 mg of

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pesticide-free wool from Merino sheep were immersed in 20 mL volumes of these solutions in 50 mL beakers for 60 s and agitated to ensure complete wetting. Wool was then removed from the beaker, allowed to drain, and a standard method of shaking seven times with a brisk wrist movement used to remove excess fluid. The treated wool was added to 28 mL capacity flat bottom glass bottles and at least 20 adult lice, freshly collected from live sheep, were added to each bottle. There were two control groups, one with lice in dry untreated wool and one with lice in wool treated with emulsifier without TTO, and three replicates for each treatment or control. The assay bottles were held in an incubator at 36.5 C and 65% RH and lice were assessed as live or knocked down or dead at 24 and 48 h PT. To test the residual effect of TTO, 50 mg amount of pesticide-free wool was thoroughly wetted with 500 μL of TTO at the desired concentration solubilized in water with 2% ALK. Concentrations of up to 7.5% TTO were tested and control preparations were treated with ALK without TTO. The wool was dried in a fume hood at RT for 1.5 h then inserted into 20 mL flat bottom glass vials. Twenty lice were added to each, the vials placed in an incubator at 36 C and 65% RH and lice assessed at 24 and 48 h PT as earlier.

Fumigation assay James and Callander’s study9 also included two methods to test for fumigant effects. The first method used fumigation chambers. Test arenas (Fig. 3e.1A) consisted of 90 mm glass Petri dishes with uncovered 55 mm glass Petri dish FIGURE 3E.1 Fumigation assay design by James and Callander9 Chambers used to assess fumigant effects of TTO when it was delivered (A) on a glass surface or (B) applied to wool. TTO, Tea tree oil. Reprinted with Permission from Publisher.

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bottoms within to contain the lice. Twenty adult lice were counted onto 55 mm filter papers and placed in the small Petri dishes. Measure volumes of test solution were dispensed onto 21 3 21 mm glass coverslips placed within the large dishes, but outside of the 55 mm dishes. Lids were applied to the large dishes which were held in an incubator at 36 C and 65% RH. Lice were inspected at 3 h and the numbers of lice knocked down or dead recorded. The small dishes containing lice were removed from the larger dishes and returned uncovered to the incubator. After 24 h, they were reexamined to identify lice that recovered. To determine if the fumes of TTO applied to wool caused mortality of lice not actually contacted by TTO, the apparatus shown in Fig. 3e.1B was used. Glass tubing, 50 mm in length and 15 mm diameter, was glued through holes drilled into the screw tops of 80 3 27 mm diameter glass vials. A piece of nylon mesh was secured across the bottom end of each tube and the chamber so formed used to contain 20 lice in 20 mg of untreated wool. Test wool was weighed to 400 mg amounts and submerged in 20 mL of test solution for 30 s. It was then removed and placed onto paper toweling which was folded over the wool and gently compressed to remove free liquid. Concentrations of TTO between 0.25% and 2% emulsified in water with 1% ALK were tested, with dry wool and wool treated with 1% ALK as controls. The treated or control wool was added to the bottom of the glass vials taking care to position the wool so that there was no contact with untreated wool when the lids were screwed in place. There were three replicates for each concentration and mortality of lice was assessed after 24 h. To test for fumigant effects against eggs, lice were collected from a heavily infested sheep and held overnight in the laboratory on wool that had been clipped some weeks previously from an infested sheep and stored at 225 C. The wool was inspected the next morning and newly deposited eggs that were opaque and apparently healthy were carefully removed from the wool, together with wool fibers when attached. Eighty eggs were counted to 8 groups of 10, placed on filter paper in 50 mm Petri dishes and added to the fumigation chambers (Fig. 3e.1A). Treatments were 60 μL and 15 μL of 100% TTO added to the cover slips in the fumigation chambers as previously described. There were three replicates for each of the two TTO treatments and two for the control (no TTO). Eggs were exposed in the fumigation chamber in an incubator set at 36.5 C and 65% RH for 24 h before being transferred to clean dishes, maintained at similar temperature and RH. They were assessed daily under a microscope over 14 days and classed as hatched where nymphs had successfully emerged completely. In a further preliminary experiment to simulate the effects of immersion dipping, pesticide-free wool was weighed to 400 mg lots and immersed for 30 s in 1% TTO emulsified in water with 1% ALK solution with TTO, or left untreated. Treated wool was then removed from the solution with forceps and placed onto paper toweling. Lice eggs were collected as described earlier but were not dipped with the wool. Instead, to facilitate observations and

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because of the potential for eggs to be lost during dipping, in each treatment, the wool was parted after removal from the solution, 10 eggs positioned in the middle and the wool folded back over the eggs. The preparations were then placed in 28 mL McCartney bottles in an incubator at 36 C and 65% RH for 14 days to allow viable eggs to hatch. Eggs were inspected daily at 45 3 magnification with a dissecting microscope to determine whether embryos had developed and whether or not hatching had occurred. Experiments were of similar design, with 10
20 lice/replicate and 2
3 replicates for each test concentration or control. Analysis was with one-way ANOVA, with appropriate transformations to normalize the data where necessary.

Feather assay Al-Quraishy et al.10 reported on the effects of a neem seed extract on biting lice (featherlings) of chickens. Several infested feathers were cut off from the plumage of chickens and/or ducks and were transported in a closed plastic tube to the laboratory. There, the feathers with the nymph and adult mallophages [L. caponis (wing louse), Columbicola sp. (elongate feather louse), Menapon gallinae (shaft louse), or Philopterus thuringiacus] were placed under the microscope and covered by a tiny droplet of a freshly prepared 1:33 water-diluted test product (neem oil extract, i.e., MiteStop). They were then observed for the next 20 min. The microscopic inspection of the in vitro treatment of the lice on cutoff (or dropped down feathers) demonstrated that the adult lice of all genera reacted in an identical manner when coming into contact with the 1:33 water-diluted product. If a droplet of this dilution was placed on a feather close to a louse, the specimen ran away at high speed. If a small droplet of this product was placed directly onto a louse, the parasite tried to get away from the wet region. However, even if the louse succeeded in leaving the product-covered spot, its movements stopped at a distance of about 5 mm and its legs started to have uncoordinated movements, while the intestine showed contractions. About 10 min after covering the louse by the diluted product, the slowed movement of the legs had stopped completely as well as the contractions of the intestine so that after 1 h, all treated specimens were dead, whereas the controls were alive after 2 days when kept in the feathers in a closed plastic tube at room temperature of 22 C.

Immersion assay In vitro immersion tests were carried out by Khater et al.11 using the slender pigeon louse, Columbicola columbae. The lice (per replicate) were dipped in a wrapped filter paper (9 cm diameter). Ten lice were immersed for 60 s in 100 mL solution from each concentration of tested chemical and the solution was continuously stirred during the process. Each test substance was diluted

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in water to different concentrations. The applied concentrations were 0.004%, 0.03%, 0.06%, 0.25%, 0.5%, and 1% of the test chemicals. The immersed lice were then held in the lower half of a 9-cm glass Petri dish. After 30 min, the liquid had spread out and no excess moisture was left in the dish. Two control groups were treated with distilled water only or with Tween 80 and distilled water as dependent upon the active ingredient being tested. Four replicates were used for each concentration (40 lice/concentration). Bioassays were performed at 27 C 6 2 C and 75% 6 5% RH. Lice were examined, under a stereoscope, at different time intervals (1, 4, and 24 h PT). Death of lice was defined as the lack of limb movement and failure to respond when the logs were stroked forceps. The number of deaths was recorded. The testing for in vitro LT, that is, the time-response bioassay, was similar to the standard concentration-response described earlier with the following exception; lice were exposed to a single concentration for each trial. The mortality was initially assessed 1 h after being subjected to test materials, followed by mortality assessment at 2, 4, and 24 h PT. The concentrations used were 0.01%, 0.03%, 0.06%, 0.25%, and 0.5%. The mortality data were subjected to probit transformation followed by regression analysis to determine the lethal concentration (LC) and LT values.

In vivo method(s) Efficacy studies of biting lice of ruminants should be conducted in accordance with the WAAVP guideline for the evaluation of ectoparasiticides against biting lice of ruminants (see APPENDIX A of Ref. [12]).

Experimental infestations: cattle Briggs et al.13 applied fungal conidia to louse populations that were contained in circular arenas glued to the backs of cattle. Since B750 species of fungi in 56 genera are known to be pathogens of arthropods,14 the aim of the Briggs et al.13 study was to consider whether Metarhizium anisopliae might also be a viable candidate pathogen for the control of B. bovis on cattle. B. bovis were obtained from Holstein cows. The lice were removed with forceps from the upper back, shoulders, and rump of the cattle and placed in 15 cm Petri dishes. Ventilation holes of about 2 cm in diameter were covered with fine mesh and were cut into the lids of these dishes. Stock dishes of lice were maintained at 33 C and B55% RH in a dark incubator and the lice were provided ad libitum with bovine skin scurf and hair supplemented with Brewer’s yeast. Only adult females were selected for the experiments and were used for up to 2 weeks after collection. For the study, 7 cm diameter circular arenas were glued with contact cement to the flanks of Holstein steers (12
18-months old) 24 h prior to

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treatment. The arenas were made from the rim of flexible plastic cups with cloth Stockinette enclosing the arena, surrounded by mesh (fiberglass window screen) coated in rubber cement to form the attachment surface to the hide. To maximize the contact for the adhesive between the rubber mesh and the cattle hide, cattle were shaved with an electric razor to 2
4 mm around the area where the arena was to be attached. To facilitate retrieval of the lice after each trial, the area of hair enclosed by the arena was also shaved, but to a length of 8
10 mm. Six arenas were glued along the back of each cow, with three on each side of the midline. Cattle were allocated to various trials and after each trial, the area where the arena had been washed swabbed with 70% alcohol. For each trial, two Holstein steers were maintained in individual crates in a walk-in climate-controlled room at 15 C with a 16L:8D hour cycle and were fed silage and grass hay ad libitum. Adult female B. bovis were selected at random from the stock dish as maintained earlier. Groups of 10 female lice were placed in sterile 5 cm Petri dishes, then tipped into the arena 10 min prior to treatment. The fungal suspensions and excipient were applied to the arenas using spray bottles. Sprays were applied (at a distance of B10 cm) to give a total application of 0.2 mL/arena. The Stockinette cloth covers of the arenas were then closed with metal ties to prevent contamination. Seven days PT, all lice were removed from each arena, with controls being removed first, followed by the excipient and then the fungal treated arenas. The lice were collected into sterilized 5 cm diameter Petri dishes with dampened sterile filter paper to minimize static charge. The arena was removed to locate any lice not found in the initial search. All of the recovered lice were submerged in sterile water for 2
3 min to ensure any live lice were killed; lice were then sterilized in 2% sodium hypochlorite solution and incubated at 26 C in sterile individual microtiter plate chambers, one dead louse/individual well. The plates were sealed with Parafilm and incubated at 26 C. Lice were inspected every 24 h for 3 weeks after removal from the cattle and infections were confirmed visually by the emergence of M. anisopliae hyphae from cadavers and sporulation. After this time, dead lice with no signs of hyphal growth were considered to be not infected. In this particular study, the highest concentration of M. anisopliae was effective in infecting lice.

Natural transfer: cattle One way to evaluate the persistence of a topically administered product to prevent the establishment of lice infestations is by natural transfer. An example study schedule, utilizing 32 louse-free cattle (principal animals) and 32 cattle (lice donor cattle) will include the following elements: Three weeks before the TD 0, cattle are treated topically with a shortlived insecticide to eliminate any existing lice. Daily observations for

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health problems will also occur at this time. Treatment with the short-lived insecticide can be rerepeated about 5 days prior to TD 0. The trial is a randomized block design. On TD 1, the principal animals are examined for the absence of lice and are ranked by descending body weight and consecutively assigned to replicate four animals each until eight replicates have been formed. Within replicates, the animals are randomly assigned to each of four treatment groups (in this example: Group 1—unmedicated controls; Group 2—pour-on insecticide administered once on day 0; Group 3—pouron insecticide administered once on day 7, and Group 4—pour-on insecticide administered once on TD 14). Blocks of four similar pens are formed. Replicates are randomly allocated to pen blocks and one pen within each block is randomly assigned to each animal. The principal animals are housed individually in pens that prevent contact between animals except for the 7-day exposure period. Pens are 3 3 3 m2. Donor animals are ranked according to descending louse counts of the predominant species (should multiple species be present) on TD 34 or 35. The four heaviest infested donor animals are assigned to Replicate 1, the next heaviest to Replicate 2, etc. Within replicates, one donor animal of those assigned to the replicate is randomly assigned to each pen on TD 35. Each principal animal is exposed to one donor animal for 7 days, after which time the donor animal is removed. The lice are counted on the donor animals following their removal from the principal animals. On TD 45 and 52, the lice are counted on each principal animal. Each principal animal is subjected to a thorough body search on TD 1 or 0 to confirm the absence of lice. The donors are also subjected to a thorough body search on TD 34 or 35 and on TD 42 following their removal from the principal animal’s pens and the principal animal counts are performed on TD 45 and 52 as follows: Lice will be recorded by species and at each examination, the number of lice present in the following 12 predilection sites is recorded: 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12.

Right eye Right cheek Muzzle Left eye Left cheek Dewlap Right neck Left neck Withers Topline, midway between withers and tailhead Tailhead Rump/escutcheon

2.5 cm band surrounding the eye 5 hair partings 3 15 cm 5 cm band over the dorsum of the nose 2.5 cm band surrounding the eye 5 hair partings 3 15 cm 5 hair partings 3 15 cm 5 hair partings 3 15 cm 5 hair partings 3 15 cm 5 hair partings 3 15 cm 5 hair partings 3 15 cm 5 hair partings 3 15 cm 5 hair partings 3 15 cm

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Statistical analyses of the data will be performed for each of the lice species present in the study, using ANOVA, least squares means, or other appropriate statistical models. Percent efficacy was based on geometric means. geometric mean count control group 2 mean count treated group %Efficacy 5 100 3 mean count control group

Natural infestations: cattle Potential insecticide treatments, such as pour-ons, can also be evaluated against natural lice infestations of cattle. An example is provided for a 40 animals study with 4 groups of 10 animals each. The allocation of animals to treatments and pasture lots is according to a completely randomized block design with one-way treatment structure. Experimental unit for treatment is a lot. Pasture groups will be formed based on pretreatment louse counts (mean of TD 7 and 4), so as to minimize differences in distribution of louse counts across pens. Approximately 45 cattle should be available for use in this study and acclimated to the test facilities at least 2 weeks prior to treatment. On TD 7
4, lice counts, by species, are performed on each animal and the 40 animals with the highest counts (of the lice species of most interest) and of the best health are selected for the study. The cattle are weighed on TD 3 and this weight is used to calculate the doses of the test product. On TD 1, lice counts are again conducted. Prior to treatment on TD 0, the cattle are clinically examined for overall health and treated on TD 0 in accordance with the study designs, with Group 1 being the untreated/or excipient controls. Lice counts are conducted on TD 7, 14, 21, 28, and 35 (or longer should persistent activity warrant). The counts are performed according to procedures outlined in the protocol but will most likely include some of the same predilection locations described earlier in the natural transfer section. The lice will be counted by species should multimode species be present. Louse count reduction for each species presented by treated group by time point is generated using geometric means and expressed as percentage efficacy relative to the control group mean count.

Natural or artificial infestations: sheep Module 5 (lice: insecticide resistance: diagnosis, management, and prevention) pp. 183
216 of the FAO publication “Resistance Management and Integrated Parasite Control in Ruminants” (http://www.fao.org/ag/againfo/ resources/en/pubs_ah.html) includes a description of pen trials using either sheep with obvious insecticide resistant B. ovis or groups artificially infested with suspect resistant lice. Later, B. ovis is transferred either mechanically or through contact with infested sheep. All sheep should have 2
5 lice/parting

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before treatment. Where shorn sheep are required, 1
2 cm of wool should be left to maintain the louse population. After shearing, the animals should be left undisturbed for several days prior to treatment. Prior to treatment, study groups of not less than seven sheep should be allocated which are equally weighed for mean louse counts and counts for the groups should be similarly ranked. Relevant animals should be treated with the product under investigation (for suspicion of resistance) according to the manufacturer’s recommendations. For plunge dipping, the correct volume of water must be added to the dip bath and the required volume of insecticide concentrate added to the water. This solution should be mixed for at least 5 min prior to use. For pour-ons, the equipment to be used for dosing and weighing must be calibrated to ensure proper dosage administration. Lice counts are made before and after treatment, weekly up to 8 weeks PT and monthly thereafter up to 18 weeks. Lice are counted on 10 evenly spaced partings on each side of the sheep and if no lice are found, additional 20 partings should be examined. In accordance with this FAO publication, the arithmetic mean louse count is calculated for each treated and control group and the percentage reductions in mean louse counts determined using the Henderson
Tilton formula: Rð%Þ 5 100 3

ð1 2 Ta 3 Cb Þ 3 Tb Ca

where Ta is the mean posttreatment count on treated sheep, Tb is the mean pretreatment count on treated sheep, Ca is the mean posttreatment count on control sheep, and Cb is the mean pretreatment count on control sheep.

Natural infestations: goats Garg et al.15 described in vivo studies on goats with natural infestations of Damalinia caprae where the burden of lice was assessed by summation of the total number of lice counted by using the standard counting technique in 10 3 10 cm coat openings at each of the five sites on the neck, shoulder, withers, flank, and rump regions of all goats on TD 0, prior to treatment, and then on TD 7, 14, 21, 28, 35, and 42 PT. The goats from each group were housed in separate pens; they were allowed, however, to graze daily but without contact with the other test group or mix with any other animals of the flock. The Henderson
Tilton formula was used to determine the percent efficacy after treatment. Fourie et al.16 described an in vivo study with Angora goats infested with the red louse, Damalinia limbata.

Natural infestations (avoidance assay): chickens Yoon et al.,17 using a previously validated poultry model by Ketzis et al.,18 determined the efficacy of a lice deterrent against natural infestations of chewing lice Menapon spp. and Menacanthus spp.

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In this study,17 33 adult layer hens, naturally infested with poultry chewing lice, were used. None of the chickens had been previously treated with an insecticide. Six chickens were randomly selected from the 33 birds, were treated with the active ingredient (i.e., insecticide) in the shampoo to be tested, to kill all the lice. A volume of 100
110 mL was applied to each chicken until fully covered. The feathers were moved backward and the skin and feathers were sprayed with the insecticide using a spray bottle. After 15 min, the treated chickens were washed with Dawn dishwashing liquid. Approximately 10 mL of dishwashing liquid was mixed with water in a bucket, the chicken was placed in the bucket, and the feathers gently washed by hand. The washing liquid was rinsed off with a hose, the chickens were towel dried and gently air dried with a blow drier. Once the chickens were completely dry, they were inspected to determine if all lice were killed. Lice-free chickens were assigned to Group 2 (control group) and separately housed for 2 days. These chickens were examined for lice prior to the starting of the study to ensure that no eggs had hatched and that they were still louse free. After 2 days, the remaining 27 chickens were inspected to determine the number of lice on each chicken. Ten of the 27 chickens were randomly selected and were treated with the shampoo product under evaluation. Treatment with the shampoo consisted of spraying the formulation onto each chicken until the chicken was considered fully covered (100
144 mL of product/chicken was used depending on the size of the chicken). The feathers were moved backward and the skin and feathers were sprayed with the formulation until all feathers and skin were wet with the treatment. After 15 min, a small amount of water (B20 mL) was applied to the chicken to allow the soaping agent in the product to react. The chicken was gently washed by hand with the soap from the product and completely rinsed with water. The chickens were then towel dried and blow dried and were assigned to Group 1 (test group). The six chickens from Group 2 were rinsed with water and dried as described. The remaining 17 chickens without any treatment were assigned to Group 3 (lice reservoir group). The timing of PT intervals began at the time that rinsing commenced on TD 0. All lice assessments were made by individuals blinded to the treatments. The general design was based on the WAAVP guidelines (see Appendix A for Ref. [12]) for assessing persistent activity of products for lice on cattle. To assess the number of lice on a chicken, five B2 3 2 cm areas were inspected and the number of lice was counted. The areas included the ventral and dorsal surface of each wing, the breast, the tail feathers, and the back. Initial lice assessments were performed on all chickens in all three groups at 8 h 6 18 min PT. Lice assessments were then performed in all chickens (all groups) at 24, 48, 72, 96, 120, 148, and 172 h PT. The arithmetic and geometric mean number of lice for each group were calculated. Groups 1 and 3 were compared pretreatment to determine that

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they were not statistically different using a Kruskal
Wallis test. A Fisher’s exact test was used to compare the number of chickens with lice at each time point in Groups 1 and 2 using the total number of chickens infested at each time point as well as the accumulated number.

Natural infestations: horses Two examples of in vivo lice studies on horses are provided, using two different techniques to assess efficacy. Reeves and Miller19 conducted an in vivo study using Bovicola equi in Shetland ponies. Louse infestations were measured on five Shetland ponies by counting all lice in a known area. Ponies were haltered, numbered, and 182 cm2 of hair were sheared with an electric clipper from the flank, midline, and hindquarters. The hair was clipped down to the skin and all hair clippings were collected in individual plastic bags. The sheared areas were visually checked for lice and if lice were found, they were placed in the plastic bags. The ponies were randomly allocated to control and treatment groups. Each group was isolated in pens with no physical contact with the other horses. The treated animals received the product as a pour-on applied along the back midline. After 6 weeks, the ponies were haltered and lice were counted as previously described. Animals were sheared on the opposite side. Louse infestations were compared with a chi-square test. A study by Castilla-Castano et al.20 evaluated the tolerance and efficacy of a pour-on product in a treatment and in a prevention study using natural Damalinia equi infestations in horses. In the treatment study, seven adult horses received the solution from mane to tailhead on TD 0 and four adult horses, living separately, served as nontreated controls. All were naturally infested with lice. The lice burden was recorded by counting the number of live parasites, bilaterally, over seven anatomic regions. Lesions were also assessed and scored on a 0
3 scale. Evaluation was performed on TD 0 and subsequently weekly until TD 56 in treated horses and on TD 0 and TD 56 in control horses. In the prevention study, six adult horses free of parasites were similarly treated on TD 2 and 30. Two adult horses, naturally infested with D. equi and left untreated, were mixed with the treated horses from TD 0 to 60. Evaluation was performed similarly to the treatment study on all horses, every 14 days, until TD 60.

Natural infestations: dogs Two examples of in vivo lice studies are provided, one employing experimentally infested dogs and one using natural lice infestations. Pollmeier et al.21 conducted a laboratory study using dogs that were experimentally infested with T. canis by exposing them to T. canis infested donor dogs. In this study, the principals were kept together with the donor

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dogs in an inside pen with an outside run. To assist in the spread of the infestation by close contact between animals, dogs were held in the inside pen overnight. The development of typical symptoms (itching and lesions) and the health status of animals were observed daily. Lice were counted in 38 hair coat-partings at 16 predilection sites on TD 2, 7, and weekly to TD 63. In addition, a whole-body comb count was conducted on TD 63. Counters were blinded as to treatment. The total number of live lice counted in the 38 hair coat-partings from each dog at each examination and the whole-body count on TD 63 were transformed to the natural logarithm of (count 11) for calculation of geometric means by treatment group and counting time. Louse counts were analyzed at each examination using Friedman’s test, a nonparametric analysis for a randomized block design based on the ranks of the counts in each replicate. Each treated group was compared to the control group. A significance level of 0.05 was used. Efficacy was calculated based on geometric mean whole-body comb count for each treated group relative to the control group. Endris et al.22 reported on the efficacy of a spot-on product against dogs naturally infested with T. canis. Fourteen dogs, naturally infested with T. canis, were randomly allocated to two groups (control and treatment) using lice counts determined on TD 0 prior to treatment. Dogs were ranked in descending order according to the number of lice counted. The first two dogs formed the first replicate, the next two dogs the second replicate, etc. until seven replicates were formed. Within replicates, dogs were randomly assigned to one of the two treatment groups. Treatment was administered once on TD 0. (At the completion of the trial, the untreated controls were treated to remove their lice populations.) Lice were counted in each dog before treatment on TD 0 and after treatment on TD 7, 14, 21, and 28. Counting was done by observing lice exposed as a medium-toothed comb was gently moved through the hair from posterior to anterior at six designated sites: head, tail, belly, each side, and an 8-cm strip the length of the body along the back. Lice were not removed during counting, and the total number of lice observed from all six sites was used as the index of infestation. Counts of lice at each evaluation time were transformed to the natural logarithm of (count 11) for calculation of geometric means.

