Arachnida

Chapter 4

Arachnida Chapter 4a

Arachnida, Metastigmata, Argasidae 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

Arachnida Metastigmata Argasidae The family Argasidae is generally considered to include 198 species in two families, Argasidae (genus Argas) and Ornithodorinae (genera Otobius, Ornithodoros, Anticola, and Nothoaspis).1 The typical argasid life cycle is a multihost feeding pattern or cycle2 with the exceptions of Ornithodoros lahorensis that is a two-host tick and Otobius megnini that is a one-host tick. The nymphal cycle of host contact, rapid feeding, engorgement, detachment, and ecdysis occurs several times in niche environments (nests or burrows of hosts), and the precise number of nymphal stages is indeterminate, but eight is the highest number recorded.3 The scutum is absent, the body of adult soft ticks is dorsoventrally flattened appearing ovoid from the dorsal surface with anteroventrally located mouthparts. Nymphs and adults have eight legs, while the larvae have six legs. Female ticks measure up to 1.2 cm and males are usually smaller measuring 7.5 mm. Nymphs measure 3.5
6.5 mm in length.4 All development stages are hematophagous. Unlike ixodid nymphs and adults that feed once during each stage taking several days to complete engorgement, nymphs and adult soft ticks feed repeatedly within a minutes or hours. Nymphs attach for 5
10 days, while nymphs and adults only attach for B30 min. Females oviposit small egg masses (,500 eggs/cycle) in hidden places after each blood meal and Parasiticide Screening, Vol 1. DOI: https://doi.org/10.1016/B978-0-12-813890-8.00004-3 © 2019 Elsevier Inc. All rights reserved.

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larvae hatch in 2
6 weeks. The duration of the entire life cycle can be from 3
6 months up to 2
3 years. Soft ticks are drought resistant. Diapause plays a major role in the regulation of the development of many argasid species that must survive in empty nests or burrows for extended periods of time until a host returns or arrives.5,6 Argasid ticks generally are not considered major species for the determination of efficacy in the acaricide/drug development process. Testing of compounds for control of the parasites typically consider species of major veterinary and human medical concern. These tests would entail dose titration/determination studies followed by a series of dose confirmation studies in both the laboratory and field. Testing of an acaricide/drug for activity against minor species such as the argasids entail testing a dose and formulation already determined to be efficacious against major parasite species, such as pyrethroids, and for which the dose and formulation have been shown to be safe for the target animal species, for human consumption of tissue from treated animals, and for the environment. However, the use of soft ticks in the discovery of systemic parasiticides has become a major component of screening paradigms as discussed in the “In vitro method(s)” section of Ornithodoros species.

Otobius megnini—Duge´s, 1884—spinose ear tick Biology and life cycle The spinose ear tick is generally associated with semiarid and arid environments. It is the most common soft tick that parasitizes domestic mammals in North America, especially prevalent in arid regions of the southwestern United States. The eastern boundary of their range is B97th meridian west.7 It has been reported in Argentina, Brazil, Chile, Mexico, Australia, South Africa, and India.8 This spiny soft tick has a single host life cycle in which the larval and two nymphal stages feed deep within the external ear canal of their definitive hosts (cattle, sheep, horses, dogs, cats, lions, goats, hogs, coyotes, deer, mountain sheep, rabbits, ostrich, peccary, and pronghorn sheep) for long periods of time. The inner ear can become ulcerated and the outer ear excoriated and raw. When the second-stage nymphs have completed feeding, they drop from the host to the ground and molt to adults. The adult ticks are nonparasitic, have vestigial hypostomes, do not feed, and live in the environment of the host. They appear to have a violin shape due to a constriction at the middle of their body. The cuticle is covered with spines. Female ticks lay up to 1500 eggs in a 2-week period and can survive for as long as 2 years. Eggs are laid on or near the ground in crevices in walls and around feeding troughs. Eggs can remain viable for up to 6 months, and the incubation period can be about 18 days or more. Only the larvae and nymphs are parasitic. Newly emerged, unfed larvae measures B0.5 mm in size and can live without feeding on a host for more

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than 2 months. The body is oval with six long legs. Engorged larvae measure B4.0 3 2.5 mm.9 Larvae crawl up vegetation, fence posts, and feed bunks to await a host. Once on a host, the larvae crawl to the ear and attach just below the hair line. Larvae engorge for 5
10 days in the ear, molt to the first nymphal stage, which can feed for periods of B30 days or as long as 7 months. The first nymphal stage is similar to newly emerged larvae, but with more slender legs. The second nymphal stage has a blueish gray body with pale yellow legs (four pairs), mouthparts, and spines. The second nymphal stage reattach in the ears of the infested host and feed for B3
4 weeks, growing to B8 mm. Engorged nymphs then leave the ear, drop to the ground where they hide, and undergo the final molt to the adult tick.

Rearing method(s) A tick colony was initiated from fully engorged nymphs collected from horses and maintained at 10 C, 22 C, and 28 C.10 Immature stages were reared on New Zealand rabbits. Only larvae weighing .0.9 mg (84.9%) molted to nymphs with a premolting period of 5.6 6 1.7 days. None of the larvae maintained at 10 C 6 1 C molted. Larval survival was significantly higher at 28 C 6 1 C (44.2 6 4.3 days) than at 22 C 6 1 C (35.6 6 9 days). At 28 C 6 1 C the majority of nymphs (95.6%) weighing over 10 mg molted into adults within 12.5 6 3.1 days. Nymphs kept at 10 C 6 1 C underwent diapause for 81.1 6 72.0 days. Larger females ( . 13 mg; 93.0%) laid eggs for 44.6 6 17.8 days with a mean preoviposition period of 10.3 6 5.1 days. Females survived longer (369.8 6 128.3 days) than males (210.4 6 54.1 days; Student’s t-test; t 5 5.9, P 5 .001). Some females that laid eggs were parthenogenic. O. megnini successfully completed the life cycle within 123 days and had only one nymphal instar. O. megnini ticks were collected from cattle in Argentina to study the biological parameters of larvae, nymphs, and adult ticks.11 Groups of nymphs were also maintained at three different photoperiods at 25 C. The abundance of immature stages was greatest during January
April and August
October in the first and second year of the study, respectively. No larvae successfully molted. Nymphs weighing ,17 mg also failed to molt, but 89% of heavier nymphs molted into adults. Nymphs molting to males weighed less (49.5 6 16.09 mg) than nymphs molting to females (98.1 6 34.08 mg). The premolt period was similar for nymphs molting to either sex and was temperature dependent, in that, for nymphs that molted to females the premolt period was significantly longer (P , .01) for nymphs maintained at 25 C compared with nymphs kept at 27 C. No effect of