Natural infestations: cats Two examples of in vivo lice studies are provided, one employing experimentally infested cats and one using natural lice infestations. Pollmeier et al.23 conducted a laboratory study using cats that were experimentally infested with F. subrostratus. The cats were experimentally infested by exposing them to F. subrostratus infested donor cats; the test subjects were kept together with the donors in cages. The development of

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typical symptoms (itching and lesions) and the health status of animals were observed daily. On TD 1, the number of live lice found in 27 hair coatpartings, located at 14 potential predilection sites or this parasite, was counted. Cats within replicates were randomly allocated to one of the treatment groups. Treatments were applied topically on TD 0. The cats were housed individually in pens that prevented contact between cats from the time of allocation to study completion. Lice were counted in 27 hair coatpartings at the 14 predilection sites on TD 2 and 7 and weekly to TD 42. In addition, a whole-body search was conducted on TD 42. The total number of live lice counted in the 27 hair coat-partings from each cat at each examination and the whole-body counts on TD 42 were transformed to the natural logarithm (count 11) for calculation of geometric means by treatment group and counting time. Lice counts were analyzed at each examination using Friedman’s test. Each treated group was compared to the control group. A significance level of 0.05 was used. Efficacy was calculated, based on geometric mean whole body counts for each treated group relative to the control group. Shanks et al.24 evaluated a spot-on product on cats naturally infested with F. subrostratus. Eighteen cats, of various ages and body weights, were used in this study following an assessment of their lice burdens. The cats were allocated randomly to either the treatment or placebo control group (excipients less active) and both treatments were administered on TD 0 as a single spot-on on the skin of each animal’s back at the base of the neck in front of the shoulder blades. The presence of live lice (nymphs and/or adults) was confirmed prior to treatment. Clinical assessments of louse infestation were repeated on TD 7, 21, 35, and 42 and the numbers of live lice present on each animal were estimated by counting them in 5 cm long coat partings made at 16 predilection sites for lice. The numbers of live lice counted on each animal on each day were analyzed by a repeated measurement analysis. A natural log transformation was applied to the total counts before they were analyzed by a mixed linear model (least square means).

References 1. Hopkins DE, Chamberlain WF. In vitro colonization of the goat biting lice, Bovicola crassipes and B. limbata. Ann Entomol Soc Am 1969;62(4):826
8. 2. Hopkins DE, Chamberlain WF. In vitro colonization of the sheep biting louse, Bovicola ovis. Ann Entomol Soc Am 1970;63(4):1196
7. 3. Hopkins DE, Chamberlain WF. In vitro colonization of the cattle biting louse, Bovicola bovis. Ann Entomol Soc Am 1972;65(3):771
2. 4. Hopkins DE, Chamberlain WF, Gingrich AR. Mallophaga: In vitro testing of artificial diets. Ann Entomol Soc Am 1976;69(3):538
40. 5. Stockdale HJ, Raun ES. Biology of the chicken body louse, Menacanthus stramineus. Ann Entomol Soc Am 1965;58(6):802
5.

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6. Maturano R, Daemon E. Reproduction, development and habits of the large turkey louse Chelopistes meleagridis (Phthiraptera: Ischnocera) under laboratory conditions. Braz J Biol 2014;74(3):712
19. 7. Sands B, Ellse L, Wall R. Residual and ovicidal efficacy of essential oil-based formulations in vitro against the donkey chewing louse Bovicola ocellatus. Med Vet Entomol 2016;30:78
84. 8. Levot GW, Hughes PB. Laboratory studies on resistance to cypermethrin in Damalinia ovis (Schrank) (Phthiraptera: Trichodectodae). J Aust Entomol Soc 1990;29:237
59. 9. James PJ, Callander JT. Bioactivity of tea tree oil from Melaleuca alternifolia against sheep lice (Bovicola ovis Schrank) in vitro. Vet Parasitol 2012;187:498
504. 10. Al-Quraishy S, Abdel-Ghaffar F, Al-Rasheid KAS, Mehlhorn J, Mehlhorn H. Effects of a neem seed extract (MiteStops) on mallophages (featherlings) of chicken: in vivo and in vitro studies. Parasitol Res 2012;110:617
22. 11. Khater HF, El-Shorbagy MM, Seddiek SA. Lousicidal efficacy of camphor oil, D-phenothrin, and deltamethrin against the slender pigeon louse, Columbicola columbae. Int J Vet Sci Med 2014;2:7
13. 12. Holdsworth PA, Vercruysse J, Rehbein S, Peter RJ, Letonja T, Green P. World Association for the Advancement of Veterinary Parasitology (W.A.A.V.P.) guidelines for evaluating the efficacy of ectoparasiticides against biting lice, sucking lice and sheep keds on ruminants. Vet Parasitol 2006;136:45
54. 13. Briggs LL, Colwell DD, Wall R. Control of the cattle louse Bovicola bovis with the fungal pathogen Metarhizium anisopliae. Vet Parasitol 2006;142:344
9. 14. Kirk PM, Cannon PF, David JC, Staplers JA. Ainsworth and Bisby’s dictionary of the fungi. 9th ed Wallingford, UK: CAB International; 2001. 15. Garg SK, Katoch R, Bhushan C. Efficacy of flumethrin pour-on against Damalinia caprae on goats (Capra hircus). Trop Anim Health Prod 1998;30:273
8. 16. Fourie LJ, Kok DJ, Allan MJ, Oberem PT. The efficacy of diflubenzuron against the body louse (Damalinia limbata) of Angora goats. Vet Parasitol 1995;59:257
62. 17. Yoon KS, Ketzis JK, Andrews SW, Wu CS, Honraet K, Staljanssens D, et al. In vitro and in vivo evaluation of infestation deterrents against lice. J Med Entomol 2015;52(5):970
8. 18. Ketzis JK, Clements K, Honraet K. Use of a poultry model to assess the transfer inhibition effect lice (Pediculus humanus capitis) products. Parasitol Res 2014;113:1943
8. 19. Reeves WK, Miller MM. Control of Bovicola equi (Phthiraptera: Trichodectidae) with dimilin and permethrin. J Vector Ecol 2009;160. Scientific Note. 20. Castilla-Castano E, Vischi A, Navarro C, Lecru LA, Ribeiro C, Pradier S, et al. Control of lice infestation in horses using a 10 mg/mL deltamethrin topical application. Irish Vet J 2017;70:22 Available from: https://doi.org/10.1186/s13620-017-0100-2. 21. Pollmeier M, Pengo G, Jeannin P, Soll M. Evaluation of the efficacy of fipronil formulations in the treatment and control of biting lice, Trichodectes canis (De Geer, 1778) on dogs. Vet Parasitol 2002;107:127
36. 22. Endris RG, Reuter VE, Nelson J, Nelson JA. Efficacy of a topical spot-on containing 65% permethrin against the dog louse Trichodectes canis (Mallophaga:Trichodectidae). Vet Ther 2001;2(2):135
9. 23. Pollmeier M, Pengo G, Longo M, Jeannin P. Effective treatment and control of biting lice. Felicola subrostratus (Nitzsch in Burmeister, 1838) on cats using fipronil formulations. Vet Parasitol 2004;121:157
65. 24. Shanks DJ, Gautier P, McTier TL, Evans NA, Pengo G, Rowan TG. Efficacy of selamectin against biting lice on dogs and cats. Vet Rec 2003;152:234
7.

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Chapter 3f

Arthropoda, Insecta, Siphonaptera Alan A. Marchiondo, MS, PhD1, Larry R. Cruthers, MS, PhD2 and Daniel E. Snyder, DVM, PhD, Dipl. ACVM3 1 Adobe Veterinary Parasitology Consulting LLC, Santa Fe, NM, United States, 2LCruthers Consulting, Chesapeake, VA, United States, 3Daniel E. Snyder, DVM, PhD, Dipl. ACVM, Consulting LLC, Indianapolis, IN, United States

Arthropoda Insecta Siphonaptera

Pulicidae Ctenocephalides felis felis Bouche´, 1835—cat flea Biology and life cycle The cat flea, Ctenocephalides felis felis (Bouche´, 1835), is a near ubiquitous problem globally being found in Australia, European Union (Austria, Czechoslovakia, France, Germany, Ireland, Poland, United Kingdom of Great Britain, and Northern Ireland), Japan, Scandinavia (Denmark and Sweden), Africa (Egypt, Western Cape, and provinces of Gauteng, North West, Limpopo, and Mpumalanga), South America (Argentina), and the United States including Puerto Rico on both cats and dogs. The dog flea, Ctenocephalides canis (Curtis, 1826), is largely confined to dogs and more restricted in its distribution to the European Union (Austria, Czechoslovakia, Germany, Great Britain, Ireland, and Poland), Japan, Scandinavia (Denmark and Sweden), South Africa, South America, and United States (Indiana, Florida, and Georgia).1,2 Both adult cat and dog fleas are characterized by caudally directed spines distributed over their body and a genal as well as a prenatal comb. But cat fleas have a head that is longer than it is tall; the first and second genal spines are of almost equal length.3 Dog fleas (C. canis) have a head that is as tall as it is long, and the first genal spine is Bhalf the length of the second spine. The fleas of dogs and cats are wingless, laterally compressed, obligatory hematophagous insects of the order Siphonaptera (Greek siphon 5 “tube” or “siphon” 1 a 5 “not” 1 ptera 5 “wing,” “sucking-wingless”). Adult cat fleas of both sexes are generally 2
5 mm long, with females being larger than males. Female cat fleas tend to develop before male fleas (protogony).4 Male cat fleas can be differentiated from female fleas by their generally smaller size and the presence of the prominent male clasper.5 The abdomen of the male cat flea has a more abrupt sloping shape as compared to the female flea. Female cat fleas have a larger abdomen with more caudally placed sensilium and postanal hairs.

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The life cycle of the cat flea is holometabolous; it involves complete metamorphosis, meaning a pupal stage occurs during development.6 A blood meal is required by both sexes to initiate reproduction, with mating and ovipositioning beginning initially within 20
36 h after feeding on a host. Female cat fleas ingest B10 times more blood than male fleas during 24 h time periods.7 Typically, all developmental stages of the cat flea life cycle take place in the animal’s local environment and are present at the same time. Adult cat fleas on the pet account for B5% of the total population. The balance of the flea population is made up of the developing stages (B50% eggs, B35% larvae, and B10% pupae) in the animal’s environment. The developmental cycle of the cat flea from egg to adult occurs within the ranges of 13 C
32 C, 50%
92% RH.8 The entire life cycle of the cat flea can be completed as early as 12
14 days or as long as 140
190 days depending on temperature and humidity.9,10 Under ideal laboratory environmental conditions (18.3 C, 70% RH), the entire life cycle of the flea can be completed in as little as 3 weeks. Outside the laboratory under most household conditions, the majority of cat fleas complete their entire life cycle within 3
4 weeks. Preemerged adult fleas in cocoons (pupal cases) rapidly emerge in response to stimulation and the newly emerged adult fleas will then jump onto a passing host (heated target) initiating an infestation. Fleas possess specialized, powerful legs being able to jump 34 cm with direction, but not precision.11,12 Unfed and fed adult cat fleas exhibit positive phototaxis and negative geotaxis.13 Newly emerged unfed adult fleas begin to seek a host for a blood meal almost immediately after emerging from the cocoon3,14 but can survive outside of the cocoon waiting for a host for 20 days in cool, dry air, and 62 days in humid conditions.10 About 95% of adult cat fleas die within 15 days at 24 C, 78% RH13 and within 12.3 days under ambient conditions averaging 22.5 C, 60% RH.3 No life cycle stages can survive for 10 days at 3 C or 5 days at 21 C.15 Upon infesting a suitable host the adult cat flea starts taking a blood meal within seconds, and a female cat flea can consume an average of 13.6 6 2.7 μL of blood/day, about 15 times their body weight.13,16 About 98% of adult female fleas feed within 12 h of being placed on a host.13 The maximum time of feeding of the cat flea is 5
10 min.17 The blood feeding is necessary for oviposition as well as for successful mating and nutrition,18 and male fleas feed less frequently than females.17 The mating of cat fleas occurs on the host within the first 8
24 h, with most female cat fleas having mated by 34 h.19,20 Adult female cat fleas begin to lay eggs on the host within 36
48 h after taking a blood meal, reaching maximum production between 4 and 9 days with the capability of producing eggs over 100 days.21 A female cat flea can lay as many as 11
46 eggs/day with over 2000 eggs being laid in her lifetime.13,21
24 The adult fleas are permanent ectoparasites on the host with an average of 85% of females and 58% of males staying continuously on a cat

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for at least 50 days,21 unless the population approaches the threshold of about 200 fleas. Along with the laying of flea eggs, flea feces (also called flea dirt or flea frass) that consists of undigested blood components is also deposited into the environment and which subsequently serves as the principal source of food for the larval stages. The cat flea eggs are nonsticky with a smooth surface, slightly transparent to pearly white, widely oval with rounded ends, and measure 0.5 3 0.3 mm. The eggs fall off the host along with the flea feces within 8 h of oviposition,25 develop, and hatch in 1
10 days depending on temperature (50% egg hatch at 35 C) and RH (80% egg hatch at RH ranges of 50%
92%). The free-living cat flea larvae measure 2
5 mm having a white body with yellow-to-brownish head but become brownish in color after ingesting dried blood feces deposited in the environment from the feeding adult fleas on the host.4 The larvae have 13 body segments,26 3 thoracic segments and 10 abdominal segments27 with chewing mouthparts.28 The cat flea larvae are negatively phototactic and positively geotactic.29 They undergo two molts, thus having three instars that progress in size and length with the third instar measuring B4
5 mm in length.30,31 The cat flea larvae have a minimal nutritional requirement of dried blood to develop.32 The flea larvae are thigmotactic (that is, they recognize tactile stimulation and react to mechanical contact).33 Development of the larvae varies from 5 to 11 days depending on temperature and RH (,50% RH causing desiccation just as is the case with egg development).9,13,34 The third instar spins a pupal cocoon composed of silk with the outer surface becoming coated with debris from its surroundings.34
36 Pupation is triggered by declining levels of juvenile hormone.37 Interestingly, when larvae were not allowed to orient against a perpendicular structure, most larvae failed to spin an enclosing cocoon, yielding what has been termed “naked flea pupae” with 95% surviving and developing to adult fleas.38 The larva within the cocoon undergoes two molts to complete its development to the adult stage. The female fleas pupate within B32 h, while males pupate within B44 h, B12 h later than females at 27 C 6 2 C.38 The females develop into adults about B1.6 days faster than males.38 Within the cocoon the flea develops within 7
19 days into an adult.9,35 The adult flea then emerges from the cocoon upon proper stimulation (vibrations, physical pressure, CO2, and heat). The adult cat fleas begin to emerge as early as 3
5 days following pupation and reach a peak in 8
9 days at 27 C and 80% RH.9,39 The female adult fleas generally emerge from the cocoons 1
5 days before the males.

Rearing method(s) Maintenance of cat fleas on animals Maintenance and rearing of a laboratory colony of cat fleas are essential for conducting parasiticide screening tests. This can be done in-house or the

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different flea life cycle stages, in particular unfed adult fleas, can be purchased from vendors who maintain the life cycle. The two major laboratory methods of cat flea rearing involve maintaining a flea colony on a suitable host, such as the cat (the natural host of C. felis), or developing methods for rearing cat fleas on an artificial feeding system. Numerous procedures have been developed for the laboratory rearing of cat fleas on cats39
43 and on artificial feeding systems.44
47

Production of Ctenocephalides felis on cats The purpose of this procedure is to describe a method for the production of C. felis (cat flea) on cats. Choice and husbandry of animal Male or female domestic short hair cats can be used for laboratory flea rearing. The utilized cats, negative for feline leukemia, can be neutered or spayed if the facility requires these procedures and preferentially declawed (to reduce excessive self-grooming). When choosing a cat for this procedure, it is important to select a cat that will use a litter box. The cats that will not use a litter box will ruin the flea production “chamber” (see description later) due to urine contamination, and the weekly collection of flea eggs will be lost. Weigh the selected cat prior to placing in the chamber of appropriate size based on the size of the cat. A physical examination of a flea donor cat should be performed by an experienced veterinarian and will include, but not be limited to, rectal temperature, thoracic auscultation, skin and hair coat assessment, including any signs of FAD, and the assessment of their general physical condition. The cats, which are identified as unsuitable by the physical examination, should not be used. The cats may be treated with a commercial anthelmintic, but not any topical or systemic ectoparasiticides, and vaccinated to protect against common bacterial and viral pathogens. The cats used for flea rearing duties will usually remain in the flea chamber for 4
6 weeks as a “colony status” animal, and at least 6 weeks as an “off colony” animal, where they will be reentered into the colony rotation after the minimal 6 weeks of resting period. Any cat that displays severe pain or distress (exhibited as depression, anorexia, indolence, or loss of body weight) or cannot acclimate to the flea chamber will be removed immediately from the flea chamber, removed from “colony status” and colony rotation, treated with the proper medications and/or ectoparasiticides or, as a last resort, euthanized, if necessary. The cats may be given canned food containing liver (in lieu of commercial dry food), while in the flea chambers, and are given water ad libitum. Each cat should be given 1 cc of vitamin B12 and 1 cc iron by IM injection. These injections should be given when the cat has completed its time in

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the flea production chamber before returning the animal to the animal colony. Each cat should be weighed and washed with a shampoo that will rid the cat of the fleas. After the cat is dried, it should also be combed with a flea comb to ensure that all fleas are removed before being placed back in the colony. Treatment with an ultrafast acting, highly effective, and short acting pulicidal insecticide may be used to insure that all fleas are killed prior to putting them back into the general cat colony. Animal care and welfare is governed by SOPs of the facility, national animal welfare regulations, and other applicable legal and national regulatory requirements. Flea chamber setup The construction of the chamber, which serves as an indoor housing unit, must meet the standards of any applicable animal welfare regulations and on site IACUC or other animal welfare approval processes. A number of stainless steel cages with grated floor chambers have been commercially developed in the United States and have flea collection jars underneath (Fig. 3f.1) (Percival Manufacturing Co., Boone, IA; Unifab Corporation, Kalamazoo, MI; Winkle Manufacturing, Glen Elder, KS). Almost all chambers are based on the basic design by Hudson and Price.39 The chamber herein described is B3 3 3 3 3 ft2 metal chamber (box) with a funnel-shaped bottom with a collection hole that is open or connected to a collecting jar so that flea eggs and flea feces fall from the coat of the cat into a collecting vessel beneath. A metal grate floor (made of 0.5 in. rubber coated expanded metal) is inserted. The chamber is fitted with a metal grate that serves as the “lid” or top of the chamber. The chamber is equipped with a resting shelf or board (made of 0.5 in. rubber coated expanded metal) for the cat to perch, a litter box as well as a food and water bowl (Fig. 3f.2). FIGURE 3F.1 Commercial cat flea cages manufactured in the Unites States with collecting jars underneath. Personal photograph by AAM.

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FIGURE 3F.2 Inner area of a cat flea chamber equipped with a resting shelf (board) for the cat with food and water bowls. Personal photograph by AAM.

Ground corn cobs, 0.25 in. diameter, sifted through a No. 12-mesh sieve (1.68 mm) or other suitable commercial material is used as litter. Start with an empty, clean, dry chamber. Place a collecting tray, jar, or pan under the collection hole or under the metal grate flooring. A cat is placed into the chamber and infested with adult fleas. The cats may or not be fitted with an Elizabethan collar to further reduce grooming activity and prevent them from ingesting adult fleas. Colony cats are infested Bonce/week with as few as 50
200 adult fleas (ratio of 1 male:3 females or 50:50 based on counts of 3 flea samples of 100 fleas each) applied on the nape of the neck. Any cat that exhibits signs of FAD such as hair loss will only be infested with up to 100 fleas every other week or not used. Replace the chamber lid. After the cat has acclimated to its flea burden, place a water bowl into the chamber. Expose the colony chamber to at least 12 h of light/ day.

Collection of flea life cycle stages Flea egg collection and larval maintenance procedures At B1 week PI after the cat is first placed in the flea chamber, material (containing the flea eggs) from the collection container (pan, jar, and tray) is collected daily. Prior to each daily egg collection procedure, the cat is manually brushed/rubbed by vigorously moving one’s gloved hands over its entire body to remove shed hair and dislodge eggs and flea feces so they fall into the egg collection device. Flea egg collections are made at B24 h intervals from each cage containing a colony animal. The cat is then removed from the chamber. Bowls, resting board, and litter box are removed, cleaned, and sanitized (5% sodium hypochlorite and water). This cleaning procedure should be done minimally at least once/week or as according to the facility SOPs while the cat is being maintained in the flea chamber.

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Using a paintbrush or other suitable object, any material adhering to the subflooring or slanted walls of the flea collection chamber is brushed through the collection hole into the dry pan/tray/jar. The chamber and grates are cleaned and sanitized while the cat is being maintained in the flea chamber. The daily collected debris and material obtained from the collection pan/ tray/jar can be transferred to a different clean container for transport to the laboratory for further processing. The flea eggs and flea feces are separated from the litter, and other unnecessary animal or food debris using a No. 35mesh sieve (500 μm) or No. 40-mesh sieve (420 μm) stacked over a No. 200-mesh sieve (74 μm). The eggs are collected in the bottom sieve to be transferred to a Petri dish as described below. If the collected material from the flea collection chamber is damp, collect it and dry it for several hours before beginning the sieving process to isolate the flea eggs. However, if the flea collection pan/tray/jar is soiled with cat urine or excessive water and is deemed unsuitable, do not collect eggs from this daily collection for this cat and replace with a clean pan/tray/jar under that collecting chamber. Cover the bottom of an empty 9 cm diameter Petri dish with clean, fine (playground type) sand and cover the sand with dried blood or alternatively add B180 mL or 1
3 mm deep of flea rearing medium (see the section on flea rearing media), mixed thoroughly and placed into a labeled (collection date and animal identification) Petri dish. Appropriate metal pans or trays containing flea rearing medium may be substituted for the Petri dishes during the flea rearing process, if desired. Add the daily collection of flea eggs for each cat to the Petri dish and cover with the lid. Egg collections from multiple cats can be combined into a single Petri dish, if desired. However, for tracking/record keeping, it may be advisable to use eggs from the same cat, thus enabling one to go back and eliminate a cat from the colony if not a good producer. The Petri dishes containing eggs should allow adequate ventilation if covered. These Petri dishes may be further placed in properly labeled (collection date, cat identification) pans/trays to segregate daily flea collections prior to being transported to and placed into an incubator/growth chamber or other suitable environment. The incubator or growth chamber should be set at and constantly maintained at an optimum humidity and temperature, which should be checked daily, for optimal flea development and survival. A daily log for the flea insectary to record relevant environmental and flea colony data should be maintained as well as a log for flea colony animal usage. After flea eggs are collected from the sieves, they can be further cleaned with a small, fine brush in a Petri dish. Flea egg enumeration can also be performed in a Petri dish lined with gridded construction paper, if desired. Eggs hatch in 1
3 days, and larvae feed for 6
8 days. Larvae (75%
85%) hatch within 48
72 h. Once larvae are observed in the Petri dish, flea rearing medium is added as needed until pupation has ceased. The larvae (late second or third instars) can be separated from the rearing medium during the 4
5 days using a No. 20-mesh sieve (0.841 mm). These larvae can

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then be used in larval bioassays. The pupation begins 7
8 days after larvae hatch from the eggs. Flea pupae collection procedure Flea pupae can be collected at 3, 6, 9, 13
16 days past the egg collection date. Place a No. 16-mesh sieve (1.19 mm) to a No. 20-mesh sieve (0.841 mm) into a litter pan and pour the contents of one flea egg collection Petri dish into the sieve. Gently detach pupae adhered to the Petri dish with a fine brush and pour them into the sieve. Tap the bottom of the sieve until only pupae and larvae remain in the sieve. Recovered larvae can be returned to the Petri dish with their original rearing media to allow them to pupate. Pupae can develop with or without a cocoon.38,48 Place a funnel into a graduated cylinder and pour the pupae collected from the sieve into the funnel. Place a maximum of 10 mL or up to 1000 pupae per container (1 mL 5 B80
85 pupae) or B1/2 in. (one finger width) of pupae in a 7 dram vial. Label each container with the dates of egg and pupal collection and Bnumber of pupae contained. It is advisable to weigh and record each cat’s pupal production for historical purposes. Return the vials (and pans if more collections need to be made) to the incubator or other appropriate locations. Keep collecting from the pans at least four times or until all the larvae have developed, and all the pupae have been collected. After the last collection the remaining medium should be discarded. Keep the pupae collections from each cat separate and record the following information: how many fleas were infested on each cat, how many grams of pupae resulted from each infestation, and any pertinent information affecting the cat colony. Adult flea collection procedure Adult fleas begin emerging from pupae 7
10 days after the initial pupation. Female fleas typically begin emerging around day 16 with males emerging 3
4 days later. Emergence occurs over several days. Total adult flea emergence from eggs should be .50%. Typically, total adult flea emergence will run from 60% to 65%. Choose pupae incubation containers from incubator that are ,43 days past their egg collection date. The adult fleas can be collected from pupae by two methods. In the first method, cocoons (pupae) are placed in specially constructed emergence jars consisting of an open container suspended 15
20 cm above the bottom of the jar (Fig. 3f.3). A flea collecting apparatus has also been described with a useful diagram,49 and a similar method was used by Gilbert.43 As the fleas emerge from their pupal case, they jump to the bottom of the jar and are separated from the cocoons. A few fleas may remain in or on the suspended lid. Before opening the emergence jar the fleas can be immobilized with CO2.

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Parasiticide Screening, Vol 1 FIGURE 3F.3 Adult flea emergence jar with pupae in the suspended lid and emerged flea in the bottom of the jar. Personal photograph by AAM.

A more sophisticated version of this method employs an oblong plexiglass box on a stand with a covered opening on top of the box to place incubation chambers of cocoons onto a shelf. The emerged adults jump from the chambers onto a platform. The adults can then be emptied into a collecting dish. In the second method, pupae in incubation container(s) are placed in the bottom of a flea aspiration chamber at least 30.5 cm (12 in.) high plastic container (5 gal plastic bucket works well) to prevent escape, and the lid of the incubation chamber is removed to allow the fleas to jump out into the aspiration chamber. When fleas are needed for in vitro or in vivo studies, connect the outlet/inlet hoses from the vacuum pump to the flea collection test tube. Using the vacuum hose attached to a pipette, begin aspiration by placing the pipette above the target flea and count as they enter the air intake. Take care not to place the pipette over fleas that are closer than B1 cm apart to avoid aspirating more than one flea at a time. Only aspirate fleas that are capable of standing on their legs in a vertical position. Aspirate 100
200 adult fleas in test tubes for the infestation of colony animals or for other research needs. If it is used for research purposes or for efficacy studies on dogs or cats, use newly hatched adult fleas that are ,14 days of age for consistency of results. Newly emerged fleas can be held 7
11 days in ventilated test tubes with 95% dying within 15 days at 24 C and 78% RH.

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Incubator temperature and relative humidity The incubator used to store the pans and flea pupae should be kept at 27.8 C 6 5 C with at least 80% RH (optimum RH .85%). The minimum temperature in the room containing the cat flea production chambers should be $ 21 C and $ 50% RH. The temperature and humidity in the chamber should be maintained at 27 C
28 C and 80%
85% RH. Supplemental heaters and/or humidifiers are used as needed. Flea rearing medium Various flea rearing media diets have been described.39,41,43 Most diets consist of pet or rodent chow, dried blood, and Brewer’s yeast with or without sand. Moser et al.50 showed that dried blood alone performs as well as any other diet. Once a flea rearing diet recipe and procedure has been established in a laboratory, it must not change, and any deviation may result in the failure of the flea colony. Anticipate your needs so as not to run out of needed materials. The recipe herein provided is well tested and reliable. 1. Dog food: Before placing dry dog food (any good quality brand such as Purina Pro Plan) in a blender, it must first be crushed in a grinder, with a hammer, or similar object, or else the blender may be damaged. Grind the dog food in the blender at high speed. Do not put more than onefourth cup of food into the blender at one time (just enough to cover the blades). For efficiency of time, grind 1 month worth of meal at a time (B3000 g). 2. Pass the ground dog food through a No. 25 sieve (0.707 mm). After several passes through this sieve, discard that which will not pass through. Push the ground meal through the sieve using a paintbrush. Sieve holes will get plugged with meal, but consistent knocking and pushing of meal through the holes will result in success. 3. Dry bovine blood is commercially obtained (California Spray Dry, Stockton, CA)51 or collected fresh from a slaughter house, placed in large metal trays (B1 cm deep), and dried for several hours at 50 C.52 The dried blood is ground or hammer milled (Straub model 4E grinding mill) and passed through a No. 30 (0.595 mm) or No. 35 (0.500 mm) sieve to produce a uniform-sized particle. 4. Brewer’s yeast is commercially provided in powder form. 5. Sand (playground) is sifted through a No. 18 (1.00 mm) and a No. 20 (0.841 mm) mesh sieve. Autoclaving the sand prior to mixing with larval rearing medium will greatly reduce or eliminate fungal growth. To make flea rearing medium, weigh out: a. 100 g ground dog food, b. 15 g ground blood, and c. 10 g powered Brewer’s yeast. Always use this ratio when making this flea food recipe.

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Another recipe with sand is as follows: Larval food

Dog chow Dried beef blood Brewer’s yeast

20 parts (weight) 5 parts (weight) 2 parts (weight)

Sand: Larval food ratio of 4:1 (volume) Flea rearing problems 1. Mites Mites can kill the colony, therefore always be on the watch for the mites. Upon completion of sieving each day, all sieves should be thoroughly cleaned with warm soapy water. If mites are present, the outside of the pans should be wiped with 3% hydrogen peroxide before being returned to the incubator. The pans should be washed with warm soapy water after the final collection has been completed. The incubators should be cleaned at least once a month with 3% hydrogen peroxide. If mites are discovered, the incubators should be cleaned more often. The ingredients other than 3% hydrogen peroxide or hot water may contain harmful chemicals that may harm the flea colony. If mites are present in the incubators, any content in the incubators should also be wiped down with 3% hydrogen peroxide. The mites feed on mold, and this is the beginning of mite problems. Should mold occur in any pan, throw away the spoiled contents. In order to kill the mites the incubator must be completely dried out and run for B48 h with a desiccant to kill the mites. 2. Small fleas This is usually caused by not using the correct sieve size during the preparation of the flea food mixture. If too large a sieve is used, the food particles are too large for the larval flea’s small mouthparts. This results in embryo change and small fleas that could ultimately result in the loss of the colony. Colony rejuvenation Periodically (every 6
10 years unless dictated otherwise by a regulatory agency), wild sourced fleas (outside sources from untreated cat or dog) should be introduced into the flea colony to keep the colony viable and to introduce new genetics. This information should be duly recorded in rearing records. Rearing Ctenocephalides felis felis on mice A mass rearing method of C. felis fed on mice has been developed using an apparatus to house flea infested mice on a 50 mesh wire net that allows flea eggs to fall through and be collected. Duration from egg to cocoon formation

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of C. felis fed on mice was about 8 days, that from egg to adult emergence was about 18 days, and the accumulated emergence rate was 89.2%. The average number of eggs deposited was 10.3 eggs/day/female flea; maximum egg deposition was 21.6 eggs/day/female flea at 4 days after infestation, and total number of eggs produced was 402 eggs/female flea. The longevity of C. felis reared on mice was more than 40 days. This rearing method is very simple and useful for producing a large number of cat fleas in a small space.53 Rearing Ctenocephalides canis on dogs The biology, life cycle, and rearing methods of C. canis are similar to those used to rear C. felis felis on cats and have been used to successfully establish colonies of the dog flea on dogs in the laboratory.54 Artificial dog The name “artificial dog” was coined by Cornell University News Service reporter Roger Segelken for the first fully successful artificial feeding system for adult fleas described in 1988.47 The late Dr. Jay R. Georgi, Professor Emeritus of Parasitology, and Dr. Susan E. Wade, Cornell University, College of Veterinary Medicine, adopted the name and invented the original artificial dog, which was based on an apparatus designed to feed mosquitoes and underwent three variations in cage design all of which were published47 and patented.55,56 The system for breeding fleas comprises a transparent acrylic box, about 16 in. on a side, with an electric heater to warm the fleas (38 C
40 C) and the citrated bovine blood (35 C
40 C) that the fleas imbibe (replaced fresh B24 h). Depending on the desired

FIGURE 3F.4 The “assay type” artificial dog accepts 104 1-in. cages and is suitable for rearing fleas and testing parasiticides for activity against adult fleas. Personal photograph by DES.

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rearing chamber capacity, 20 or 25 2-in., 40 2-in., or 104 1-in. cylindrical rearing cages (Fig. 3f.4) are constructed within the box, with slightly less than 300 fleas per cage. The flea rearing output from the 20 or 25 cages was 11,000
14,000 fleas/day. The final version of the cage consisted of a top screen with a 300 μm nylon feeding screen cemented to the underside of the cage top, a removable support nylon screen with 1000 μm openings, and a bottom screen with 50 μm openings. The support screen rested on a ledge wall about 0.08 in. deeper than the male part, thus forming a “feeding space” between the feeding screen and support screen, which themselves have a combined thickness of 0.03 in. The cage had a hole in its bottom half for loading the fleas. The cage could be placed under negative pressure, and fleas introduced through a tube inserted in the hole, which was closed with a screw (nylon, no. 3
32) at completion of the loading operation. An aluminum cylinder filled with citrated bovine blood and covered with Parafilm, an artificial skin (feeding membrane) through which the insects feed, fits on top of the rearing cage. The feeding membrane can be made by grasping a 1 3 4 in. piece of Parafilm between the four fingers and the side of the thumb of each hand and then stretching it about 6 in. wide. Then allow the Parafilm to relax, applying it to the open end of the blood-feeding chamber with just enough tension to remove any wrinkles, and press the overlapping Parafilm against the sides of the chamber to secure the membrane in place. The surplus Parafilm can be trimmed off with a razor blade or sharp knife. The fleas feed on the upside through the Parafilm membrane, and their eggs and feces are collected at the bottom in plastic dishes filled with sand and culture medium. The egg and larvae collections are allowed to develop at 27 C and 80%
85% RH in a separate incubator. Emerging adult fleas are collected within 14 days. A vacuum stage or system (actually a vacuum cleaner) can be used to restrain the fleas. An airtight box with a 2-in. diameter hole (cage port) on top and another hole on the side to receive a vacuum cleaner hose serve well for installing the fleas in 2 in. cages. The cage port can be reduced to accommodate 1 in. cages. With a flea cage over the vacuum port and the vacuum cleaner creating a high wind in the vicinity of the cage bottom, any flea introduced will be held against the bottom screen on their sides. The vacuum stage clamps on the side of an aquarium tank, enamel bucket, or a 5 gal plastic tub. The inside of the tub can be wiped with an antistatic sheet (fabric softener) to keep the fleas from sticking to the wall. Fleas released in the bottom of the tub are transferred by means of a tube against the bottom on the vacuum port. Merely holding one end of the tube against the bottom screen if the cage provides sufficient airflow, this allows one to move the fleas from the bottom of the tub to the cage bottom where they are held flat on their sides by the vacuum. Install the cage top, once the cage has been loaded with fleas. Dr. Georgi concluded the fact that fleas feeding on cats produces so many more eggs than do fleas fed artificially and suggests that considerable

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room remains for improvement in this artificial rearing system.57 Nevertheless, the artificial dog serves as a reliable and productive system for rearing adult fleas off of a mammalian host and subscribes to the guiding principles underpinning the humane use of animals in scientific research called the three Rs58: (1) replace the use of animals with alternative techniques or avoid the use of animals altogether, (2) reduce the number of animals used to a minimum, to obtain information from fewer animals or information from the same number of animals, and (3) refine the way experiments are carried out to make sure animals suffer as little as possible. The artificial dog membrane feeding system47 has undergone various modifications, particularly for evaluating parasitcides.59
61 However, the basic design principles of the model by Wade and Georgi47 remain the same. The use of the artificial dog membrane feeding system for the testing of parasiticides will be covered in the “Membrane feeding—artificial dog feeding chamber—adult fleas” section.

In vitro method(s) In vitro techniques are often used in the discovery and preclinical phases of the development of potential pulicidal compounds. Methods are available to determine LC50 values62 by contact exposure, for example, of adult fleas to a range of concentrations of the investigational chemical in or on liquid or solid media, such as impregnated filter papers or whole blood. Such studies can provide useful data on the spectrum of potential insecticidal activity, on synergistic activities, or on resistance profiles, but they rarely give more than a broad indication of the dosage required for topical or systemic application to an animal.2 Contact bioassay Topical application—adult flea A topical (microdroplet) bioassay has been described for the cat flea.63 Adult fleas are immobilized with CO2. Dilutions of a test compound are applied to the individual fleas in 0.1 μL acetone. Mortality is recorded 24 h later, and LC50 and LC95 levels are determined. LD95 values in ng/flea are determined for the test compounds. Experiments repeated 2
5 years later, with five of the previously tested compounds, in a different facility, and showed only small shifts in potency (0.38 to twofold of the original LD50 values).64 Filter paper—background The filter paper (contact) method in which acetone solutions of test chemicals are used to treat filter paper to which fleas are subsequently exposed was first published by Burden and Smittle.65 This method employs the use of filter paper disks or strips66 treated with known concentrations of a test compound in a glass vial containing newly emerged adult fleas. Two major factors influence the reliability of continuous

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exposure testing on treated filter paper: (1) RH and (2) the solvent(s) used to solubilize and deposit the test compound. First, RH can affect the toxicity of the test compound with the activity of chlorpyrifos increasing as the RH increases.67 On porous substrates, such as filter paper, the water vapor has a greater affinity for the paper than does the test compound.68 Second, the solvents used to treat filter paper must completely dissolve the test compound which will have its own specific physicochemical characteristics. Single solvents, such as acetone, or mixtures of solvents may be required. It may be desirable to use a highly volatile solvent that evaporates quickly leaving a fine residue of the test compound on the filter paper. To secure uniform contact between insect and chemical, oils were recommended earlier for specific test compounds69: olive oil for organophosphates and carbamates,70 and silicon oil for pyrethroids.71 In some cases the test compound might become impregnated into the filter paper, in which case, the activity of the test compound will increase as residue migrates out to the surface over time. The effectiveness of a test compound depends very much on the test method. Azamethiphos was highly effective against the fleas in topical and nylon carpet tests, whereas it was ineffective in the filter paper test.72 Finally, cat flea isolates may vary in their sensitivity to insecticides as the El-Labs source (Soquel, CA), isolated in 1969 that are reared and fed on an artificial system, tend to be much more sensitive to toxicants than other isolates.73 Filter paper—adult fleas Containers of developing and newly emerged adult fleas collected from the artificial dog or in vivo from flea rearing systems on cats are placed in a cylindrical chamber (5 gal plastic bucket) that allows the fleas to jump out of the containers into the chamber. The desired number of live fleas are aspirated and counted from the chamber into a manometer tube and transferred to a test tube. A small piece of screen is placed in the test tube to provide a resting area. The tube is stoppered to prevent the escape of fleas and to allow air exchange. A stock solution of the experimental compound is prepared by dissolving the active ingredient in a solvent solution (examples include acetone, DMSO, and dimethylformamide) or ratio mixtures of solvents such as dimethylformamide:acetone (1:9) to ensure the active ingredient is well dissolved. An aliquot of 20 μL of stock solution is added to 380 μL of DI water and mixed thoroughly to yield a solution of 100 ppm concentration of the active ingredient. The 100 ppm solution is then serially diluted with DI water to yield concentrations of 25, 6.25, 1.6, and 0.4 ppm. This dilution series is only an example as the serial dilution concentrations will vary based on the potency of the investigational compound. Then duplicate filter paper disks (Whatman Hardened Ashless, Cat. No. 1540 321, 21 mm diameter) are treated with

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75 μL of each of the prepared dilution series and allowed to air dry overnight at RT in a ventilation hood. Duplicate untreated and blank solvent treated filter disks are used as a control standard. For each dilution, one of the duplicate filter disks is placed into the bottom of a Wheaton Liquid Scintillation Vial (22 mm), and the other filter disk is placed into the cap of the vial. Replicates of each dilution for use in multiple scintillation vials can be performed as well. Newly emerged adult cat fleas (B10) are added with a flea vacuum device to each container, and then the vial cap is securely closed. The vials are incubated at 27 C and 80% RH. Flea mortality (number of fleas dead or unable to make forward motion) is assessed at 24, 48, and 72 h (or longer) intervals. Lethal probit concentration values are determined. Alternative filter paper methods The treated Whatman filter paper disks (3 cm diameter) are inserted into the drilled-out top of a 7-dram plastic or glass vial and the other disk placed inside the vial. The vials are incubated at 27 C and 80% RH. The flea mortality is assessed every 24 h for 4 days. On TD 5 the vials are opened and the precise mortality recorded. The lethal probit concentration values are determined. The contact filter paper bioassay can be modified for other treated substrates, such as glass cover slips, plastic or glass Petri dishes, dog hair,74,75 fiber glass, woven woolen fabric,76 and nylon.77 Filter paper—flea eggs The cat flea eggs are evaluated in vitro to determine the effect of test compounds (adulticides and IGRs) directly (contact exposure) or indirectly (exposure from the treated host or uptake by the female flea).59,78 The flea eggs are collected as previously described in vivo from a colony cat or in vitro from an artificial dog membrane feeding system. The membrane feeding system is useful to obtain flea eggs of a known age. The flea eggs are collected from trays or other collection devices placed underneath cats naturally or experimentally infested with fleas. The eggs and debris are passed through a series of sieves No. 10-mesh (2.00 mm), No. 16mesh (1.19 mm), No. 20-mesh (841 μm), and No. 60-mesh (250 μm),with the eggs being retained on the No. 60-mesh screen. Collected flea eggs are separated from debris with a fine camel’s hair brush, carefully transferred onto a folded piece of black construction paper, and the eggs are counted. The collected eggs are examined microscopically for damage. Normal eggs appear oval, full, and opalescent, whereas abnormal eggs usually appear dull, dark, and partially collapsed. Flea ova are very susceptible to environmental conditions. When working with flea eggs, the ambient temperature should be $ 21 C and $ 50% RH. Flea ova must be protected from cold shock, so all

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collection surfaces should be warmed to ambient RT prior to use. Damaged eggs are removed and replaced. The aliquot of eggs on the black construction paper is then transferred to a glass vial, Petri dish, or plastic cup for testing. Treated paper disks from each of five concentrations of test compounds are seeded on TD 0 with 100 newly collected flea ova (,18-h-old), covered with rearing medium, and incubated at 26 C
28 C and 80%
85% RH. Direct counts of larvae (ovicidal activity), pupae (larvicidal activity), and adult fleas (inhibition of adult emergence) are made on TD 7, 13, and 28, respectively. Also, these methods can be applied to test the activity of test compounds on carpet [short nap, foam backed, 5.25 diameter (0.1503 ft2)] by treating carpet swatches with serial dilutions of a test compound (mg/ft2). The swatches are seeded on TD 7, 35, and 114 with 100 newly collected flea eggs (,18-h-old). The swatches are covered with rearing medium 3 days after egg seeding and incubated at 26 C
28 C and 80%
85% RH. The duplicate or triplicate swatches per concentration are recommended. The direct counts and efficacy determination of larvae (ovicidal activity at 72 h), pupae (larvicidal activity), and adult fleas (inhibition of adult emergence at 35 days) are made on TD 3, 7, 14, and 35 postseeding of ova.

Filter paper—flea larvae A number of in vitro contact assays have been developed to determine flea larvicidal activity of the test compounds.60,79,80 The flea larvae are collected by sieving flea eggs collected from in vivo or in vitro rearing methods. The treated paper disks from each of five concentrations of test compounds are seeded on TD 0 with 30 first instars81 or 7-day-old second instars as they were found to be optimal because they had the most uniform size and at this age exhibited a greater need to feed.80 The larvae are covered with rearing medium and incubated at 26 C
28 C and 80%
85% RH. The direct counts of pupae (larvicidal activity) and adult fleas (inhibition of adult emergence) are made on TD 13 and 28, respectively. The dilutions of test compounds in acetone (100 μL) are individually added to a glass vial (1.5 cm 3 3.0 cm) and mixed with 20 mg of sand and 10 mg of flea feces. The vials of treated media are ball milled for 1.5 h to remove the acetone. First instars (B30) or 7-day-old second instars are added to each vial. The vials are sealed with dental roll plugs, placed in a plastic bag (to maintain 75% RH) and incubated at 28 C. Reading of the assay is first done on TD 0 and then on 3 consecutive days at about the same time each day. The numbers of live (exhibiting motility) and dead (often difficult to identify because they become desiccated) larvae are recorded. The number of dead larvae is calculated by subtracting the live larvae from the total count on TD 0.

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A validated larval development inhibition bioassay was developed to determine the baseline susceptibility of field-collected strains of cat fleas to imidacloprid.82,83 The bioassay involved the freshly hatched flea larvae exposed to a standard flea larval rearing medium impregnated with imidacloprid at a series of incorporation rates. The proportion of larvae that failed to develop through to the adult stage is recorded and compared to larvae reared in untreated control rearing medium. A dose-mortality response is calculated for each incorporation rate by probit analysis. A unique feature of this bioassay is the manner of exposing only hatched larvae, which was overcome by the use of a streak of nontoxic acid-free glue placed on the underside of a Petri dish lid. The lid is then placed over a dish containing rearing medium. When the flea eggs adhered to the underside of the lid hatch, the larvae fall onto the rearing medium, and the number of hatched eggs can be counted under a dissecting microscope. Advantages of this bioassay are (1) smaller numbers of adult fleas are required, because only flea eggs are collected for the test; (2) IGRs and other novel insecticides can be evaluated; and (3) using a discriminating dose for the detection of reduced susceptibility in field strains can be determined with as few as 40 eggs.

Glass vial contact bioassay Contact—adult flea Rather than treat filter paper disks with the investigational compound, the inner surface of a rounded glass flat-bottomed vial (7-dram screw cap) can be used. The test compounds dissolved in acetone are serially diluted and pipetted (50 μL) into the flat-bottomed glass vials, placed on a roller or hand rolled to evenly distribute the compound on the inner surface, and allowed to dry 18
24 h at RT in a fume hood. The vials of acetone only (50 μL) are used as controls. The replicates of each dilution are tested. Total of 20 fleas (newly emerged adults aged 1
3 days) are aspirated into numbered test tubes and randomly allocated to the treatments. The fleas are then transferred from their numbered test tube to their assigned treatment glass vial, and a plastic drilled-out vial top with an inner lining of nylon mesh sufficient to provide ventilation is then secured. Treated and untreated vials with fleas are held at RT on a lab countertop. The number of fleas dead or unable to make forward motion at 24 and 48 h after introduction into the glass vials is recorded. The probit analysis of flea mortality for each test compound is used to estimate LC50 and LC95 values for each test compound at the 24 and 48 h time points. The contact glass vial bioassay can also be used to test compounds against flea eggs, larvae, and pupae.

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Contact—flea eggs and larvae Exposure of flea eggs and hatched larvae (first instars) to sand and flea feces treated with a compound in a vial has been described.81,84 The dilutions of test compounds (100 μL) are added to a glass vial (1.5 3 3.0 cm) with 20 mg of sand and 10 mg of flea feces. The treated vials are ball milled for 1.5 h to remove the acetone. The flea eggs (B30) or larvae (B30 first instars) as clean as possible without flea feces are added to the vials. The vials are sealed with a dental roll plug and incubated at 28 C and 75% RH. The number of emerged larvae in the egg assay or the number of dead and live larvae in the larval assay is recorded for 3 consecutive days and the percent mortality determined (quality control of 20% or less mortality of negative control treatment). Contact—pet hair The rate and the extent of migration of an adulticide or IGR can be evaluated on the surface of treated cats up to 72 h as assessed by larval hatch (ovicidal activity) of cat flea eggs from adult fleas exposed in vitro to hair clippings. The hair clippings (1.0
1.5-in.2, B0.25 g) are collected at 1, 4, 24, 48, and 72 h from between the shoulder blades, mid backline, tail heat, and chest of cats untreated and treated with a spot-on formulation. Aliquots of 50
100 cat flea eggs are sprinkled onto each hair sample in Petri dishes. The dishes are shaken to allow the eggs to fall down into the hair sample to ensure all flea eggs have contact with the hair clippings. After 30 min exposure of the eggs to the hair clippings, the eggs are removed to a clean Petri dish and incubated at 26 C
28 C and 75%
80% RH. The larval hatch is determined and recorded 72 h after incubation, and the percent hatch (ovicidal activity) is determined. Using the artificial membrane system, adult fleas are exposed to adulticides or IGRs residues on dog hair while they fed on blood through a Parafilm membrane.59

Ingestion/feeding assays Membrane feeding—watch glass or Petri dish—adult fleas Membrane feeding is carried out using a flat-bed incubator apparatus maintained at 37 C
40 C. Test solutions are added to 5 mL of citrated bovine blood in a 50-mm glass beveled-edge watch glass or 50-mm diameter Petri dish and sealed with a thin Nesco/Parafilm membrane. Control fleas are fed on blood containing 0.5% (v/v) DMSO. A small cage or plastic test tube with the plastic lid bored out and replaced with a fine mesh containing 30
35 adult fleas is inverted over the container so the fleas can feed through the membrane on the warm blood. The tubes with fleas are placed in contact with membranes to allow fleas to feed through the mesh top. The fleas are allowed to feed for 2 h. On removal from the feeding apparatus, fleas were

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maintained at 25 C and 75% RH. Efficacy recordings (a subjective visual assessment of flea viability based on motility) are made at 2 and 24 h after the initiation of feeding. The screening assays are conducted in duplicate and comparative assays in triplicate. Each compound is tested at half-log intervals with endpoint data recorded as LC80 (the lowest concentration to cause $ 80% mortality) in μg/mL.85 Membrane feeding—artificial dog feeding chamber—adult fleas The modifications to the artificial dog membrane feeding system47 have been made for evaluating parasitcides.59
61 In brief the modified apparatus is a plexiglass chamber divided into a heated upper and a nonheated lower half. A plexiglass divider separates the heated upper and nonheated lower sections and has a manifold holding a series of 25 (5 cm) or 104 (2.5 cm) cages. The bovine blood, with or without concentrations of parasiticides, is introduced into plastic sleeves with Parafilm stretched across each sleeve bottom to form a membrane. Each blood or blood-parasiticide filled sleeve is placed into the heated upper half with the Parafilm membrane interfacing with one of the 25 (5 cm) or 104 (2.5 cm) cages. A small cage with 25 adult fleas is attached to the bottom of each plastic sleeve and the fleas feed on the warm blood through the Parafilm membrane. The blood in the heated chamber is maintained at 40 C, and the fleas in the bottom half are maintained at 28 C and 85% RH. After 24 h or a suitable PT time of feeding the fleas are assessed for mortality. This assay is also applicable to the testing of IGRs.59,60 Shoop et al.86 and Meinke et al.87 utilized the artificial dog feeding system60 to screen nodulisporamide compounds for intrinsic systemic potency. The flea feeding apparatus has been used to test avermectins as well.61 96-Well feeding assays—adult fleas A mixed sex adult population of fleas is placed in a suitably formatted 96well plate allowing fleas to access and feed on treated blood via an artificial feeding system.88 The fleas are fed on the treated blood for 24 h, after which the compound effect is recorded. Insecticidal activity is determined on the basis of the number of dead fleas recovered from the feeding system. 96-Well feeding assay—flea larvae Insecticide doses are prepared in acetone stock solutions at 10 mg/mL. Since acetone is corrosive to plastic microwells, serial dilutions are made using methanol, and 100 μL of each dilution is added to each well. Each dose is replicated at least 16 times. For the contact bioassay the insecticide dilutions in the 96-well plates (surface area 0.34 cm2) are allowed to dry for B2 h in a fume hood yielding surface coatings of 10
0.087 ppm depending on the insecticide being tested. For the oral (feeding) bioassay, B0.5 mg of

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powdered porcine blood curds is added to each well containing insecticide and allowed to dry as above, thus yielding treated blood curds from 10 to 0.78 ppm. The 96-well treated plates are placed upside down on a 96-well plate containing one-third instar for each well. The two plates held close together are turned upside down to allow the larvae to fall into the treated wells. The plates are incubated at 27 C and 75% RH. The number of surviving larvae is counted after 24 h, and LD50 and LD90 values are determined.89

Repellency bioassays A repellent, by definition, is a chemical substance causing oriented movements away from the source.90 While this definition is easily applied to flying insects, it is more difficult to define for crawling insects on a mammalian host, such as fleas. In vitro repellency bioassays for fleas generally subscribe to measuring repellency by movements away from the chemical substance or an antifeeding effect. In vivo assays are less straightforward because movement away (repellency) from the treated animal would be measured by causing the infesting flea to leave quickly or fall off (expellency) soon after contact with the hair coat or skin of the treated animal. Thus interfering with the feeding process preventing the flea from taking a blood meal (antifeeding) or killing the flea before it can take a blood meal (antifeeding). The repellent activity against fleas is controversial.91 Filter paper—adult flea Two long strips of filter paper are impregnated with test compounds (dissolved in ethanol) or ethanol only (control). After drying at RT the two filter papers are glued together along the long side and inserted into a glass tube containing nonfed adult cat fleas. The distribution of cat fleas in each half of the filter paper is recorded after 30 min to calculate repellency.92 Results showed that the essential oil of Cinnamomum osmophloeum (from leaf), Taiwania cryptomerioides (from heartwood), and Plectranthus amboinicus (from leaf) exhibits repellent activity against cat fleas in a dose-dependent manner. Moreover, the repellent activities against cat fleas of 2% transcinnamaldehyde (the main constituent of C. osmophloeum essential oil) and 0.5% thymol (the main constituent of P. amboinicus essential oil) were 97.6% and 90.6% and persisted for up to 4 and 8 h, respectively. These results are comparable to those of 15% DEET. Olfactometer tubes In vitro flea repellency systems include Y- or T-tube olfactometers. The four-arm airflow olfactometer93 might be an option, but no reports have been published as yet for flea repellency.

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Antifeeding bioassays Glass or plastic watch glass—adult flea This assay is essentially the same as the membrane feeding assay using a glass or plastic watch glass. A small cage or plastic test tube containing 30
35 adult fleas is inverted over the watch glass so that the fleas can feed on the warmblood through the treated Parafilm membrane. The fleas are allowed to feed for B6 h. The number of blood-engorged fleas is counted at B1, 3, and 6 h. The endpoint data are recorded as an ED80 in μg/mL. Synergistic activities Synergy (1650s, Greek and Latin synergia “cooperation,” working together) is defined as the interaction of two or more agents, elements, or forces where their combined effect is greater than the sum of their individual effects.94 Two examples of synergistic activities of compounds against fleas are as follows: 1. Laboratory flea egg eclosion (hatching) tests and cat/dog flea ovicidal efficacy tests were conducted in support of the patent, and a patent declaration was submitted on September 24, 1999. Based on data provided in this declaration as summarized below, the patent was granted.95 Flea egg eclosion test—filter paper—eggs This test determines the emergence of an insect larva from an egg. The inhibitory activity of fipronil and (S)-methoprene alone and in combination was evaluated on egg eclosion, larval development, and adult cat flea emergence. The dilutions were prepared in acetone at 0, 10, and 50 ppm. Fipronil and (S)-methoprene combination concentrations were prepared at 10 ppm each, 50 ppm each, 10 ppm fipronil/50 ppm (S)-methoprene, and 50 ppm fipronil/10 ppm (S)-methoprene. The treated filter paper disks of four replicates per concentration were placed into individual Petri dishes. The cat flea eggs (B26
28, ,24-h-old) were deposited onto the surface of each disk, and the dishes were incubated at B23 C
29 C and B75%
90% RH. The number of larvae that hatched at B72 h was recorded, place in rearing medium and incubated. The number of pupae and emerged adult fleas was recorded 35 days after deposition of the flea eggs onto the treated disks. The fipronil alone exhibited limited ovicidal activity (,10%) at 10 and 50 ppm but provided a significant larvicidal effect at both doses. The (S)-methoprene alone exhibited ovicidal efficacies of 28.4% at 10 ppm and 88% at 50 ppm. The (S)-methoprene was lethal against the developing flea larvae yielding 100% inhibition of adult flea development at 10 and 50 ppm. The fipronil/ (S)-methoprene combination at 10 ppm yielded an ovicidal efficacy of 45.5% that was 1.6 times greater than (S)-methoprene alone, thus indicating a synergistic effect in combination with fipronil. Likewise, the combination of

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fipronil/(S)-methoprene at 50 ppm each yielded an ovicidal efficacy of 97.3% or 1.1 times greater than (S)-methoprene alone. At 10 ppm fipronil/ 50 ppm (S)-methoprene the ovicidal efficacy was 88.7%, which was comparable to the ovicidal efficacy of 88.9% of (S)-methoprene alone at 50 ppm. In contrast, 50 ppm fipronil/10 ppm (S)-methoprene exhibited an ovicidal efficacy of 92.9% or 3.3 times greater than (S)-methoprene alone at 10 ppm. All combinations of fipronil and (S)-methoprene were lethal to developing flea larvae, resulting in 100% inhibition of adult flea emergence.96,97 These in vitro data were subjected to the methodology of calculating synergistic responses of Colby.98 Although this methodology is directed to determine the synergistic efficacy of herbicides, the same concept can be used to determine if a combination of insecticides, parasiticides, pesticides, etc. exhibits synergistic efficacy. The results showed that 10 ppm fipronil 1 10 ppm methoprene yielded an expected efficacy of combination of 26.9%; observed efficacy 5 41.9%; 50 ppm fipronil 1 50 ppm methoprene: expected efficacy 5 86.7%; observed efficacy 5 93.1%; 10 ppm fipronil 1 50 ppm methoprene: expected efficacy 5 86.0%; observed efficacy 5 84.7%; 50 ppm fipronil 1 10 ppm methoprene: expected efficacy 5 30.7%; observed efficacy 5 88.9%. These results were consistent with data reported in the patent declaration, that is, at the 10:10 and 50:10 combinations, there is an enhanced efficacy that suggests synergism at lower concentrations of (S)-methoprene. Thus the calculation using the Colby equation confirmed that the combination was synergistic. In vivo dose-confirmation studies In vivo studies using fipronil/(S)-methoprene combination spot-on products have been reported (Frontline Plus) for cats99 and dogs.96,100,101 Both cat and dog in vivo studies demonstrated synergistic ovicidal activity of fipronil in combination with (S)-methoprene. In the dog study the percent reduction in larval hatch of the combination was enhanced (B1.5 times) over (S)-methoprene alone from TD 57
85. Likewise, the percent reduction of adult development of fipronil/(S)-methoprene treatment was enhanced by B1.3 times over (S)-methoprene alone. These in vivo dog efficacy data were later subjected to the synergistic calculations of Colby.98 The comparison of the expected efficacy using the Colby equation98 and the actual observed efficacy demonstrated that starting at TD 57, the combination of fipronil and (S)-methoprene exhibited a synergistic efficacy that was surprising and unexpected. This is a unique example of demonstrating and confirming in vitro results with in vivo data on the host species. Another example has been demonstrated with dinotefuran and fipronil.102 2. The IGRs (S)-methoprene and pyriproxyfen are widely used as topical treatments of pets or applied to the indoor environment to control cat

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fleas. The toxicity of (S)-methoprene, pyriproxyfen, and the combinations of both IGRs to cat flea larvae has been determined.103 The LC50 of (S)methoprene and pyriproxyfen applied to larval rearing media was 0.39 and 0.19 ppm, respectively. The combinations of (S)-methoprene:pyriproxyfen in ratios of 1:1, 5:1, 10:1, and 20:1 produced LC50 values of 0.06, 0.09, 0.19, and 0.13 ppm, respectively. The pyriproxyfen synergized the activity of (S)-methoprene as indicated by the combination indices (CI). The ratio of (S)-methoprene:pyriproxyfen (40:1) provided an LC50 of 0.42 ppm, and the pyriproxyfen was not synergistic. The combinations of pyriproxyfen:(S)-methoprene in ratios of 5:1, 10:1, and 20:1 provided LC50s of 0.14, 0.20, 0.20 ppm, respectively, and the (S)-methoprene did not synergize the activity of pyriproxyfen. The dose-reduction indices indicated that the concentrations of IGRs in the combinations of (S)methoprene:pyriproxyfen (ratios of 20:1 or less) could be reduced by at least one-third of the amount required by (S)-methoprene alone to provide similar larval mortality. The combinations of (S)-methoprene and pyriproxyfen may be effective in increasing the residual activity on pets and assist in reducing the likelihood of insecticide resistance developing to IGRs.

In vivo method(s) Adult flea efficacy studies Dogs and cats Guidelines for flea efficacy studies of topical and systemic parasiticides on dogs and cats are provided in Marchiondo et al. (see Appendix A of Ref. [2]), as well as regional regulatory guidelines. The WAAVP guidelines on evaluating the efficacy of parasiticides against fleas cover in vitro and in vivo study designs for the assessment of efficacy against adult fleas, flea eggs, and larvae; controlled study designs; efficacy; repellency; and antifeeding (see Appendix A of Ref. [2]). Types of studies include dose determination, does confirmation, simulated home environment (SHE), speed of kill, and field trials. In addition to the guideline, detailed methodology on challenge, infestation, and evaluation is provided herein. Mice Topical The mice (CD-1) are anesthetized, and varying microliter volumes of a test solution, suspension, or emulsion of a parasiticide or placebo are applied along the topline of the mouse. Anesthetized mice are infested with 10
15 unfed cat fleas. The efficacy is evaluated at various time intervals up to 1 h postinfestation. The efficacy is expressed as a percentage reduction of the flea population for test concentrations compared to the placebo control treated mice.

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Systemic Studies have been performed to determine whether cat fleas that feed on laboratory mice that have been treated PO with test compounds in this small animal bioassay can detect systemically active. In these studies, acepromazine maleate was used to temporarily sedate various strains of mice and allow fleas a window of time to feed undisturbed. For validation of the model, CD-1 mice were dosed PO with seven known insecticides at 30, 10, and 1 mg/kg. The mice were sedated with 0.0125 mL acepromazine maleate IP and infested with fleas. After 2 h, fleas were removed, one-third were examined immediately to confirm the occurrence of feeding with 77% found to have ingested a blood meal. The remaining fleas were incubated for 24 h to determine mortality. Nitenpyram, the active ingredient in Capstar, was highly active ( . 94%) at 1 mg/kg. Selamectin, the active ingredient in Revolution, was very active (86%) at 10 mg/kg, but inactive at 1 mg/kg. Fipronil, the active ingredient of Frontline TopSpot, was very active (83%) at 30 mg/kg, moderately active (54%) at 10 mg/kg, and inactive at 1 mg/kg. Cythioate, the active ingredient in Proban, and nodulisporic acid, an oral insecticide, were moderately active (64% and 55%, respectively) at 10 mg/kg, but both were inactive at 1 mg/kg. Lufenuron and ivermectin exhibited no efficacy at any level tested. These findings suggest that this mouse model can effectively identify systemic flea-control leads and, subsequently, reduce or replace the use of dogs and cats in early efficacy studies prior to confirming results in the target animal species for which the product is being developed. A patent application on a flea feeding mouse model apparatus104 to test the systemic activity of parasiticides includes a containment system comprising a subject-restraining apparatus and a removable housing that contains the fleas in proximity with the nonsedated blood host (CD-1 mouse) in order to obtain in vivo flea feeding data. A mouse is treated PO with a parasiticide and then placed headfirst into a ventilated cylindrical 8 3 2.5 cm tube restraining apparatus so that its nose protrudes from the opening at one end of the tube. A glass tube containing fleas is then inserted perpendicularly into an opening in the restraining apparatus so that the mesh end of the flea containing glass tube is in contact with the abdomen of the mouse. The mesh side of the glass tube containing the fleas is pressed against the mouse to allow the fleas to feed undisturbed for a predetermined length of time. The tube containing the fleas is then removed and placed directly into an incubator at 28 C and 85% RH with a 12L:12D hour photoperiod cycle. The nonsedated mouse is then released. This invention allows a known quantity of fleas to feed on a test blood source undisturbed, followed by the recovery of all of the fleas for further observation and analysis, with minimal stress and manipulation of the blood source (test animal host) and no anesthesia of the animal host.

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Rats Topical Laboratory rats are dosed topically with a test parasiticide (60 mg/mL in DPGMME, dose volume 0.11 mL/kg). The topical treatments of placebo (DPGMME) and test compound are applied above the shoulders on the upper neck to avoid direct application to the flea feeding site. At 6 and 24 h, B20 unfed adult fleas, contained in a plastic 50 mL tube with a mesh top feeding membrane, are fed for 10 min on a preclipped area on the shoulder/back region. The rats are contained in a modified plastic rat restrainer during the feeding period. Following feeding, the flea tubes are stored at B20 C and B80% RH and the number of dead fleas recorded at 24 h postfeeding. In addition, fleas can be assessed for blood feeding by freezing the flea tubes and squashing the individual fleas (unpublished laboratory method—AAM). Systemic The laboratory rats are dosed PO by gavage with a parasiticide (2.5 mg/mL in PEG 200, dose volume 5 mL/kg), the same procedure as the topical evaluation described previously. Speed of kill The speed that fleas are killed, including the onset of action, after a parasiticide treatment is applied to an infested pet is one aspect of flea control that can affect pet comfort, client satisfaction, severity of FAD, and flea reproduction. A number of speed of kill studies have been reported using a variety of study designs.105
107 Basically, animals are infested with B100 fleas on TD-7 and combed on TD-5 to assess the ability of the animals to maintain infestations. Qualifying animals are randomly allocated and assigned to the desired number of treatment groups and times to be assessed PT/postinfestation (this can be specific minute-wise or hourly intervals according to the objective of the study, for example, 15
60 min, 1
48 h). The animals are weighed on TD-1 for dosage calculations and for treatments administered on TD 0. The animals are infested with B100 fleas on TD 2 (or TD 1), 7, 14, 21, and 28. After dosing on TD 0, animals in each treatment group are combed at the specified time interval. On TD 7, 14, 21, and 28, animals are combed as for TD 0. Ovicidal activity A number of study designs have been published.101,108,109 Pretreatment flea infestations are conducted on TD 12 to ensure viability of the ectoparasite on the test animals and to determine egg production and hatchability. Each animal is infested with B200 newly emerged, unfed adult fleas. Three days (72 h) following the initial infestation (TD 9), the treated animals are placed into egg collecting cages. The waste pans below the cages are cleaned, dried, and lined with a paper substrate that facilitates the collection of eggs. The animals remain in the egg collection cages overnight, and

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the eggs are collected on TD 8 (96 h PI). Flea ova (B100) are collected from below each animal and placed into separate Petri dishes. If ,100 ova are available, the entire production is placed in the Petri dish. Each Petri dish is labeled with tape that indicates the animal number, study number, and calendar date. The Petri dishes are incubated at 26 C
28 C and B80% RH. The dishes are observed (counted) for larval hatch at B72 h postcollection (TD 5). Flea growth medium is then be added to each dish. About 14 days after egg collection, pupae are sifted from the growth medium. The pupal numbers are estimated, and adult flea counts of each dish are made at 21, 28, and 35 days postcollection. The number of adult fleas is counted and recorded. All animals are combed on TD 8 following egg collection to remove the initial flea infestation population or treated with nitenpyram (minimum dose 1.0 mg/kg) at the end of the study to kill any remaining adult fleas from the test animals. Simulated home environment The clinical field performance of strategic flea treatment programs using approved products depends on three distinct factors: therapeutic efficacy, persistent (residual) activity, and impact on the reservoir of off-host developmental stages. Therapeutic and persistent activity can be measured by standard laboratory dose-confirmation studies. SHE studies are used to investigate and simulate the wider impact of strategic (e.g., preventive) animal treatments on the off-host (environmental) reservoir. To do this, animals are kept individually in pens/cages that are partly or wholly carpeted. The adult fleas are placed on the animal and eggs drop onto the carpet which provides a suitable habitat for flea larvae to hatch, develop, and pupate. Thus the flea life cycle is established and the flea population allowed to multiply freely in the pen/cage, which is the experimental unit for statistical purposes. Some pens/cages are used as untreated controls, while in/on-animal treatment interventions are applied on others. There is as yet no reliable way of measuring the number of developing fleas on or within the carpet within a pen/cage, and so PT animal flea counts are used as an indicator of the relative size of the total flea population in each pen/cage. SHE studies as described in the WAAVP guidelines (2013) (see Appendix A of Ref. [2]) can be used to simulate different clinical scenarios. 1. Prevention of a flea problem (prophylaxis)—This mimics the situation where a dog owner starts monthly treatments in anticipation of the forthcoming flea season. In this case, infestation of the dogs in a SHE study will commence after the first treatment has been applied. 2. Control of an existing flea problem—This mimics the situation where the client initiates a control program once fleas are seen on their animal (that is, during the flea season when there are already off-host life-cycle stages developing in the domestic environment). In this case the SHE study

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design will establish the flea life cycle in the pens/cages before the first treatment intervention. Note: In the EU the use of the term “control” is not widely accepted nor allowed on the indications for use. Pets are not always confined to the home and often acquire extraneous fleas which are brought back and introduced into the domestic environment. This scenario can be simulated in a SHE study by infesting flea-free animals with a small number of new adult fleas at different intervals posttreatment compared to a corresponding untreated control group. In contrast to other types of laboratory studies (in which fleas are normally destroyed after combing and counting), fleas combed off dogs in SHE trials are replaced on the animal of origin in order to minimize the effect of the procedure on flea population dynamics within the pen/cage. SHE studies have to be carefully managed as flea populations can escalate very quickly causing animal welfare concern. Thus SHE studies can provide a severe test for the product under investigation. It is often necessary to curtail the flea population by removing part of the carpet in control pens and/or by limiting the number of adult fleas placed back onto the control animals after each flea comb count. When this happens, subsequent flea count comparisons will underestimate the true efficacy of the treatment program. In published SHE studies of which there are too many to reference here, one approach is where animals in treatment groups are treated on TD 0, 30, and 60 and maintained in solid-sided cages with solid carpeted floors. Each animal is infested on TD 1, 7, and 14 with 100 unfed adult fleas. Individual flea comb counts are performed at various time intervals, for example, TD 3, 9, 16, 21, 28, 35, 42, 49, 56, 63, 70, 77, 84, 91, and 95. After each count and efficacy determination the animals are reinfested with live fleas. To augment flea challenge the carpeted area in each cage is sprinkled weekly with larval flea growth media (dried blood, yeast).110,111

Repellency Flea repellency—mice This unpublished assay, based on modified flea and tick in vitro assays, enables fleas to feed on anesthetized mice treated with potential flea repellents. The CD-1 mice (30 g) have an estimated body area of B25 cm2. For dose calculation the required compound application rate is 500 μg/cm2 with a concentration of 6.25% active ingredient yielding a dose volume of 200 μL/mouse. The mice are anesthetized with 0.1 mL of pentobarbital IP and then treated topically with the test compound. The test compound is applied in two 100 μL streaks to the dorsal and ventral midline of the mouse from the head to tail. The test compound is then spread by rubbing with a gloved finger. The anesthetized treated mouse is placed in a box (40 3 25 3 25 cm) with a fine nylon mesh lid. Total of 10 unfed immobilized adult fleas are placed onto the mouse B30 min PT and allowed to feed for

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20 min. Flea behavior observations during the flea feeding period are recorded. After the feeding period has ended, fleas are asphyxiated with CO2 and collected. The repellency of fleas is assessed by determining if a blood meal was taken. The fleas are crushed on a Whatman Benchkote pad or filter paper and recorded positive for feeding based on the presence of blood. The percentage of fed fleas is calculated for each treatment as compared to each other and the control group. The flea feeding may be inconsistent depending on the repellent activity of the test compound.

Flea repellency—rabbits Prior to testing the repellency of parasiticides on rabbits, two flea feeding sites are clipped close to the skin on a placebo rabbit and an Elizabethan collar fitted. This flea feeding test is to determine the percent flea feeding of the batch of fleas on the rabbit. The fleas (2 3 10 newly emerged unfed adults; 10 applied to each clipped area) will be contained in a nylon gauze covered chamber with a feeding area of B10 cm2 (diameter B3.5 cm) and secured to the rabbit with a “Velcro” belt and exposed for 20 min to the treated area on the rabbit. If flea feeding is $ 60%, then this batch of fleas will be used for the study. If flea feeding is ,60%, a new batch will be tested. This procedure can be repeated up to five times using the same rabbit to obtain a suitable feeding batch of fleas. Following successful flea feeding evaluation, two flea feeding sites on each test rabbit are clipped close to the skin and an Elizabethan collar fitted. The test compounds (200 μL) are diluted in IPA and applied in even streaks to each clipped flea feeding site (diameter B6 cm area, 25 cm2), using a Gilson pipette and spread by rubbing evenly with the “tipped end” of a pipette as the IPA dries to yield a site dosage of 100 μg/cm2. At intervals of 3, 4, 6, 12, and 24 h PT a challenge of 2 3 10 unfed adult fleas will be exposed for up to 20 min to the treated areas on each rabbit. This procedure will be repeated at each challenge displaying repellency $ 80%. Following each challenge, repellency (complete or partial avoidance of treated area) will be recorded, and the percent of fleas fed will be determined by chilling the fleas and then crushing them on a Benchkote pad to observe if a blood meal has been ingested in each flea. The percentage of fed fleas will be calculated for each treatment group and compared to each other and the control group. Flea knockdown, hyperactivity, mortality, and behavior effects differing from the placebo treated group should also be recorded. The unpublished rabbit model has the advantages of flea feeding of .90% on untreated animals, feeding via a nylon gauze, clipped hair with exposed treatment area, good spreading of compound on smaller treated area, no anesthesia of host required, and efficacy assessment of persist repellency.

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Flea repellency—dogs The efficacy of the topical combination of fipronil and permethrin (Frontline Tri-Act/Frontect, Merial) against the dog flea, C. canis, was evaluated on dogs.112 This study was conducted to measure the 1, 6, and 24 h efficacy, as well as the repellent activity, of the fipronil
permethrin combination on treated versus untreated dogs. Beagle dogs (12) were randomly allocated to one of two groups based on pretreatment live flea counts. The dogs in Group 1 remained untreated, whereas the dogs in Group 2 were treated once on TD 0. Each dog was infested with 100 unfed adult C. canis on TD 2, 7, 14, 21, and 28. The dogs were combed for fleas 1 and 6 h after each infestation. Following this examination, all fleas remaining on the liner at the bottom of each cage were collected and counted. All live fleas were placed back on each dog after the 1- and 6-h counts. A comb-count was performed at 24 h PI on all dogs. The treated dogs had significantly (P # .01) lower flea counts than untreated dogs at every time point. The percent efficacy was $ 99.1% at 6 and 24 h after the initial and each weekly challenge up to the month. The 1-h counts also showed good efficacy of 96.5%, 98.9%, 92.0%, 70.2%, and 55.7% on TD 2, 7, 14, 21, and 28, respectively. The repellent efficacy, assessed based on the number of fleas on the liners at 1 h, was 86.5%, 94.9%, 79.5%, 58.4%, and 43.9% on TD 2, 7, 14, 21, and 28, respectively. Antifeeding on dogs and cats Antifeeding action is defined as the ability of the agent to prevent the fleas from actually biting the host113 in order to avoid the injection of antigenic saliva at the bite site and thus prevent a rise of FBD or FAD. When dog fleas are free in the hair coat of a dog, 21.2% of dog fleas will have initiated blood feeding after B5 min, 72.5% have fed after 1 h, and 95.2% of males and 100% of females have taken a blood meal after 6 h.114 When cat fleas are free in the hair coat of a cat, 72.5% of the cat fleas have fed within 1 h after deposition with nearly all the cat fleas having taken a blood meal after 6 h.115 The dog flea (C. canis) will initiate blood feeding on dogs more slowly than the cat flea on cats, but the initial period of blood feeding was significantly longer for cat fleas on cats (11
25 min) than for dog fleas on dogs (5
6 min). Rust et al.116 demonstrated antifeeding properties of compounds in vitro as well as in vivo studies.

Pulicidae Echidnophaga gallinacea Westwood, 1875—sticktight flea Biology and life cycle The sticktight flea occurs throughout the subtropical and southern temperate climates throughout the world and is a major pest of the domestic chicken

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(Gallus gallus). This flea is said to have the greatest number of hosts, both mammalian and avian, of any species of flea.117 It infests poultry (domestic fowl and wild birds comprise the most frequent hosts) and various mammals (dogs, cats, rats, horses, foxes, pigs, skunks, rabbits, squirrels, and deer). The adult female sticktight fleas measure B2 mm in length, while the males are ,1 mm in length. The head is sharply angled on the frons with no genal or pronotal ctenidia. There are two setae behind the antenna, and females have a well-developed occipital lobe. The large mouthparts conspicuously project from the head. The biology of this flea varies from that of the other pulicid fleas, in that the female fleas tend to aggregate on the head, particularly near and around the eyes, or other bare spots (comb or wattles) once on a host. The female flea firmly attaches its mouthparts into the host skin and remains attached in an often ulcerated wart-like bleb for the duration of 2
6 weeks. Practically all oviposition observed by Parman118 took place with the fleas attached to the host. The oviposition occurs 6
10 days after the attachment. The eggs are deposited either in the sore or drop to the ground and begin developing in 4
14 days, but typically 6
8 days.118 The egg laying ceases when the female has depleted the reserve of stored sperm but may begin again if the female remates. Larvae that eclose from eggs laid in the skin lesions drop to the ground to complete development through three instars in 14
31 days. Pupal development occurs after 9
19 days, and newly emerged adults locate a host and attach within 5
8 days. The entire life cycle from egg to adult may be completed in 30
65 days.119,120

Rearing method(s) Glass shell vials and mason jars Fine dust with flea feces is collected overnight from heavy paper placed on the floor of a chicken coop holding chickens that are heavily infested with adult fleas of Echidnophaga gallinacea.118 The collected material is sieved and examined for flea eggs. A sample of material with eggs (2 cm diameter of paper sample) is placed into a 5 g shell vial that is closed using a piece of sponge dipped in water, squeezed nearly dry, and stuffed loosely into the mouth of the tube. Pint Mason jars (5 in. diameter and which hold about 0.5 quarts) and tumblers holding about a 0.5 pint also can be used and filled about half full using screened material collected from under infested chickens. A moistened sponge is pinned to a muslin cover and suspended in the jar. The container is covered with a muslin cover. All vials and jars are incubated at an average of 24.4 C; no RH measurements were given. The minimum period from oviposition to emergence of adult fleas was 30 days with a maximum period of 65 days.

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96-Well plates Freshly harvested eggs of E. gallinacea collected from California ground squirrels housed in a novel nest box121 were placed individually into the wells of a 96-well polystyrene, round-bottom ELISA plate (Cell Wells, disposable nonsterile assay plates [25855]) with a fine camel’s hair paintbrush. ELISA plates loaded with eggs are then placed inside desiccators (15.5 C, 21.1 C, and 26.7 C and 35%, 45%, 55%, 65%, 75%, and 85% RH using saturated salts solutions) and checked every 24 h for hatching. After hatching, each well is filled B2/3 with larval rearing media. The plates are checked every 24 h to record cocoon formation and covered with clear tape during the cocoon stage to prevent emerging adult fleas from escaping.122 Average egg hatch was substantially reduced at 31.5% RH, and no larvae survived more than 48 h at ,55% RH.122 On host—rat E. gallinacea has been reared in the laboratory on white laboratory rats.117 The method of rearing the flea was similar to that used by Douglas and Wheeler.123 E. gallinacea was found naturally on the common or Norwegian rat, Rattus norvegicus, in Sutro Forest near the Hopper Foundation in San Francisco. Fleas and their host were kept in 5 gal earthenware crocks with a glazed surface. The top edges of the crocks were fitted with metal bands with flanged ends, which were tightened by a bolt and nut. These bands extended slightly above the top of the crock and had holes bored on each side so that a metal rod could be slipped over the top of a heavy piece of metal screen, cut to fit the top of the crock, and set inside the band. About 2 in. of wood shavings were placed in the bottom of the crock; 1
2 teaspoons of dry, powdered horse or sheep blood is then added, along with the same quantity of dried, ground Brewer’s yeast, as recommended by Sikes.124 The shavings were slightly moistened, and the host and fleas were placed in the crock and kept in a room at 21.1 C
23.9 C and 80%
90% RH. The length of time that the rat could be maintained in each crock depended on the amount of urine it produced. When the shavings become too soggy with urine, the immature stages will not develop. The laboratory rats produce urine in considerable quantities. Confined host animals were not given drinking water but were supplied with fresh carrots, sunflower seeds, and rolled oats three times a week. The laboratory rats received in addition, dog chow. In order to maintain pure cultures of fleas, animals were brought in from the field and placed in crocks with their own ectoparasite fauna intact, at the start of a culture. Ectoparasites were collected from other rats of the same species which inhabited the area were sometimes placed in the crock with the host animal. After several weeks had passed and fleas had begun to breed in numbers, a large sample of fleas was taken out of the crock, mounted, and

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identified. Examination of living fleas involved the use of a large coverslip fastened near one end of a glass slide with a piece of cellulose tape, leaving the other end of the coverslip free. A flea can readily be placed under a coverslip with the aid of a hand bulb aspirator, and the coverslip easily held in place with the index finger while the insect is being examined. After living fleas were identified and separated to species, pure cultures were maintained. On host—ground squirrel California ground squirrels (Spermophilus beecheyi) naturally infested with E. gallinacea were live trapped from the wild and housed in a novel nest box.121,125 The squirrels were housed individually in clear polycarbonate guinea pig cages (51 3 41 3 20 cm tall) with stainless steel wire lids. Each cage was provided with a two-piece nest box constructed from 0.81 mm sheet aluminum. The upper section (16.5 3 16.5 3 18 cm tall) was available to the squirrel via a single round (7.6 cm diameter) opening in one wall and was built with a hinged lid allowing access to the animal, if necessary. The floor was made of 8-mesh (2.54 mm opening) stainless steel milling grade wire cloth riveted in place about 2 cm from the bottom. The main upper section was designed to fit inside a removable base pan (17.5 3 17.5 3 8.3 cm tall), where flea eggs accumulated as they dropped off the animal through the wire cloth. The ground corn cobs (0.25-mesh, 6.35 mm diameter) were used as bedding. The wire lids covering the individual cages were filled with laboratory rodent chow and supplied with a water bottle. Newly acquired wild caught ground squirrels were fed a variety of seeds and nuts, and mixed greens and carrots (for moisture) for the first few weeks allowing them time to learn how to feed and drink from the wire lids. The caged animals were maintained at 21.1 C 6 1 C, 30%
70% RH, 12 L:12D hour photoperiod, and room air exchange of 15 times/h. The flea eggs and debris were collected from the infested ground squirrels daily from the nest box base pans and mixed with B70 cm3 of larval rearing medium [1 part nutritive medium consisting of 0.15 spray-dried beef blood, 0.75 ground dog chow, 0.10 inactive dry yeast (Nutrex 55) by weight to 3 parts of 30mesh silica sand by volume] and incubated at 26.7 C and 75% RH. Emerged adult fleas were anesthetized with CO2. Total of 20 pairs of E. gallinacea were placed on each of five ground squirrels previously established in the laboratory to provide pure colonies of the flea. After establishment of the flea colony, periodic additions of 20 females and 20 males were applied to each animal every 2
3 weeks to maintain adequate egg production. On host—rabbit The life history of Echidnophaga myrmecobii has been described with details of the technique and apparatus used in artificial cultivation of this species in

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the laboratory.126 It was found that the natural food of the larvae for this flea species was feces of adult fleas. Dried ox or rabbit blood when mixed in sand provided a suitable artificial medium for the cultivation of these larvae. The optimal temperature for breeding the immature stages was 22 C. The oviposition was stimulated during cold weather by raising the RT to 22 C. Special cages were devised whereby eggs and feces from this flea species, uncontaminated by the excreta of their host, could be collected. A trap was constructed in which adult fleas could be conveniently collected as they hatch. This is described and illustrated in this publication.126 The methods used to separate flea eggs from foreign material, and collection of cocoons from the medium, are also described. The rabbits and rats have been parasitized by artificial methods. A flea-proof cage for housing laboratory rabbits has been devised and is described.126

In vitro method(s) No specific in vitro tests to assess the efficacy of parasiticides against Echidnophaga spp. were found in the literature. However, based on the rearing information of Echidnophaga spp., in vitro assays of parasiticides against eggs, larvae, and newly emerged adult fleas could be developed on glass-treated surfaces and treated larval rearing medium. In vivo method(s) E. gallinacea cannot be fed periodically on animals because the adult flea is a permanent parasite. Anesthetized mice and rats can be treated topically or systemically and then exposed to 25
30 unfed adult E. gallinacea. After being experimentally infested the fleas make their way to the head region of the test animal and attach. The fleas can be mechanically removed after a specified exposure period and mortality determined. Flea and tick products as well as various insecticides are recommended for treatment of the E. gallinacean on naturally infested fowl species, but no published studies have been found. Topical and systemic rodent assays could be viable options for in vivo evaluations of parasiticides against Echidnophaga spp.

Hectopsyllidae Tunga penetrans Linnaeus, 1758—sand or chigoe flea Biology and life cycle The sand flea, jigger or chigoe, is the smallest flea known and an important human parasite in neotropical and subtropical Americas and Africa.127,128 The chigoe has an angular head with an acute frontal angle and no ctenidia. The short thorax is reddish-brown, and there are no spiniform bristles on the

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metathoracic coxae. The adult female fleas measure B1 mm in length, but gravid females may swell to the size of a pea (up to 7 mm). Similar to the sticktight flea, the female chigoe burrows into the skin of the host through a wound made with her mouthparts. Over a period of 40 h, she inserts her head and body into the wound with only the last two abdominal segments being exposed. An ulcerous sore forms around the embedded female flea leaving only the last abdominal segments exposed. The adult male fleas are free living, measure B0.5 mm long, mate with the embedded female, and never embed in the host. The embedded female flea passes up to 200 eggs through a small opening to the outside of the swelling about 6 days after initial penetration and hypertrophy.129 The eggs are ovoid, measuring about 600 3 300 μm, and drop to the ground. The eggs hatch in 1
6 days (mean 3
4 days). The larval (4
10 days), pupal (5
7 days), and adult (9
15 days) stages develop with the entire life cycle occurring in about 17 days.129

Rearing method(s) On host—rat The life cycle of Tunga penetrans can be established and maintained on Wistar rats exposed in tungiasis endemic areas.129
131 Basically, Wistar rats are placed in cages at locations for 5
7 days where tungiasis is common in humans. The animals are examined daily for embedded sand fleas. The infested rats (range of 4
7 embedded sand fleas/rat) are taken to the laboratory. The eggs are collected from the infested rats on black paperboard for 3
5 h. The eggs are kept in Petri dishes with sand from the area where the rats were infested. It was found that the most effective method for the development of flea eggs was to breed them on permanent wet paperboard (Petri dishes filled with wet cotton wool and paperboard). The Petri dishes are incubated at 25 C
28 C and 50%
71% RH. When larvae hatched, they are mixed with rearing medium and incubated. The cocoons were separated from sand and adult fleas subsequently emerge. Newly emerged T. penetrans fleas can then be placed on uninfested rats where the females will embed and the males copulate and inseminate the females with egg laying soon to follow. In vitro method(s) Contact—spray—eggs Batches of pooled eggs are randomly assigned to treatment and control groups placed into Petri dishes. The eggs are sprayed with 175 μL of insecticide applied from a distance of 5 cm using a standardized hand pump spray bottle. The application rate is equal to 0.022 L/m2. Tap water is used as a control. The Petri dishes are incubated at 25 C
28 C and 50%
71% RH. After 3, 5, and 7 days PT the eggs are examined for hatching, and larvae are

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counted.132 Corrected hatch rates are calculated as: (crude hatch rate in test group)/(crude hatch rate in untreated control group). The efficacy of a product was defined as: (1—corrected hatch rate). Negative values (hatch rate in insecticide treated group is higher than in control group) were considered as 0% efficacy and 95% CI for efficacy were calculated according to an asymptotic formula.133 The calculated values below 0% or above 100% CI were set to 0% or 100%, respectively. Relative frequencies between groups were compared applying Fisher’s exact test to evaluate statistical significance.

In vivo method(s) Tungiasis (sand flea disease) is a neglected tropical disease associated with debilitating acute and chronic morbidity. Infestations are typically treated with manual removal of the flea. A proof-of-principle study was conducted to compare the topical application of a mixture of two dimeticones of low viscosity to the topical application of a 0.05% solution of KMnO4 in 47 school children in an endemic area in rural Kenya.134 The efficacy of the treatment was assessed during a follow-up period of 7 days using viability signs of the embedded parasites, alterations in the natural development of lesion morphology, and the degree of local inflammation as outcome measures. In the dimeticone group, 78% (95% CI 67%
86%) of the parasites lost all signs of viability as compared to 39% (95% CI 28%
52%) in the KMnO4 group (P , .001). In the dimeticone group, 90% (95% CI 80%
95%) of the penetrated sand fleas showed an abnormal development after 5 days, compared to 53% (95% CI 40%
66%; P , .001) in the KMnO4 group. Seven days PT, signs of local skin inflammation had significantly decreased in the dimeticone group (P , .001). This study identified the topical application of dimeticones of low viscosity as an effective means to kill embedded sand fleas.

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34. 73. Bossard RL, Dryden MW, Broce AB. Insecticide susceptibilities of cat fleas (Siphonaptera: Pulicidae) from several regions of the United Sates. J Med Entomol 2002;39(5):742
6. 74. Hunter JS, Keister DM, Jeannin P. The effect of fipronil treated dog hair on the survival of the immature stages of the cat flea, Ctenocephalides felis. In: Hinkle NC, editor. Proc fourth Int Symp ectoparasites pets. Riverside, CA; 1997. p. 83
4. 75. Correia TR, Cruz VP, Riberio FA, Melo RM, Fernandes JI, Veocal CG, et al. [Residual activity in vitro of treated dog’s hair coat with dinotefuran on larvae and adults of Ctenocephalides felis felis (Bouche´, 1835) (Siphonaptera: Pulicidae)] [in Portuguese]. Rev Bras Parasit Parasitol Vet 2008;17(4):195
9. 76. Olsen A. Focusing on the substrates when evaluating the efficacy of flea products. In: Knapp FW, editor. Proc Int Symp ectoparasites pets. Lexington, KY; 1993. p. 70
5. 77. Bossard B, Dryden MW, Broce AB. Evaluation of bioassays for insecticide resistance in cat fleas. In: Meola RW, editor. Proc third Int Symp ectoparasites pets. College Station, TX; 1995. p. 123. 78. Marchiondo AA, Riner JL, Sonenshine DE, Rowe KF, Slusser JH. Ovicidal and larvicidal modes of action of fenoxycarb against the cat flea (Siphonaptera: Pulicidae). J Med Entomol 1990;27:913
21. 79. Moser BA, Koehler PG, Patterson RS. Effect of methoprene and diflubenzuron on larval development of the cat flea (Siphonaptera: Pulicidae). J Econ Entomol 1992;85(1):112
16. 80. Zakson-Aiken M, Gregory LM, Shoop WL. Development of an assay for the screening of compounds against larvae of the cat flea (Siphonaptera: Pulicidae). J Med Entomol 2000;37(4):571
4. 81. Bettina A, Koehler PG, Patterson RS. Effect of methoprene and diflubenzuron on larval development of the cat flea (Siphonaptera: Pulicidae). J Econ Entomol 1992;85(1):112
16. 82. Rust MK, Waggoner M, Hinkle NC, Mencke N, Hansen O, Vaughn M, et al. Development of a larval bioassay for susceptibility of cat fleas (Siphonaptera: Pulicidae) to imidacloprid. J Med Entomol 2002;39(4):671
4. 83. Kopp S, Blagburn B, Coleman G, Davis W, Denholm I, Field C, et al. Monitoring field susceptibility to imidacloprid in the cat flea: a world-first initiative twelve years on. Parasitol Res 2013;112(Suppl. 1):47
56 Available from: https://doi.org/10.1007/s00436013-3280-z. 84. McTier TL, Evans NA, Martin-Short M, Gration K. Comparison of the activity of selamectin, fipronil, and imidacloprid against flea larvae (Ctenocephalides felis felis) in vitro. Vet Parasitol 2003;116(1):45
50. 85. McTier TL, Chubb N, Curtis MP, Hedges L, Inskeep GA, Knauer CS, et al. Discovery of sarolaner: a novel, orally administrated, broad-spectrum, isoxazoline ectoparasiticide for dogs. Vet Parasitol 2016;2016(222):3
11.

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86. Shoop WL, Gregory LM, Zakson-Aiken M, Michael BF, Haine HW, Ondeyka JG, et al. Systemic efficacy of nodulisporic acid against fleas on dogs. J Parasitol 2001;87 (2):419
23. 87. Meinke PT, Colletti SL, Fisher MH, Wyvratt MJ, Shih TL, Ayer MB, et al. Discovery of the development candidate N-tert-butyl nodulisporamide: a safe and efficacious once monthly oral agent for the control of fleas and ticks on companion animals. J Med Chem 2009;52:3505
15. 88. Nanchen S, Gauvry N, Goebel T, Isoxazoline derivatives as pesticides. US Patent Application 20140120147 A1. 2014. 89. Chen Y-J, Huang C-G, Hsu J-C, Wu W-J. Development of a larval bioassay method using 96-wellmicrotiter plates for evaluation of susceptibility of cat fleas (Siphonaptera: Pulicidae) to insecticides. J Med Entomol 2017;2(1):377
81. 90. Dethier VG, Browne LB, Smith CN. The designation on chemicals in terms of the response they elicit from insects. J Econ Entomol 1960;53:134
6. 91. Halos L, Baneth G, Beugnet F, Bowman AS, Chomel B, Farkas R, et al. Defining the concept of ‘tick repellency’ in veterinary medicine. Parasitology 2012;139(4):419
23. 92. Su LC, Huang CG, Chang ST, Yang SH, Hsu SH, Wu WJ, et al. An improved bioassay facilitates the screening of repellents against cat fleas, Ctenocephalides felis (Siphonaptera: Pulicidae). Pest Manage Sci 2014;70(2):264
70. 93. Vet LEM, Van Lanteren JCV, Meymans M, Meelis E. An airflow olfactometer for measuring olfactory responses. Phys Entomol 1983;8:97
106. 94. The American Heritages. Stedman’s medical dictionary. Houghton Mifflin Company; 2002. 95. Jeannin P. Insecticidal combination to control mammal fleas. In: Particular fleas on cats and dogs, US Patent 454313-2339; 1997. 96. Marchiondo AA, Green SE, Plue RE, Wallace DH, Synergistic ovicidal activity of Fipronil and (S)-Methoprene against the cat flea, Ctenocephalides felis. In: Proc sixth int symp ectoparasites pets. Westport, CT, Mayo, Ireland. 2001. 97. Marchiondo AA. Patent declaration under 37 CFR 1.132. Synergistic activity of Fipronil and (S)-Methoprene. U. S. Patent 45313-2339. Insecticidal combination to control mammal fleas, in particular fleas on cats and dogs. Identified and demonstrated the in vitro and in vivo flea ovicidal synergy of fipronil 1 (S)-methoprene combinations and products; Issued August 1, 2000. 98. Colby SR. Calculating synergistic and antagonistic responses of herbicide combinations. Weeds 1967;15(1):20
2. 99. Marchiondo AA, Green SE, Wallace DH, Barrick RA, Jeannin PC, Efficacy of a fipronil/ (S)-methoprene combination spot-on product against the cat flea, Ctenocephalides felis, on cats. In: 46th annual meeting of veterinary parasitologists, American Association, Boston, MA; 2011. 100. Marchiondo AA, Young DR, Jeannin PC. 2001. Efficacy of a fipronil/(S)-methoprene combination spot-on product against the cat flea, Ctenocephalides felis, on dogs. In: 46th annual meeting of veterinary parasitologists. American Association, Boston, MA; 2011. 101. Young DR, Jeannin PC, Boeckh A. Efficacy of fipronil/(S)-methoprene combination for dogs against shed eggs, emerging and existing adult cat fleas (Ctenocephalides felis, Bouche´). Vet Parasitol 2004;125:397
407. 102. Delcombel R, Karembe H, Nare B, Burton A, Liebenberg J, Fourie J, et al. Synergy between dinotefuran and fipronil against the cat flea (Ctenocephalides felis): improved

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103. 104. 105. 106.

107.

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113. 114.

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onset of action and residual speed of kill in adult cats. Parasit Vectors 2017;10(1):341 Available from: https://doi.org/10.1186/s13071-017-2272-8. Rust MK, Lance W, Hemsarth H. Synergism of the IGRs methoprene and pyriproxyfen against larval cat fleas (Siphonaptera: Pulicidae). J Med Entomol 2016;53(3):629
33. Rasa C.G., Heimbach H.W., Shoop W.L., Zakson-Aiken M. Flea feeding apparatus. US Patent 2004/0069235 A1; 2004. Cruthers L, Slone RL, Guerrero A, Robertson-Plouch C. Evaluation of the speed of kill of fleas and ticks with Frontline Top Spot in dogs. Vet Ther 2001;1(2):170
4. Schenker R, Tinembart O, Humbert-Droz E, Cavaliero T, Yerly B. Comparative speed of kill between nitenpyram, fipronil, imidacloprid, selamectin and cythioate against adult Ctenocephalides felis (Bouche´) on cats and dogs. Vet Parasitol 2003;112(3):249
54. Dryden MW, Smith V, Payne PA, McTier TL. Comparative speed of kill of selamectin, imidacloprid, and fipronil-(S)-methoprene spot-on formulations against fleas on cats. Vet Ther 2004;6(3):228
36. Olsen A. Ovicidal effect on the cat flea, Ctenocephalides felis (Bouche), of treating fur cats and dogs with methoprene. Int Pest Ctrl 1985;27:10
13. Dryden MW, Payne PA, Vicki S, Debra RL, Lynn A. Evaluation of the ovicidal activity of lufenuron and spinosad on fleas’ eggs from treated dogs. Intern J Appl Res Vet Med 2012;10(3):198
204. Jacobs DE, Hutchinson MJ, Ryan WG. Control of flea populations in a simulated home environment model using lufenuron, imidacloprid or fipronil. Med Vet Entomol 2011;15 (1):73
7. Snyder DE, Meyer KA, Wiseman S, Trout CM, Young DR. Speed of kill and efficacy of flavored spinosad tablets administered orally to cats in a simulated home environment for the treatment and prevention of cat flea (Ctenocephalides felis) infestations. Vet Parasitol 2013;196(3-4):492
6. Beugnet F, Soll M, Bouhsira E, Franc M. Sustained speed of kill and repellency of a novel combination of fipronil and permethrin against Ctenocephalides canis flea infestations in dogs. Parasit Vectors 2015;8:52 Available from: https://doi.org/10.1186/s13071015-0680-1. Franc M, Cadiergues MC. Antifeeding effect of several insecticidal formulations against fleas on cats. Parasite 1998;5:83
6. Cadiergues MC, Santamarta D, Mallet X, Franc M. First blood meal of Ctenocephalides canis (Siphonaptera: Pulicidae) on dogs: time to initiation of feeding and duration. J Parasitol 2001;87(1):214
15. Cadiergues MC, Hourcq P, Cantaloube B, Franc M. First blood meal of Ctenocephalides felis felis (Siphonaptera: Pulicidae) on cats: Time to initiation and duration of feeding. J Med Entomol 2000;37:34
636. Rust MK, Hinkle NC, Waggoner M, Mencke N, Hansen O, Vaughn M. The influence of imidacloprid on adult cat flea feeding. Suppl Comp Cont Educ Vet Pract 2001;2 (4A):18
21. Burroughs AL. Sylvatic plague studies. The vector efficiency of nine none species of fleas compared with Xenopsylla cheopis. J Hyg 1947;43:371
96. Parman DC. Biological notes on the hen flea, Echidnophaga gallinacean. J Agric Res 1923;23(19):1007
109. Wall R, Shearer D. Veterinary ectoparasites: biology, pathology & control. Chapter 6 Fleas (Siphonaptera). 2nd ed. Oxford, UK: Blackwell Science Ltd.; 2001. p. 143
61.

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120. Knapp FW, Scheibner RA. Fleas affecting livestock and pets. Chapter 10. In: Williams RE, Hall RD, Broce AB, Scholl PJ, editors. Livestock entomology. NY: John Wiley & Sons; 1985. p. 183
9. 121. Metzger ME, Rust MK. Abiotic factors affecting the development of fleas (Siphonaptera) of California ground squirrels (Rodentia: Sciuridae) in southern California, USA. In: Robinson WH, Rettich F, Rambo GW, editors. Proc third int conf urban pests; 1999. p. 235
9. 122. Metzger ME, Rust MK. Effect of temperature on cat flea (Siphonaptera: Pulicidae) development and overwintering. J Med Entomol 1996;34:173
8. 123. Douglas JE, Wheeler CM. Sylvatic plague studies. II. The fate of Pasteurella pestis in the flea. J Infect Dis 1943;72:18
30. 124. Sikes EK. Notes on breeding fleas, with reference to humidity and feeding. Parasitology 1931;23:243
9. 125. Metzger ME, Rust MK. Laboratory techniques for rearing the fleas (Siphonaptera: Ceratophyllidae and Pulicidae) of California ground squirrels (Rodentia: Sciuridae) using a novel nest box. J Med Entomol 2001;38(3):465
70. 126. Mules MW. Notes on the life history and artificial breeding of Echidnophaga myrmecobii Rothschild. Aus J Exp Bio Med Sci 1940;18(3):385
90. 127. Geigy R, Herbig A. Die Hypertrophie der Organe beim Weibchen von Tunga penetrans. Acta Trop 1949;6:246
62. 128. Connor DH. Diseases caused by arthropods: tungiasis. In: Binford CH, Connor DH, editors. Pathology of tropical and extraordinary diseases. Washington, DC: Armed Forces Institute of Pathology; 1976. p. 610
14. 129. Nagy N, Abari E, D’Haese, Calheiros C, Heukelbach J, Mencke N, et al. Investigations on the life cycle and morphology of Tunga penetrans in Brazil. Parasitol Res 2007;101: S233
42. 130. Witt LH, Linardi PM, Meckes O, Schwalfenberg S, Ribeiro RA, Fiedmeier H, et al. Blood-feeding of Tunga penetrans males. Med Vet Entomol 2004;18:439. 131. Calheiors CML. Aspectos biolo´gicos e ecolo´gicos de Tunga penetrans (L., 1758) ˇ (Siphonaptera: Tungidae) em a´reas endemicas brasileiras. Ciencas: Instituto Ciˇencas Biolo´gicas, Universidade Federal de Minas, Gerais; 2007. 132. Kiesewetter T, Ariza L, Martins MM, Limongi JE, da Silva JJ, Mendes J, et al. In vitro efficacy of four insecticides against eggs of Tunga penetrans (Siphonaptera). Open Derm J 2013;7:15
18. 133. Rosenheim JA, Hoy MA. Confidence intervals for the Abbott’s formula correction of bioassay data from control response. J Econ Entomol 1989;82:331
5. 134. Thielecke M, Nordin P, Ngomi N, Feldmeier H. Treatment of tungiasis with dimeticone: a proof-of-principle study in rural Kenya. PLoS Negl Trop Dis 2014;8(7):e3058 Available from: https://doi.org/10.1371/journal.pntd.0003058. eCollection 2014. 135. Santora KA, Zakson-Aiken M, Rasa C, Shoop W. Development of a mouse model to determine the systemic activity of potential flea-control compounds. Vet Parasitol 2002;104(3):257
64.

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Chapter 3g

Arthropoda, Insecta, Hemiptera Alan A. Marchiondo, MS, PhD1 and Richard G. Endris, PhD2 1 2

Adobe Veterinary Parasitology Consulting LLC, Santa Fe, NM, United States, Endris Consulting, Inc., Bridgewater, NJ, United States

Arthropoda Insecta Hemiptera The order Hemiptera (hemi 5 half and pteron 5 wing) was first recognized by Linnaeus in the volume 10 of Systema Naturae 1758.1 Hemipterans have modified piercing-sucking mouthparts that differentiate them as “true bugs,” in which the lower lip (labium) forms a sheath surrounding the elongated, slender mandibles and maxillae.2 The maxillae are modified into a pair of concentric tubes forming the salivary and food canals. The mandibles lie external and are coupled to the secondary jaws (maxillae) serving a cutting function for inserting the mouthparts into a food source. The maxillary and labial palpi are completely lost in the Hemiptera. The functioning of the modified piercing-sucking mouthparts requires that food be liquid or be suspended in a liquid medium. Saliva is passed through the salivary canal to the food source. The sucking action of the cibarial pump controls the ingestion of the liquid food and transfers it to the insect’s midgut. The suborder Heteroptera (hetero 5 other and pteron 5 wing) possess forewings (hemelytra) partly thick and protective and partly membranous giving the appearance of a half wing. Heteropterans are among the largest groups of insects (around 75,000 species worldwide) with gradual or incomplete (egg, adult-like nymphs, no pupal stage, and winged or wingless adult) metamorphosis (hemimetabolous insects). Heteropteran groups have thoracic scent glands used for defense. Contrary to earlier diagnoses, the presence of scent glands in the adult metathorax does not form a diagnosis for Heteroptera. There are seven heteropteran infraorders.2 For the purpose of this book, the infraorder Cimicomorpha is of veterinary importance because it contains the families Cimicidae Latreille 1802 (bed bugs) and Reduviidae Latreille 1807 (assassin bugs and kissing- or cone-nose bugs). These two ectoparasite hemipteran families are hematophagic, feeding on the blood of animals and humans, and have been used in various contact and systemic screening tests to determine the efficacy of experimental compounds.

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Cimex lectularius Linnaeus, 1758—bed bug Biology and life cycle Bed bugs are parasitic insects (ectoparasites) of the cimicid family that feed exclusively on blood. Cimex lectularius, the common bed bug, is the most widespread and best known species as it prefers to feed on human blood. Two other bed bug species reportedly feed on humans: Cimex hemipterus, the New and Old World tropical pest, and Leptocimex boueti, which feeds on bats and humans and is restricted to West Africa.3 There are 9 genera of bed bugs that are ectoparasites of birds and 12 genera that parasitize bats. Only Cimex contains species that feed on bats, for example, bat bugs, and birds.4 C. lectularius may have originated in the Middle East in caves inhabited by bats and humans5,6 or may have started feeding on humans when bats roosted in homes.4 Bed bugs were mentioned in ancient Greece as early as 400 B.C. and were later mentioned by Aristotle in Historia Animalium ¯ “History of Animals.” Pliny the Elder in his encyclopedia Natural History (Naturalis Historia, first published c. A.D. 77 in Rome) claimed bed bugs had medicinal value in treating ailments, such as snake bites and ear infections. Belief in the medicinal use of bed bugs persisted until at least the 18th century. The name bed bug derives from the preferred habitat of C. lectularius to colonize near or inside bedding or other sleeping areas (made of organic materials like hay, straw, leaves, pine needles, reeds, or ropes tied across a frame in early times) and later (about the 1870s) in beds, particularly within the mattress, box spring, headboard, or bed frame. They are attracted to their hosts primarily by CO2 and secondarily by body heat, though a variety of host semiochemicals play a role in attraction.7 Bed bugs hide in tight areas (cracks and crevices) during the day and typically feed during the night with all stages seeking out hosts during periods of minimal host activity, that is, sleeping. They usually feed on their hosts without being noticed piercing the skin of the host with its specialized piercing-sucking mouthparts, the stylet fascicle. The author of the nursery rhyme “Goodnight, Sleep Tight” is unknown. The first known usage of something like “sleep tight, don’t let the bed bugs bite” dates to 1881, though the word “bed” is left out. In the book Boscobel: The Novel, two children (Fred and Flossie) come to say good night to their parents saying “Good night, sleep tight. And don’t let the buggers bite.”8 Next, in the 1884, a little girl tells boaters that she hopes they “may sleep tight, where the bugs don’t bite.”9 This example may seem to refer more to mosquitoes since they are on a river, but the 1881 example8 was set during a November winter storm. Finally, in 1897, the exact phrase appears in a usage book that explains what a boy in New England would say to a companion as

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they part for the evening—“Goodnight, sleep tight, don’t let the bedbugs bite.”10 The “sleep tight” part of the rhyme may have originated from beds supported by ropes tied in a weave like a hammock, a predecessor to the box-spring, used during the 18th and early 19th centuries. To support the mattress and prevent the bed from sagging to the floor, the ropes needed to be pulled “tight.”11 Another theory was to button “tight” your pajamas and nightgown to keep the bed bugs from getting to your skin to bite. Finally, as noted by the Oxford English Dictionary,12 “sleep tight” is defined as “to sleep soundly” or “to sleep well” (usually in the imperative). Adult bed bugs are 4
5 mm (0.16
0.20 in.) long and 1.5
3 mm (0.059
0.118 in.) wide, dorsoventrally flattened, oval-shaped, and light brown to reddish-brown. Bed bugs develop from egg to adults in about 37 days (5 weeks from egg to egg at 28 C
32 C and 75%
80% RH).13 Like other heteropterans, bed bugs develop by “gradual or incomplete metamorphosis” meaning the last larval stage develops into the adult without passing through a nonfeeding pupal stage. However, unlike most heteropterans, adult bed bugs do not have functional front wings, but only small, nonfunctional hemelytral pads.4 Bed bugs have segmented abdomens with short, goldencolored hairs that give them a banded appearance. Their long, segmented proboscis projects backward between the legs when at rest but is thrust forward when taking a blood meal. The piercing-sucking mouthparts (stylet) penetrate through the skin of the host and inject saliva with anticoagulants and painkillers that aid in a painless bite.14 Bed bugs feed about every 3
7 days and engorge with blood in about 10
15 min. Female bed bugs lay 3
5 eggs/day and can produce about 500 eggs during her lifetime dependent on the frequency of blood meals. The singly laid eggs of the bed bug are whitish, elongate, oval-shaped B1 mm long and hatch within 6
10 days depending on temperature and RH. Newly hatched bed bugs undergo five instar stages (first through fifth instar that each develops in 6
7 days) before becoming adults with each instar requiring at least one full blood meal to complete each molt to the next instar.15 Bed bugs can survive a wide range of temperatures (15 C
34 C) and RH (46%
75%) with an average of 25 C and 59% RH in laboratory colonies.16 Adult beg bugs enter semihibernation below 16.1 C and can survive for at least 5 days at 210 C but die after 15 min of exposure to 232 C.17 The thermal death point for C. lectularius is 45 C with all stages of the life cycle killed by 7 min of exposure to 46 C.17 Most life-cycle stages of bed bugs are killed by freezing with 95% mortality after 3 days at 212 C.18 Bed bugs surviving low humidity show high desiccation tolerance with earlier life-cycle stages being more susceptible to drying than later ones.19 Washing bedding at 60 C was found to be effective against all life-cycle stages, as was tumble drying on a hot cycle ( . 40 C) for at least 30 min, and dry cleaning with perchloroethylene, and freezing at 217 C for at least 2 h.

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Using data loggers it was also shown that 2.5 kg of loosely packed, dry laundry takes B8 h to reach 217 C. Soaking for 24 h in detergent-free water was found to be effective against active stages but had no effect on eggs.20 Under optimal conditions, bed bugs may live up to 1 year, although a life span of 4
6 months is more common.

Rearing method(s) Physical factors (temperature, RH, and photoperiod) and physiological factors (type and frequency of blood meal) play important roles in the laboratory rearing of bed bug colonies.16 Any change in these factors produces changes in life-cycle duration. Temperature and blood meal are the most important factors, with a marked impact on the life cycle of laboratory populations, depending on the species. A wide range of temperatures (15 C
34 C) and RH (46%
75%) with an average of 25 C and 59% were found for these colonies. Two widely used blood sources for the colonies were rabbits and humans. Typically, bed bugs are maintained in plastic containers (4.7 cm in height and 5 cm in diameter) with folded paper as harborages in an incubator at 26 6 1 C, 40 6 10% RH, and a photoperiod of 12L:12D hour and fed weekly on defibrinated rabbit blood using an artificial membrane-feeding system.

In vitro rearing methods Bed bugs can be maintained in either plastic, screw-cap containers, or glass Mason jars. Opening from the bottom and fitted with a mesh top, screw-cap containers are clear polystyrene, and measure B7 cm 3 5.5 cm (4 oz), while Mason jars are 16 oz (473 mL) wide-mouth jars.3 A Dremel rotating tool is used to remove the top of the container before gluing plankton screening to the top of the jar. This creates a screened surface for bed bugs to feed through. There are several artificial feeding methods for feeding bed bugs. The “water-bath method”21 involves specialized glassware, a circulating water bath, and a plastic membrane. The bottom of the glassware is covered with stretched Parafilm M or Nescofilm that acts as an artificial membrane. Other artificial feeding methods that were subsequently developed involve the mosquito membrane and Petri dishes feeding methods. Water-bath feeding method22 This system was originally developed by Garcia et al.,23 and later modified.21,22 The system functions by using a hot water bath coupled with a water pump to circulate heated water through a series of Nalgene tubes. The artificial feeding system uses five tubes (each B45 cm) to connect a series

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of four custom-made glass bed bug feeders measuring 60 3 60 mm supplied with caps and hose barbs. Each feeder is held in place with a chemistry clamp and stand. Feeders can be lowered or raised, brought forward or backward, or rotated left or right. Using circulated water, each container is heated to the temperature set for the water bath. System limit control and temperature control dials are both set to maintain a temperature of B40 C throughout feeding. As each container is hollow (from top to bottom) in the center, a piece of Nescofilm or Parafilm M is stretched over the bottom of the glass to contain blood that is injected into the upper opening of the feeder through the use of a pipette. Bed bug containers with mesh tops are then placed beneath and against this layer of artificial membrane. After detecting and moving to the location of the heat/blood source, bed bugs insert their mouthparts through the mesh and pierce the artificial membrane to obtain a blood meal. Bed bugs are fed defibrinated rabbit blood once/week. The feeding reservoir holds up to B4 mL of blood, but 2 mL is sufficient to eliminate air from the feeding unit. Eighty mL of blood can feed B8
10 densely populated containers, or several thousand adult bed bugs, for 2 weeks. The blood is inserted into the membrane and kept at 37 C
40 C using a warm water bath and pump that circulates the water through the feeders. Filter paper strips are placed inside the insect container to allow the bed bugs to reach the mesh lid, which is in contact with the membrane containing the blood. Lights are turned off to promote feeding. Bed bugs should be allowed to feed for B30 min. After feeding, bed bugs are housed in their original containers or vials with paper inserts. Mosquito (Hemotek) membrane
feeding system The mosquito membrane
feeding system ([email protected])24 is a water-bath feeding system that can be used to artificially feed bed bugs. The apparatus uses an electric heating element to maintain the temperature of the defibrinated rabbit blood. The blood can also be warmed by using a glass mosquito feeder that has an outer chamber for circulating 37 C water and an inner chamber into which the blood is injected and contained. Parafilm serves as an artificial membrane and is stretched over the lower portion of the inner chamber. The assembled feeder is held on top of the screened box containing bed bugs. Petri dish feeding method A small amount of defibrinated rabbit blood placed into the lid of a Petri dish, which is then covered with stretched Parafilm M plastic. The bottom of the Petri dish is used to push down the Parafilm so that it is in contact with the blood. A hotplate is used to keep the temperature of the blood at 37 C and the insect containers are placed upside down on the Parafilm so that the bed bugs can feed through the mesh lid. The Petri dish method shares some

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disadvantages with the water-bath method, such as expensive heating equipment and the leaking of blood. The advantages of this method include its quick setup and reduce some of the drawbacks from the current water-bath method, such as the possibility of flooding bed bug rearing jars with water or blood and the need for expensive custom-made glassware. Bed bugs fed during a 6 week time period showed that there was no significant difference in the numbers that fed using the Petri dish method compared with those that fed using the water-bath method.25 Development of the nymphs also showed that there was no significant difference in the time required to produce adults by either method.

In vivo feeding method(s) Animal hosts The use of live animal hosts, such as chickens, pigeons, rabbits, guinea pigs, and mice, offers another reliable in vivo feeding method. The skin of the animal is plucked or shaven of all feathers and hair.26 The host animals are then anesthetized. Feeding methods whereby bed bugs are placed onto the exposed skin of the confined host include a set of two rearing vials positioned on the middorsal sides on the host and strapped around the ventral area or feeding bed bugs contained in a jar from below.27,28 Bed bugs kept in screw-top plastic dishes (10 cm diameter 3 7 cm high) on half pieces of circular filter paper (Whatman No. 3, 9 cm) covered with 60-mesh organdie (sheerest and crispest cotton cloth made) held in place with the open-center screw top or taped in place are placed on the belly of a closely clipped rabbit with the organdie cloth in contact with the skin.29 The bed bugs then feed for 15 min from the insect container through the container membrane. Higher fecundity was reported when C. lectularius were fed on rabbits in comparison to humans, chickens, and pigeons.25 The fed insects are then maintained at B26.7 C and 50%
70% RH. Bed bug colonies feed on animal hosts or humans are housed in plastic dishes as described earlier and maintained at 28 6 2 C.29 Oviposition occurs on the filter paper. To obtain bed bugs of known age or development stage, filter papers with eggs attached are transferred to other dishes at known intervals and the stage is fed on a host. Rabbit Bed bug colonies are maintained in a temperature-controlled room at 28 6 2 C.29 The bugs are kept in a screw-top plastic dishes (10 cm diameter 3 7 cm high) on half pieces of circular filter paper (Whatman No. 3, 9 cm). The dishes are covered with 60-mesh organdie, held in place with the open-center screw top. If screw-top dishes are not available, the organdie covers are taped in place. Oviposition occurs on the filter paper. Young bed bugs are fed on a closely clipped rabbit held in a stanchion. For this process,

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the dish is simply placed on the belly of the rabbit with the organdie cloth in contact with the skin. The dish is secured by rubber bands for 15 min, which is adequate time from all bed bugs to feed. The same procedure is followed for the older nymphs and adult, with feeding once a week for maintenance, or twice a week in an active colony being used for testing. In order to obtain bed bugs of a known age or stage of development, filter papers with eggs attached can be transferred to other dishes at known intervals. The dishes are cleaned when large numbers of cast skins and heavy blood stains accumulate on the organdie covers. Mold formations can be controlled by good sanitary practices and a supply of filter papers adequate for the number of bed bugs. Dishes should not be overcrowded. Humans An effective method of in vivo feeding laboratory reared bed bugs colonies in terms of cost and reliability is the use of a human volunteer. Fifteen bed bugs obtained from a colony are placed in 4-dram vials with a screen covering a circular opening in its cap. The container vial is usually held up to the arm or leg of the volunteer, with the mesh lid touching the skin. The screen mesh is monofilament plankton netting, 60 3 60 filament count/in. which is large enough to allow bed bugs to feed, but small enough to prevent their escape.30 After feeding, insects are examined with a microscope to confirm feeding by the presence of an extended abdomen with a red appearance. While this method is the most cost-efficient and the threat of bed bug mortality from blood leakage is eliminated, it provides an uncomfortable experience for the host during and post feeding because of allergic reactions due to bed bug saliva.31 This method using human participants would be overseen by an IRB to ensure the rights and welfare of human subjects during their participation according to the rules and regulations of the Protection of Human Subjects in the Code of Federal Regulations (45 CFR 46). As an alternative, expired human blood collected from a blood transfusion facility could be used to artificially feed beg bugs.

In vitro method(s) Contact methods Residual surface tests Regulatory agencies32 have provided laboratory testing methods for the registration of products for control of adult and immature stages of bed bugs. These dry residue tests involve application of test material to surfaces, such as unpainted and painted plywood, commercial linoleum tile, concrete board, cotton sheeting, and nylon carpet and are applicable to bed bug product evaluation and comparative testing33 as well as product registration.

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Direct spray bioassay Twenty adult bed bugs, large nymphs (fourth
fifth instars), or 2-day or 3-day-old eggs are placed on the filter paper in each small plastic dish (5.5 cm in diameter and 1.5 cm in height). They are sprayed with a test compound using a Potter spray tower at the application rate of 4.07 mg/cm2 (1 gal/ 1000 ft2). This application rate is a standard “point of run off” and allows for fair comparisons among different compounds.34 Bed bug stages in the control group are sprayed with water. Each treatment is replicated three times. After treatment, bed bugs in their respective treatment groups are immediately transferred to clean 1.5-cm diameter screened plastic Petri dishes with a paper harborage after treatment. The Petri dishes are placed in an incubator at 27 C and 50% RH with a photoperiod of 12L:12D h. Mortality data are taken at 1, 3, 5, 7, and 10 days PT. A bed bug is considered dead if it was not moving or could not right itself when it is prodded with forceps. Dry residue contact bioassay Fabric The test compounds are applied to 10 3 10 cm cardboard panels covered with white 100% cotton fabric at the rate of 4.07 mg/cm2 using a Potter spray tower. The control panels are sprayed with water. After 1 day, bed bugs are released onto the treated fabric and confined with a plastic ring (9 cm in diameter and 2 cm in height) for 5 min. The bugs are then transferred to clean Petri dishes. Each treatment is replicated three times. Mortality is recorded at 1, 3, 5, 7, and 10 days after exposure.34

Filter paper The filter paper method was described in 1968 by Burden and Smittle.35 Basically, test compounds are dissolved in acetone to evaluate their efficacy against adult bed bugs at 0 (control) and serial dilutions (mg/m2). The test compounds are applied uniformly to filter paper (Whatman No. 1, 15 3 55 mm, 11-cm diameter) using a pipette with a volume to achieve the desired test concentration on the filter paper. Three replicate batches of 10 adults (7 females plus 3 males) are used/treatment. Each batch of bed bugs is confined continuously on the filter paper and fed at weekly intervals. At weekly intervals, the filter paper, together with attached eggs, is removed from each Petri dish and immediately replaced with a clean filter paper that had also been subjected to the same treatment at TD 0. The removed filter papers were individually incubated for 2 weeks to determine the viability of the attached eggs. The papers were then discarded. The following parameters are assessed weekly: mortality of adults, number of eggs laid, hatching success of eggs, and morphology and survival of emerging nymphs.36

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Glass: vial test The adult vial test was originally developed by Plapp et al.37 for the adult tobacco budworm, Heliothis virescens, and has been modified for bed bugs.38 This insecticide-coated glass assay was used to determine the susceptibility of adult bed bugs to test compounds. All test compounds were dissolved in acetone and serially diluted. Treated vials were hand rotated until all surfaces within vials had been coated and the acetone had completely evaporated, leaving a uniformly applied insecticidal residue on the inner surface. Vials treated with only acetone were used as controls. For each treatment, 10 adult blood-fed bed bugs were used. Each treatment of 10 bed bugs was replicated 3 times for each concentration. Vials were placed upright in a ventilated cabinet within a fume hood and maintained at a constant temperature of 25 C and 80% RH for 24 or 48 h. Mortality of bed bugs was determined immediately after the 24-h period. A bed bug was considered dead if it was not moving or could not right itself when probed. Percentage mortality was measured as the proportion of 30 bed bugs dead after a 24-h exposure to the test compounds. All data were subjected to probit analysis. The performance of test compounds using three residual dry surface contact bioassays (oil-based insecticide films on filter paper, and acetone-based insecticide deposits on two substrates: filter paper and glass) was assessed against a susceptible strain of C. lectularius and two resistant strains of C. hemipterus.39 Substrate type significantly affected (P , .05) the test compound knockdown response of the susceptible strain in acetone-based test compound bioassays, with longer survival time on filter paper than on the glass surface. With the exception of deltamethrin, the different diluents (oil and acetone) also significantly affected (P , .05) the insecticide knockdown response of the susceptible strain in the filter paper-based bioassays, with longer survival time with acetone as the diluent. The lower effectiveness of the insecticide acetone-based treatment on filter paper may be due to crystal bloom. This occurs when a compound, dissolved in a volatile solvent, is applied onto absorptive surfaces. The effect is reduced on nonabsorptive surfaces and slowed down with oil-based insecticides, whereby the oil forms a film on absorptive surfaces. These findings suggest that if absorptive surfaces are used in bioassays for testing active ingredients, then oil-based insecticides should be preferably used.

Membrane feeding assay The in vitro maintenance technique used successfully to rear C. lectularius by feeding for .2 years all nymphal stages and adults through Parafilm M sealing film on different types of blood can be used successfully to screen compounds in vitro. Using this feeding technique, the subsequent egg production of female bed bugs was remarkably high. The blood was maintained at 37 C to enhance the attachment of the bugs. The effect of anticoagulation

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methods for the blood meal was investigated, and heparinized blood was found the most suitable for feeding bed bugs. All stages of the bed bugs fed weekly on blood in the artificial feeding system remained attached for up to 0.5
1.0 h, until completion of their blood meals, and all reached engorged weights. More than 90% of the bed bugs fed artificially on whole blood, and they molted or laid eggs successful.21 This in vitro screening method allows for the testing of experimental compounds against all life-cycle stages of the bed bug.

Repellency Petri dish assay Plastic Petri dishes (11.4 cm diameter 3 3.8 cm height) are used to quickly evaluate the comparative repellency of test compounds. Filter papers are cut into two equal halves; one half was treated with a repellent using a Potter spray tower at 2.47 mg/cm2 or 0.61 gal/1000 ft2 of ethanol solution. The other half was sprayed with 95% ethanol. A small piece of filter paper was also treated with the same repellent and folded to a tent shape with the treated side facing down. The paper tent was placed on the repellent-treated side and the dishes were left uncovered throughout the assay. In the control dish, one-half of the filter paper and the harborage were treated with 95% ethanol. The other half of the filter paper was not treated. In the repellency assay,40 nine males and six large nymphs were released into each Petri dish. The location of bed bugs in each Petri dish was recorded at 3, 5, 9, and 24 h PT. Each treatment was replicated four times in both assays. The assays were initiated at B2
5 h into the dark cycle. The experiments were conducted in a room at 22 C
26 C and a photoperiod of 12L:12D hour cycle. Repellency indices from Petri dish assays are calculated according to the formula: Repellency index 5 C 2 T/C 3 100, where C is the mean numbers of bed bugs on the treated filter paper halves in all control dishes, and T is the number of bed bugs on treated filter paper half in one test dish.41 Repellency indices are compared using ANOVA followed by Tukey’s HSD test. The bed bug count data in arena assays comparing different chemicals were analyzed. Arena assay This bioassay is to determine if bed bugs avoid contacting substrates treated with selected test compounds.40,42 Paper surgical tape was treated with test compounds using a Potter spray tower at 4.07 mg/cm2. The control tape was treated with water. After 24 h, the tape is placed on the exterior wall of black Climbup Insect Interceptors (10 cm in diameter 3 2.2 cm in height). The interior surface of the Climbup interceptors was coated with a light layer of fluoropolymer resin to prevent trapped bed bugs from escaping. Plastic tray

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arenas (80 3 75 3 5 cm; length by width by height) with a brown paper lined bottom are used. A layer of fluoropolymer resin is applied to inner walls of the arenas to prevent the bugs from escaping. A wooden stool (26.5 cm in length 3 26.5 cm in width) with four legs was placed in each arena. A filter paper (15 cm in diameter) was placed on the center of each arena below the stool, and then a plastic ring (13.3 cm in diameter 3 6.4 cm in height) is placed on the filter paper to confine the bed bugs. A piece of folded cardboard and folded fabric is placed on the filter paper to provide harborages for bed bugs. Four arenas are placed in a nonventilated room at 25 6 1 C and a photoperiod of 12L:12D h. Seventy bed bugs (35 fourth
fifth instar nymphs and 35 adult males) were confined with a plastic ring. The bugs were acclimated for B15 h before the start of the experiment. Each of the four interceptors under each stool was treated with test compound or water. At 1 h after the onset of the dark cycle, CO2 is released from a gas cylinder to the top of the stool at 100 mL/min to stimulate bed bug activity. The plastic ring confining the bugs was removed at 1.5 h after the dark cycle to initiate the experiment. The numbers of bed bugs trapped in the interceptors and those in the arenas are collected and counted after 4 h with the aid of a red light. Most (80%) of the bugs responsive to CO2 stimulation were trapped in the Climbup interceptors within 4 h. Abbott’s formula43 was used to calculate corrected mortality. Percentage corrected mortality, percentage egg hatch and survival, and percentage trap catch values were arcsine square root transformed to meet the assumptions of normality and homogeneity of variances. The repeated-measures analysis of the mortality data was done using a mixed model to determine differences between treatments and their interaction with time. Each source of variation, between and within treatments (day and TD), was included as an effect in the model. Replicates were included as random effect. One-way ANOVA was used when only one observation period was selected to compare the treatments. When the interactions were significant, Tukey’s HSD (α 5 0.05) was used to separate the means. The arena assay examining the changes in repellency of test compounds was analyzed.

Triple-bowl assay This bioassay is designed to evaluate the efficacy of test compound-treated bands for repelling bed bugs under conditions mimicking the natural environment.40 The experimental setup consists of three inverted plastic dog bowls (600 mL in volume and 18 cm in diameter 3 64 cm in height), placed next to one another with a wooden rod serving as a bridge between the three bowls. The inner surfaces of the dog bowls are coated with a layer of fluoropolymer resin to prevent trapped bed bugs from escaping. One piece of filter paper (10 cm in diameter) and a piece of black cloth are placed at both ends of the wooden rod to provide harborages for bed bugs. A piece of cloth is placed at the bottom of the center bowl to allow bed bugs trapped in the bottom to be

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able to climb back to the harborage located at the wooden rod, whereas bed bugs captured in either of the two side bowls cannot return to the harborages associated with the wooden rod. Eight plastic containers, each with 100 bed bugs (B90% adult males and 10% fourth
fifth instar nymphs), are prepared 1 day before the test. The Irvington strain was selected for this experiment because the strain was only kept in the laboratory for 1 month and the bugs were very responsive to host cues. The bioassay utilized 100 bed bugs that were released into the center bowl at 2 h into the dark cycle. After 15 min of acclimation, two wooden rods were placed horizontally between the bowls to allow bed bugs to cross between the bowls. One wooden rod was wrapped with a 2.5-cm-wide repellent-treated fabric tape. The other rod was wrapped with a 95% ethanoltreated fabric band as control. The test compounds were applied to the bands using the same method as described in the arena assay 1 h before the test.40 The experiment was conducted at 27 C
29 C and lighted with a 25 W transparent red light bulb. CO2 (100% concentration) was released from three 5 lb CO2 cylinders each at 100 mL/min to stimulate bed bug foraging movement. Bed bugs would naturally disperse both vertically or horizontally from the center bowl after being stimulated. The three CO2 release points were B1.5 m above the test devices. Eight sets of devices were set up in the room. The number of bed bugs found in the two side bowls was counted after 2 h. Once counted, the bed bugs were returned to the center bowl and the wooden rods removed. The room was vented for 10 min using a fan. The triple-bowl assay examining the changes in repellency of test compounds was analyzed.

Contact—vapor chamber assay—larvae The exposure chamber employed was the following but refer to the original paper for a diagram.44 Half pint card cream jars were the receptacles used (capacity 5 330 mL). A small tin box without its lid was placed at the bottom to contain the insecticide, which, if liquid, was soaked up on several thicknesses of blotting paper. The fleas in larval, cocoon, or adult stages as the case might be were placed in glass-bottomed boxes 1 in. in diameter, either open at the top or with a covering of fine gauze. The box was fastened to the side of the jar by a pin about 2 in. above the bottom so that contact with the insecticide was impossible and any vapor that reached the insects must have risen above the box containing them and have entered it from above. In experiments with the egg stage, the ova were placed on a slip of blotting paper in glass tubes 3 in. high and 1 in. in diameter. The tube was uncorked and stood upright in the card jar, its base being kept from contact with the tin containing the insecticide. The jars were covered with a piece of thin cotton cloth which would allow air to pass. The amount of liquid insecticide used was 0.5 mL or 0.5 g depending on the compound. Larval stages of C. lectularius were tested at 15.6 C
18.6 C and RH at 80%.

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In vivo method(s) Mice Systemic antiectoparasitic activity of test compounds using bed bugs was first reported by Linquist et al.45 and fifth instars of C. lectularius were used to assess the systemic activity of chemicals given PO to rabbits.46 A mouse/bed bug assay for systemic efficacy and safety was used to screen for intrinsic systemic nodulisporamide compounds for use in the monthly control of fleas and ticks on companion animals.47 A murine model using a blood-sucking ectoparasite, for example, bed bug, was used prior to testing suitable derivatives in dogs and cats. Analogs with suitably useful activity in the artificial membrane flea assay subsequently were screened using a rodent model against a blood-sucking ectoparasite: the common bed bug, C. lectularius.48 The C. lectularius assay served to identify not only intrinsic systemic efficacy (ED50 for efficacy) but also functioned as a potential preliminary filter for overt mammalian toxicity. The Schistosoma mansoni/mouse/tail exposure technique49 was adapted to a systemic mouse/bed bug assay. A sheet metal platform was fitted with seven cylindrical plastic containers (24 3 80 mm) with a 7-mm diameter hole in the bottom of the tubes and in the cap. Adult female mice (CD-1, .28 g bw) were fed diets containing test compounds [ppm amounts of “pure” compound dissolved in acetone:DMSO (9:1), ground lab chow (10
20 g) added prior to dryness, 45 g ground lab chow added, and the entire diet mixed in a mechanical grinder to yield dose titrations]. The test compound diets were fed to mice ad libitum for five consecutive days. On the fifth day of diet treatment, the mice were restrained without sedation in cylindrical plastic containers and each mouse tail was quickly dipped in a solvent solution of ethanol:acetone:DMSO (7:2:1), then tap water, followed by a distilled water rinse, and wiped dry with tissue in order to remove as much of the medicated diet from the skin and fur as possible. The tail was inserted into a 17 3 60 mm shell vial containing five, unfed, fourth instar C. lectularius on a piece of filter paper (7 3 25 mm). A plastic foam stopper was used to plug the vial with the mouse tail in place. The mice and bed bugs were held in place on a 15.2 3 30.4 cm stainless steel (24 gauge) sheet metal platform by broom clips and fuse clips. Following 10
20 min exposure on mice in a darkened area, the vials containing the engorged instars were removed, plugged, and incubated at 21.7 C
22.2 C and 60%
70% RH. The engorged insects were examined daily until at least 80% of the controls had molted to the fifth instar. The criteria for activity were death (no detectable movement), paralysis (unable to maintain a grip on filter paper), or molt delay (failure to molt to fifth instar in the same time as 80% of the controls). Each dose (ppm) group rate was compared to the appropriate control group rate with chisquare test. P-values ,.05 were statistically significant.

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The C. lectularius in vivo assay48 utilizes female CD-1 mice (n 5 3) given compounds PO via gavage (four dose levels: 1.0, 0.5, 0.25, and 0.125 mg/kg) 24 h prior to feeding the fourth instars. The single PO dosed compounds were dissolved in a mixture of polyethylene glycol 400 and DMSO (1:2, v/v) and administered at a rate of 0.5 mL/100 g, with concentrations adjusted to provide the specified levels in mg/kg. Following treatment (24 h for single PO doses), the mice were restrained in plastic containers from which their tails were put through a small hole. The tail was washed, wiped dry, and inserted into a shell vial containing five unfed fourth instar C. lectularius. The vials containing the engorged insects were removed, plugged with a foam stopper, and examined daily until 80% or more of the control instars had molted to the fifth instar. Death, paralysis, or molt delay relative to the placebo-treated controls served as the criteria for activity.47

Reduviidae spp. Latreille, 1807—assassin bugs and kissingor cone-nose bugs Biology and life cycle The assassin bug family Reduviidae, subfamily Triatominae of the Hemiptera, contains the Triatomine and kissing bugs that transmit the tropical parasitic disease Chagas disease, also known as American trypanosomiasis, caused by the protozoan Trypanosoma cruzi. The disease was named after the Brazilian physician and epidemiologist Carlos Chagas, who first described the flagellate protozoan in 1909 from the intestines of reduviid bugs.50 T. cruzi is commonly spread to humans and other mammals by the bloodsucking “kissing bugs,” the triatomine bugs, especially the most relevant vectors to humans, Triatoma infestans Klug, 1834 (South America) and Rhodnius prolixus Sta˙l, 1859 (South-Central America and Panama). Other sylvatic triatomine species, such as Panstrongylus geniculatus Latreille, 1811, found in specific South American regions (Venezuela, Colombia, and Argentina) transmit T. cruzi to wild animals (sylvatic cycles) and are putative vectors of T. cruzi to humans. Insect vectors become infected while taking a blood meal from an infected host. The trypanosomes are taken to the posterior part of the insect gut (posterior station or stercorarian trypanosomes), where they develop into amastigotes with a short free flagellum. These spheromastigotes are reproduced by binary fission and move onto the rectal wall of the bug and become the infectious stage (metacyclic trypomastigotes with an undulating membrane). The trypanosome cycle in insects requires about 2 weeks but varies depending on temperature and humidity. The bugs seek out and live in dark, cool places like cracks and crevices inside the home during the day but emerge at night to feed. They tend to feed on the face of humans around the

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eyes and lips (kissing bugs) during sleeping and are able to feed painlessly enough so as not to awaken a sleeping host.51,52 While the bugs are feeding or at the end of a blood meal, the bug defecates passing the infective metacyclic stage in the feces deposited on the skin near the site of the bite. The irritation of the bite or mere presence of the bug causes the person to scratch or rub the bite area, thus mechanically introducing the infective stage into the bite wound or the ocular and oral mucous membranes. The most recognized external mark of acute Chagas’ disease is the unilateral swelling of the face (Roman˜a’s sign) that includes the eyelids on the side of the face near or at the site of the bite wound. Once the metacyclic stage is inside the mammalian host, they invade cells differentiating into intracellular amastigotes which multiply by binary fission. Further intracellular differentiation yields typomastigotes, which are released into the bloodstream of the host. The chronic form of Chagas’ disease may develop over 10
30 years and affects the heart with major cardiac manifestations leading to heart failure, causes “mega” condition of the esophagus and colon, and affects the peripheral nervous system (neuritis resulting in altered tendon reflexes and sensory impairment). Assassin bugs (family Reduviidae) belong to the true bug order, Heteroptera (Hemiptera), and are characterized by a thin necklike structure connecting the narrow head to the body. A characteristic of the family is that the short three-segmented beak is curved and lies in a groove between the front legs. They are hemimetabolous and exopterygote (wing buds develop externally) insects with the immature stages having a similar body form to adults and there is no pupal stage. The total duration of the life cycle of the triatomine bug, from egg to adult, varies from 4 to 24 months, depending on the species and environmental conditions. The most important vector species usually have 1
2 cycles/year. Female bugs are inseminated shortly after they molt and can produce eggs for 1 year after copulation. Females can lay from 80 to 100 eggs in 10
20 days. Embryonic development lasts 10 days. Reduviids develop through five instars to the sexually mature adult and require a blood meal to molt to the next stage. A blood meal takes about 20 min. The third-stage instars have visible wing progenitors and the fifth instar is characterized by developing wings. Female triatomines develop into the adult stage before males, but males are larger than females. Adults can live an average of 8
16 months, although T. infestans in Bolivia has a life span of 3 years. Adults possess thin, transparent wings and can glide over 100 m.

Rearing method(s) Artificial membrane feeding Reduviid bugs can rear a laboratory insectarium at 26 6 1 C and 70 6 1% RH. The bugs are fed with defibrinated human blood using a mosquito

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(Hemotek) membrane
feeding system. A mass rearing apparatus has been described.53 Under laboratory conditions, the mean time from egg to adult was 269 days, with a wide range of duration (174
598 days) and the times required for first, second, third, fourth, and fifth instar development were 33, 37, 41, 61, and 69 days, respectively, with a mortality of 22%. Optimum production was attained with a regimen of 22 days of bug fasting, in which 76% of the nymphs reached the fifth instar with a weight range from 201 to 300 mg.54 Triatoma dimidiata presented a development time with a broad range for some individuals, possibly due to the irregularity in the food availability. A temperature-controlled membrane-feeding apparatus for use with blood-sucking arthropods has been described which is relatively simple in design, efficient, and trouble free.55 Easily constructed from an ordinary Mason-type fruit jar, readily available laboratory glassware, and unsophisticated electrical parts, its construction is economical and involves no special techniques. An electric heater coil for raising temperature of the blood to body heat is built into each feeding device and other than flexible connecting wires, no troublesome linkage as such is used with hydraulic systems employed. By means of a variac (variable autotransformer), the temperature can be adjusted with ease. The feeding arthropods have access to a maximal membrane surface area, so the device is adaptable to mass feeding of colonized vectors. A variety of membranes, either artificial such as a rubber condom, or of animal origin such as sheep cecum or pig intestine, can be employed, depending on the type of arthropod with which the device is used. Arthropods fed using this apparatus included R. prolixus, C. lectularius, Culex pipiens, and Omithodorus moubata. An in vitro artificial mass feeding technique was developed allowing as many as 60 bugs to engorge completely during a 10-min period.56 No differences in molting rate and mortality index were found between the method described here and the rabbit feeding procedure. The method has the advantage in that diets to be used may be in the form of suspension including whole blood nutrient meals. Maintenance of large colonies was found feasible eliminating the rabbit host. R. prolixus was successfully reared in vitro for three generations on a diet of defibrinated pig blood fed through Parafilm or silicone-rubber membranes.57 Reproduction in terms of the number and sizes of eggs produced and survival from egg to adult was superior to and equal to that reported for insects fed on live hosts. The technique appears equally applicable to the rearing of Panstrongylus megistus, T. infestans, and T. brasiliensis. The inadequacy of cow blood as a diet for R. prolixus was manifested in a decrease in adult size after two generations and in a reduction of egg weight from 30 to around 16 mg/female/week. This was accompanied by a corresponding reduction in egg hatch from 90% to about 50%.

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Animal hosts Techniques have been developed for small-scale rearing of R. prolixus on rabbits.58 R. prolixus bugs are presented in tubes closed with silk to the shaved stomachs of rabbits. The bugs are held in 1-lb glass jars at 28 C between feeds. Large-scale rearing by this method was precluded by the amount of handling required. Feeding on defibrinated blood resulted in a survival rate to the adult stage of ,10%, but sheep on which up to 6600 fourth instars could be fed weekly proved satisfactory hosts, although they had to be replaced after 1
9 weeks as they developed a resistance which prevented bug feeding. A large-scale rearing technique involved presenting second through fifth instars to sheep in wire cages covered with nylon netting, while first instars and adults are fed on rabbits; between feeds, the bugs are housed in 3.5
7-lb glass jars or in net-covered cages. A production of 4000 fed fourth instar or larger bugs/week for 18 months was achieved, sometimes reaching 7000/week. Triatomine bugs are fed in containers strapped to the ears of semilop eared rabbits restrained in boxes with only the head protruding.59 The feeding containers are left on the rabbits’ ears for B30 min in a darkened room. The feeding containers for first instars are specially constructed from Perspex, while all other stages are fed in Geigy cages covered with nylon tights. After feeding, the containers are placed on blotting paper for 24 h. The blotting paper absorbs a large quantity of water which the bugs excrete immediately after a blood meal. The bugs are then transferred to Kilner jars containing chromatography paper. These are incubated at 27 C, where they remain until the bugs have molted or laid eggs and are ready for another blood meal. Cast skins and eggs are removed when the bugs are transferred from Kilner jars to feeding containers. To reduce the risk of escapes, bugs are immobilized with CO2 before being transferred from one container to another, and all handling of bugs is performed in trays with an oil surround. Using 25 rabbits, a weekly output of 200
500 fed fifth instars of R. prolixus can be maintained. Tolerance of host animals to loss of blood determines the selection of host for maintenance of large laboratory colonies of Triatoma spp.60 Rabbits tolerate frequent and numerous feedings. Chickens, pigeons, and guinea pigs are widely used laboratory hosts. Wild and laboratory rats and mice must be used with caution to avoid the effects of blood loss. Chickens and pigeons are of special interest as a food source for Triatoma in trypanosometransmission studies. Host species are exposed to the adult insects in a confined chamber either in an inner cage or allowed to move freely within the chamber. The ability of Triatoma spp. to produce viable eggs and larvae and the rate of reproduction are conditioned by the volume of the blood meal and temperature (82% hatch rate of T. infestans eggs at 25 C). First and second instars drown from

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surface contact or immersion in the urine of the host. Hosts can be elevated with special devices for collecting waste or confined to an inner wire basket within the bug cage. Absorbent liners, pleated toweling, or blotters can be used in the feeding jars and culture chambers. Bugs can be confined to the culture jar and fed through gauze covers of nylon mesh from a shaved surface of the host. Laboratory rearing conditions of T. infestans include a temperature range of 18.5 C
28.7 C and 80% RH with females producing 249 eggs. Egg hatch occurs in 27
48 days and nymphal development in 134 days with an adult longevity of 182
470 days.61

In vitro method(s) Contact Filter paper Whatman No. 1 filter paper circles (4 cm diameter) were treated with test compound solutions in acetone to determine estimated EC50 values against first instar nymphs of T. infestans.62 After allowing the solvent to evaporate for 30 min, the circles were placed on the floor of plastic containers (4 cm diameter 3 4 cm high). Groups of 10 nymphs were added to each container, covered with plastic tops, and placed in a chamber with a constant temperature of 28 C and 60%
90% RH. The control groups were exposed to filter papers treated only with acetone. The effects of the test compound (incoordination and knockdown) were recorded 24 h later. The incoordination has been previously defined as the incapacity of the insects to leave a filter paper disk (15 cm diameter).63 At least four different concentrations of a pyrethroid, ranging between 0.07 and 700 μg/cm2, were used to estimate each EC50 value. Each experiment was replicated a minimum of five times and the EC50 values were calculated using the probit method.64 Differences between the values were considered significant (P , .05) when the respective 95% confidence limits did not overlap. Cardboard boxes To test the immediate and persistent activity of LCT under laboratory conditions (27 C and 75%
80% RH) on laboratory reared T. infestans, a series of six experiments was performed.65 The interior surface (400 cm2) of 36 specially manufactured cardboard boxes was uniformly sprayed with 2 mL of the following substrates: 12 with 0.6% LCT suspension, 12 with 1.0% LCT suspension, and 12 with water (controls). At 24 h 30, 90, 150, 270, and 360 days later, in 2 pairs of each 6 boxes, lots of 10 T. infestans fourth nymphs were deposited inside. In each of the six experiments, daily observations of the conditions of the insects (unaffected, affected in terms of various grades of progressive uncoordination, or dead) after exposure to the insecticide were performed to end with the register of these conditions 96 h later. After

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7 and 10 days of exposure, the immediate and persistent effect of LCT was 100% mortality.

Repellency A video-tracking technique was performed in a glass ring (2.5 cm high 3 5 cm diameter).62 The test arena floor was covered with a rectangular piece (9 3 8 cm) of white No. 1 filter paper. Filter papers were treated with 0.5 mL of the test compounds in acetone solutions or with acetone only (control), 30 min before starting each experiment in order to allow the solvent to evaporate. The floor of the test arena was covered with the treated filter paper, and a group of three nymphs of T. infestans was placed on the paper. Rectangular pieces of filter paper were cut into halves (Zones I and II). Zone I was treated with 0.25 mL of acetone, Zone II was treated with 0.25 mL of a DEET solution in acetone (7 or 70 μg/cm2). After the acetone evaporation (30 min), the filter paper halves were fitted together to make a single layer to cover the test arena floor. For the controls, both filter paper halves were treated only with acetone. In order to determine the distribution of the nymphs on the test arena, the viewing field image was divided into two zones using the Multiple Zones Motion Monitor for Videomex software. In each zone, the area (expressed in pixels) occupied by the nymphs during the experiment was recorded. A closed-circuit black and white video camera was placed 15 cm above the center of the test arena. An image analyzer converted the analog signal input from the video camera to digital data. The resolution was 256 3 192 pixel and the acquisition and processing speed was 30 frames/s. In the monitor, the video signal colors are inverted and therefore white objects appeared to be black and vice versa. Thus the presence of insects in the arena was determined by a visual contrast between the individuals (white) and arena background (dark) and scored as the number of pixels. The locomotor activity was recorded using the Multiple Zones Motion Monitor for Videomex software, which records the movement of multiple objects in one area. Each set of data was imported and handled on a personal computer. The distribution of the nymphs was recorded for 30 min. The results were expressed as a repellency coefficient (RC) 5 (1 A(II)/A(I)) 3 100, where A(I) is the area occupied by insects in Zone I, and A(II) the area occupied in Zone II. When RC 5 0, the distribution of the insects is random; as RC values increase, the repellent effect is higher. Each experiment was repeated four times, and the results were analyzed using one-way ANOVA. The level of significance considered was 0.05. The video-tracking technique has been used to evaluate the repellency of volatile compounds against first instars of R. prolixus.66

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The kissing bugs, Triatoma rubida, Triatoma protracta, and Triatoma recurva, are common hematophagous bugs in southeastern Arizona and responsible for severe allergic reactions in some individuals who are bitten.67 They also possess the potential to transmit the blood parasite, T. cruzi. DEET, picaridin, tea tree oil, peppermint oil, and citronella oil were tested for repellency to T. rubida and its ability to probe and feed on a small restrained rat.68 No long-range repellency was observed with any of the test materials. Only citronella oil was able to stop all probing and feeding by T. rubida on a restrained mouse. Further tests were conducted for repellency on mice of the major citronella oil components—geraniol, citronellol, limonene, and citronellal.69 Different concentrations and combinations were tested. All components of citronella oil demonstrated some inhibition of feeding, ranging from very weak inhibition (limonene) to significant inhibition (geraniol and citronellol). A mixture of geraniol and citronellol was found to be repellant at concentrations of 0.165 and 0.165 v%, respectively, for all three triatome species. Citronellol and limonene had no significant repellent activity. The repellent activity of citronella oil appears to be acting through direct contact with the bugs rather than diffusion of vapors.

In vivo method(s) Feeding assays Rat An in vivo rat-Triatoma model has been developed to evaluate the antifeeding and insecticidal efficacies of test compounds.70 All rats were anesthetized with an IP injection of a combination of 90 mg/kg ketamine and 10 mg/kg xylazine. Each rat was placed in a transparent plastic container (50 3 20 3 20 cm3) in which it was then exposed for 1 h to starved nymphal instars (n 5 12) and male and female adults (n 5 4) on TD 1, 7, 14, 21, and 28 PT. Each rat represented the experimental unit. The whole experiment was conducted in a slightly darkened room at a mean 6 SD temperature of 22 6 1 C. After the 1-h exposure period, the bugs were categorized and counted as fed or unfed and dead or live. Live bugs from each group were maintained in separate incubators and survival rates at 24 h after the end of the exposure was recorded. The primary end points were the fed and live bug counts. The arithmetic mean numbers of engorged bugs and live bugs were calculated at each time point. Antifeeding and insecticidal efficacies were calculated using Abbott’s formula.45 Statistical analyses were performed. A rat-Triatoma model71 was used to explore the antifeeding and insecticidal efficacy of a topical ectoparasiticide, dinotefuran
permethrin
pyriproxyfen (DPP), against T. infestans, a vector of T. cruzi, for which dogs are domestic reservoir hosts. Twenty rats (160.7 6 3.6 g) were divided into

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two equal groups: untreated and treated. The minimal recommended dose of DPP was therefore estimated to be 0.07 mL/rat. All doses were administered on TD 0 and were applied directly to the skin as a line-on treatment along the spine using a micropipette to ensure accurate and complete dosing. All rats were anesthetized with an IP injection of a combination of 90 mg/kg ketamine and 10 mg/kg xylazine. Each rat was placed in a transparent plastic container (50 3 20 3 20 cm3) in which it was then exposed for 1 h to starved nymphal instars (n 5 12) and male and female adults (n 5 4) on TD 1, 7, 14, 21, and 28 PT. The whole experiment was conducted in a slightly darkened room at 22 6 1 C. After the 1-h exposure period, the bugs were categorized and counted as fed or unfed and dead or live. Live bugs from each group were maintained in separate incubators and survival rates at 24 h after the end of the exposure was recorded. Insecticidal efficacy was also assessed after incubation of the insects for 24 h postexposure. Antifeeding efficacy was 96.7%, 84.7%, 80.5%, 81.5%, and 42.6% on TD 1, 7, 14, 21, and 28, respectively. Insecticidal efficacy evaluated at 1 and 24 h after exposure on TD 1, 7, 14, 21, and 28 was 100, 91.2, 82.5, 80.0, and 29.1, and 100%, 100%, 100%, 96.0%, and 49.9%, respectively.

Topical A study was conducted to evaluate the mortality and blood intake of T. infestans fed on goats that had been treated with different doses of pour-on insecticide.72 Third-instar nymphs were fed on female goats 1
2-months-old that had been treated with 0, 5, 10 or 15 cm3 of a pour-on formulation of cypermethrin. After feeding, the nymphs were taken to the laboratory for mortality and blood intake measurements. The insects were kept in appropriately labeled plastic jars at 26 C
28 C and 50%
70% RH. Blood intake was calculated as the difference between individual jar weights immediately before and after each feeding divided by the number of living nymphs per jar. The nymphs were weighed as a group and not individually, to avoid excessive manipulation that could artificially increase the mortality rate. The insects were considered dead when they were motionless after 14 days of contacting a treated goat. The exposure of T. infestans to animals treated at different PT intervals revealed a residual activity of the insecticide. The mortality rate in the treated groups was higher than in the control groups until 30 days PT (P 5 .03), except in the group treated with 5 cm3, in which no mortality was detected after 7 days PT. Rainfall affected the triatomicide effect, reducing the time of residual activity. The cypermethrin pour-on treatment decreased the blood intake of T. infestans. Thirty days after the cypermethrin application, nymph mortality was 16 6 13% with both doses (10 and 15 cm3). The 15 cm3 dose did not result in higher insect mortality or increased persistence compared to the 10 cm3 dose.

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References 1. Linnaeus C. Systema naturae per regna tria naturae, secundum classes, ordines, genera, species, cum characteribus, differentiis, synonymis, locis. Tomus I. Editio decima, Reformata. 10th ed. Stockholm: Laurentius Salvius [in Latin]; 1758. p. 1
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31 Available from: https://doi.org/10.1021/pr1000169. 15. Shukla U. Economic zoology. 4th ed. Rastogi; 2009. p. 73. 16. Cannet A, Akhoundi M, Berenger J-M, Michel G, Marty P, Delaunay P. A review of data on laboratory colonies of bed bugs (Cimicidae), an insect of emerging medical relevance. Parasite 2015;22(21):1
7. 17. Quarles W. Bed bugs bounce back. IPM Practitioner. BIRC. 2007;24(3/4):1
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93. 20. Naylor RA, Boase CJ. Practical solutions for treating laundry infested with Cimex lectularius (Hemiptera: Cimicidae). J Econ Entomol 2010;103(1):136
9. 21. Montes C, Cuadrillero C, Vilella D. Maintenance of a laboratory colony of Cimex lectularius (Hemiptera: Cimicidae) using an artificial feeding. J Med Entomol 2002;39(4):675
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22. Garcia ES, Macarini JD, Garcia MLM, Ubatuba FB. Alimentacao de Rhodnius prolixus no Laboratorio. Ann Acad Braz Diene 1975;47(3/4):537
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20. 26. Araujo RN, Costa FS, Gontijo NF, Goncalves TCM, Pereira MH. The feeding process of Cimex lectularius (Linnaeus 1758) and Cimex hemipterus (Fabricius 1803) on different blood meal sources. J Insect Physiol 2009;55(12):1151
7. 27. Davis NT. The morphology and functional anatomy of the male and female reproductive systems of Cimex lectularius L. (Heteroptera, Cimicidae). Ann Entomol Soc Am 1956;49:466
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14. 29. Gilbert IH. Laboratory rearing of cockroaches, bed-bugs, human lice and fleas. Bull WHO 1964;31(4):561
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52. 32. Environmental Protection Agency. Draft product performance test guidelines OCSPP 810.390: laboratory testing methods for bed bug pesticide products. Environmental Protection Agency EPA712; 2012. 33. Todd RG. Efficacy of bed bug control products in lab bioassays: do they make it past the starting gate? Am Entomol 2006;52:113
16. 34. Singh N, Wang C, Cooper R. Potential of essential oil-based pesticides and detergents for bed bug control. J Econ Entomol 2014;107(6):2163
70 Available from: https://doi.org/ 10.1603/EC14328. 35. Burden GS, Smittle BJ. Laboratory methods for evaluation of toxicants for the bed bug and the oriental rat flea. J Econ Entomol 1968;61(6):1565
7. 36. Naylor R, Bajomi D, Boase C. Efficacy of (S)-methoprene against Cimex lectularius (Hemipters: Cimicidae). H-8200 Veszpre´m, Pa´pai u´t 37/a In: Robinson WH, Bajomi D, editors. Proceedings of the sixth international conference on urban pests. Hungary: OOK-Press Kft.; 2008. p. 115
21. 37. Plapp FW, McWhorter GM, Vance WH. Monitoring for pyrethroid resistance in the tobacco budworm in Texas-1996. Proceedings of the 1987 beltwide cotton production conference. Memphis, TN: National Cotton Council; 1996. p. 324
6. 38. Steelman CD, Szalanski AL, Trout R, McKern JA, Solorzano C, Austin JW. Susceptibility of the bed bug Cimex lectularius L. (Heteroptera: Cimicidae) collected in poultry production. J Agric Urban Entomol 2008;25(1):41
51. 39. Dang K, Veera Singham G, Doggett SL, Lilly DG, Lee CY. Effects of different surfaces and insecticide carriers on residual insecticide bioassays against bed bugs, Cimex spp. (Hemiptera: Cimicidae). J Econ Entomol 2017;110(2):558
66. 40. Wang C, Lu L, Zhang A, Liu C. Repellency of selected chemicals against the bed bug (Hemiptera: Cimicidae). J Econ Entomol 2013;106(6):2522
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41. Todd R. Chapter 9-Repellents for protection from bed bugs: the need, the candidates, safety challenges, test methods, and the chance of success. In: Paluch GE, Coats JR, editors. Recent developments in invertebrate repellents. ACS symposium series. Washington, DC: American Chemical Society; 2011. p. 137
50. 42. Singh N, Wang C, Cooper R. Potential of essential oil-based pesticides and detergents for bed bug control. J Econ Entomol 2014;107(6):2163
70. 43. Abbott WS. A method of computing the effectiveness of an insecticide. J Econ Entomol 1925;18:265
7. 44. Bacot AW. LXXI. The effect of the vapours of various insecticides upon fleas (Ceratophyllus fasciatus and Xenopsylla cheopis) at each stage in their life-history and upon the bed bug (Cimex lectularius) in its larval stage. J Hyg (Lond) 1914;13(Suppl):665
81. 45. Linquist AW, Knipling EF, Jones HA, Madden AH. Mortality of bed bugs on rabbits given oral dosages of DDT and pyrethrum. J Econ Entomol 1944;37:128. 46. Adkins Jr FS, Sowell WL, Arant FS. Systemic effect of selected chemicals on the bed bug and lone star tick when administered to rabbits. J Econ Entomol 1955;48:139
41. 47. Meinke PT, Colletti SL, Fisher MH, Wyvratt MJ, Shih TL, Ayer MB, et al. Discovery of the development candidate N-tert-butyl nodulisporamide: a safe and efficacious once monthly oral agent for the control of fleas and ticks on companion animals. J Med Chem 2009;52:3505
15. 48. Ostlind DA, Cifelli S, Conroy JA, Mickle GW, Ewanciw DV, Andriulli FJ, et al. A novel Cimex lectularius rodent assay for the detection of systemic ectoparasiticide activity. Southwest Entomol 2001;26:181
5. 49. Campbell WC, Bartels E, Cuckler AC. A method for detecting therapeutic activity against Schistosoma mansoni in mice. J Parasitol 1978;64:699
707. 50. Chagas C. “Neue Trypanosomen”. Vorl¨aufige Mitteilung Arch Schiff Tropenhyg. 13: 120
122. Dr. Chagas named the trypanosome in honor of Oswaldo Cruz, noted Brazilian doctor and epidemiologist Chagas C (1909). “Nova tripanozomiase humana: Estudos sobre a morfolojia e o ciclo evolutivo do Schizotrypanum cruzi n. gen., n. sp., ajente etiolojico de nova entidade morbida do homem [New human trypanosomiasis. Studies about the morphology and life-cycle of Schizotripanum cruzi, etiological agent of a new morbid entity of man]”. Mem Inst Oswaldo Cruz 1909;1(2):159
218 Available from: https://doi.org/ 10.1590/S0074-02761909000200008. ISSN 0074-0276. (in Portuguese with German full translation as “Ueber eine neue Trypanosomiasis des Menschen.“). 51. Schwarz A, Medrano-Mercado N, Schaub GA, Struchiner CJ, Bargues MD, Levy MZ, et al. An updated insight into the sialotranscriptome of Triatoma infestans: developmental stage and geographic variations. PLoS Negl Trop Dis 2014;8(12):e3372 Available from: https://doi.org/10.1371/journal.pntd.0003372. 52. Lima MS, Carneiro AB, Souto-Padron T, Jurberg J, Silva-Neto MAC, Atella GC. Triatoma infestans relies on salivary lysophosphatidylcholine to enhance Trypanosoma cruzi transmission. Acta Tropica 2018;178:68
72 Available from: https://doi.org/10.1016/j.actatropica.2017.10.022. Epub 2017 Oct 28. 53. Aldana E, Otalora F, Abramson CI. A new apparatus to study behavior of triatomines under laboratory conditions. Psychol Rep 2005;96(3):825
32. 54. Reyes M, Angulo VM. [Life cycle of Triatoma dimidiata latreille, 1811 (Hemiptera, Reduviidae) under laboratory conditions: production of nymphs for biological tests]. Biomedica 2009;29(1):119
26. [Article in Spanish]. 55. Pipkin A, Connor T. A temperature-controlled feeding apparatus for hematophagous arthropods. J Med Entomol 1968;5:507
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56. McGuire EJ, Habowsky JEJ, Lumb G, De La Iglesia FA. An experimental approach to the study of drugs in invertebrate systems. I. Mass feeding of Rhodnius prolixus Stahl (Hemiptera, Reduviidae). Can J Zool 1973;51:315
18. 57. Langley PA, Pimley RW. Rearing triatomine bugs in the absence of a live host and some effects of diet on reproduction in R. prolixus Stȧl, 1859 (Hemiptera: Reduviidae). Bull Entomol Res 1978;68:243
350. 58. Gardiner B, Maddrell H. Techniques for routine and large-scale rearing of Rhodnius prolixus Stal (Hem., Reduviidae). Bull Entomol Res 1972;61:05
515. 59. Patterson JW, Maudlin I, Marashi MH. Proceedings: mass rearing of triatomine bugs on rabbits. Trans R Soc Trop Med Hyg 1974;68(1):12. 60. Wood SF. Laboratory culture of Triatoma (Hemiptera, Reduviidae). Bull WHO 1964;31:579
81. 61. Hack WW. Estudios sobre biologia del Triatoma infestans (Klug 1834) (Hemiptera, Reduviidae). An Inst Med reg 1955;4:125
47. 62. Sfara V, Zerba EN, Alzogaray RA. Toxicity of pyrethroids and repellency of diethyltoluamide in two deltamethrin-resistant colonies of Triatoma infestans Klug, 1834 (Hemiptera: Reduviidae). Mem. Ist Oswaldo Cruz 2006;101(1). Available from: https://doi.org/10.1590/ S0074-02762006000100017. 63. Alzogaray RA, Zerba EN. Comparative toxicity of deltamethrin and cis-permethrin on first instars of Triatoma infestans (Hemiptera: Reduviidae). J Med Entomol 1996;33:58
62. 64. Litchfield JT, Wilcoxon FJ. A simplified method of evaluating dose-effect experiments. J Exp Ther 1949;96:99
110. 65. Schenone H, Rojas A. [Laboratory study on the immediate and persistent insecticide activity of the pyrethroid lambda-cyhalothrin on nymphs of IV instar Triatoma infestans]. [Article in Spanish]. Bol Chil Parasitol 1992;47(1-2):35
7. 66. Lutz A, Sfara V, Alzogaray RA. Repellence produced by monoterpenes on Rhodnius prolixus (Hemiptera: Reduviidae) decreases after continuous exposure to these compounds. J Insect Sci 2014;14(1):254 Available from: https://doi.org/10.1093/jisesa/ieu116. 67. Klotz JH, Dorn PL, Logan JL, Stevens L, Pinnas JL, Schmidt JO, et al. “Kissing Bugs”: Potential disease vectors and cause of anaphylaxis. Clin Inf Dis 2010;50(12):1629
34. 68. Terriquez JA, Klotz SA, Meister EA, Klotz JH, Schmidt JO. Repellency of DEET, picaridin, and three essential oils to Triatoma rubida (Hemiptera: Reduviidae: Triatominae). J Med Entomol 2013;50(3):664
7. 69. Zamora D, Klotz SA, Meister EA, Schmidt JO. Repellency of the components of the essential oil, citronella, to Triatoma rubida, Triatoma protracta, and Triatoma recurva (Hemiptera: Reduviidae: Triatominae). J Med Entomol 2015;52(4):719
21 Available from: https://doi.org/10.1093/jme/tjv039. Epub 2015 Apr 17. 70. Diotaiuti L, Faria Filho OF, Carneiro FCF, Dias JCD, Pires HHR, Schofield CJ. Aspectos operacionais do controle do Triatoma brasiliensis. Cad Saude Publ 2000;16(2):61
7. 71. Tahir D, Davoust B, Varloud M, Berenger JM, Raoult D, Almeras L, et al. Assessment of the anti-feeding and insecticidal effects of the combination of dinotefuran, permethrin and pyriproxyfen (Vectras 3D) against Triatoma infestans on rats. Med Vet Entomol 2017;31 (2):132
9. 72. Amelotti I, Catala´ SS, Gorla DE. The residual efficacy of a cypermethrin pour-on formulation applied on goats on the mortality and blood intake of Triatoma infestans. Mem Inst Oswaldo Cruz 2012;107(8):1011
15.

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Chapter 3h

Arthropoda, Pentastomida Alan A. Marchiondo, MS, PhD Adobe Veterinary Parasitology Consulting LLC, Santa Fe, NM, United States

Arthropoda Pentastomida Linguatulidae

¨ Lingulatula serrata Frohlich, 1789—tongueworm Biology and life cycle Pentastomids or tongueworms are dioecious obligate highly specialized arthropods that live as adults in the respiratory tract of predacious reptiles, birds, and mammals. Lingulatula serrata adults live in the nasal passages and sinuses of dogs and cats. It is found worldwide, especially in warm subtropical and temperate regions, such as Asia, the Middle East, Europe, North Africa, and North and South America. Dogs, cats, wolves, foxes, and other carnivores serve as definitive hosts. Adult pentastomids are dorsoventrally flattened and have the form of a vertebrate tongue, hence the name “tongueworm.” Males measure 18
20 μm in length. Females measure 80
120 mm and begin producing eggs in about 6 months. Both males and females have an external segmented or annulated abdomen with an outer skeleton of chitin. The ventral surface of adults has a small sucking mouth flanked by two pairs of hooks. Eggs (70
90 μm) containing larvae are discharged from an infected host with the nasal secretions or passes in feces if swallowed. Cattle, water buffalo, sheep, goats, rabbits, rodents, pigs, camels, and other animals can serve as intermediate hosts.1 If the eggs are ingested by an intermediate host, larvae hatch in the small intestine, penetrate the intestinal wall, develop, and encyst in lymph nodes, lungs, and liver as nymphs.2,3 Larvae have a suboral gland whose duct pores shed their secretion caudally of the mouth onto the tegument, possibly involved in invasion into the intermediate host.4 This stage has an annulated body with two pairs of retractable hollow fangs or hooks.5 Encysted larvae can live up to 2 years in the intermediate host before they calcify and die. When the encysted larvae are eaten by a definitive host, they attach in the upper digestive tract or travel there from the stomach to reach the nasopharynx. No treatment has been attempted for infections other than surgical removal of the larvae.

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Rearing method(s) Eggs of the pentastomid species, Porocephalus crotali, have been used to experimentally infect intermediate rodent hosts.6,7 Infected intermediate hosts can then be used to experimentally infect definitive snake hosts. Infective seventh instar of P. crotali, dissected from the tissues (abdominal fat and lungs) of rat intermediate hosts, has been cultured in vitro to the adult male and female instars (stages that normally reside in the lungs of the rattlesnake definitive hosts).8 Instars (100) at a density of 2 worms/mL of culture medium [washed human RBCs resuspended in bovine serum (50:50 v/v) with 20% MEM and antibiotics] are incubated at 28 C and 5% CO2 in 500 mL bottles rotated slowly on a Rollacell system. Medium is replenished every 2
3 days. Male adult instars developed in three molts, while females required four molts. Natural mortality was about 10% over a 160-day period with 33%
51% of females and 62%
70% of males reaching the terminal instars. Male instars appeared to develop just as they would in naturally infected rattlesnakes. However, females never achieved full size nor copulated in vitro. But, some females became patent. The life cycle of Armillifer agkistrodontis has been established in a multihost model, that is, mice and rats as intermediate hosts, and snakes (Agkistrodon acutus and Python molurus) as definitive hosts.9 This pentastomid is similar to L. serrata with a morphological difference in the number of abdominal annuli. An adult gravid female A. agkistrodontis was isolated9 and mature eggs recovered after culture [washed human RBCs resuspended in bovine serum (50:50 v/v) with 20% MEM and antibiotics] for 1
2 h in an incubator at 26 6 0.5 C and 80% RH. Female Kunming mice (6
8-weeks old, 20 6 5 g) and SD rats (6
8-weeks old, 100 6 5 g) are infected PO with 40 and 80 mature eggs, respectively, using a plastic tube reaching the stomach. One rabbit and one dog were also infected, but no larvae were recovered, while mice and rats had mean recovery rates of 79.2% and 51.2%, respectively. Snakes (A. acutus and P. molurus) are fed experimentally infected mice confirmed to be infected with A. agkistrodontis by ELISA and isolation of parasite larvae. Fecal examinations were conducted on the snakes from day 60 PI onward. Snakes were dissected to determine worm burdens once fully developed eggs were recovered in their feces. Mice and rats developed larvae in 11 week PI of eggs with larvae reaching full infectivity after 16 weeks. Mature eggs first appeared in the feces of infected snakes after a prepatent period of about 10 months. Adult parasites mainly resided in the abdominal cavity of the snakes about 5
6 month PI with a shift to the lungs about 9
10 month PI. The entire life cycle under laboratory conditions was about 14 months. Nymphs and adults of Sebekia mississippiensis obtained by dissection from naturally infected mosquitofish and alligators, respectively, were cultured in vitro.10 Pentastome stages were cultured in two different systems: (1) EMEM and 10% FCS and (2) DMEM without serum. Both culture

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systems contained 10,000 IU/mg/mL penicillin and 10,000 μg/mL streptomycin and were changed weekly and maintained at 30 C in 5% CO2. Nymphs maintained in either culture system remained viable and infective to hamsters or mice for at least 1 week. Nymphs maintained in physiological saline (0.85% NaCl) at 30 C died over a 1-week period. Adults were more difficult to maintain than nymphs. Eggs deposited in vitro and held in saline at 4 C contained live larvae that survived for periods as long as 2 months. Attempts to infect hatchling alligators with eggs obtained from in vitro cultures were unsuccessful.

In vitro method(s) No specific in vitro methods for testing parasiticides against the stages of L. serrata were found in the literature. However, the models developed for related pentastomid species8,9 might be used to evaluate parasiticides in vitro.

In vivo method(s) No specific in vivo methods for testing parasiticides against the stages of L. serrata were found in the literature or known to the author. Dogs and cats infected with pentastomids might be diagnosed using an ELISA test;11 however, fecal egg counts can be used. Mice can be artificially infected with 40 eggs of a pentastomid, A. agkistrodontis, for 25
37 weeks to allow for nymph encystment in internal organs.12 Infected mice are treated PO with parasiticides and necropsied 1
3 week PT. Praziquantel, mebendazole, tribendimidine, ivermectin, artemether, and dihydroartemisinin exhibited no activity against A. agkistrodontis in infected mice. Experimentally infected mice and rats can be used to test compounds against the developing larval stage, while experimentally infected snakes could be used to test the development of pentastomids to adults in a definitive host.

References 1. 2. 3. 4.

Riley J. The biology of pentastomids. Adv Parasitol 1986;25:45
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95. John DT, Petri Jr. WA. 9th ed. Medical parasitology, 336-337. Elsevier; 2006. p. 14. Stender-Seidel S, Thomas G, Bockeler W. Investigation of different ontogenetic stages of Raillietiella sp. (Pentastomida: Cephalobaenida): suboral gland and frontal gland. Parasitol Res 2000;86:385
400. 5. Bowman DD, Lynn RC, Eberhard ML, Alcaraz A. Georgi’s parasitology for veterinarian’s. 8th ed. St. Louis, MO: Saunders; 2003. p. 76. 6. Penn Jr. G. The life history of Porocephalus crotali, a parasite of the Louisiana muskrat. J Parasitol 1942;28(4):277
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7. Esslinger JH. Development of Porocephalus crotali (Humboldt, 1808) (Pentastomida) on experimental intermediate hosts. J Parasitol 1962;48(3):452
6. 8. Buckle AC, Riley J, Hill GF. The in vitro development of the pentastomid Porocephalus crotali from the infective instar to the adult stage. Parasitology 1997;115(Pt 5):503
12. 9. Chenn S-H, Liu Q, Zhang Y-N, Chen J-X, Li H, Chen Y, et al. Multi-host model-based identification of Armillifer agkistrodontis (Pentastomida), a new zoonotic parasite from China. PLoS Negl Trop Dis 2010. Available from: https://doi.org/10.1371/journal. pntd.0000647. 10. Boyce WM, Courtney CH, Wing SR, Kurose EW. In vitro maintenance of the pentastome Sebekia mississippiensis. Proc Helm Soc Wash 1987;54(2):265
6. 11. Jones DA, Riley J. An ELISA for the detection of pentastomid infections in the rats. Parasitology 1991;103(Pt 3):331
7. 12. Xu LL, Xue J, Zhang YN, Qiang HQ, Xiao SH. Effect of six anthelmintics in oral treatment of mice infected with Armillifer agkistrodontis nymphs. [Article in Chinese]. Zhongguo Ji Sheng Chong Xue Ji Sheng Chong Bing Za Zhi 2010;28(4):277
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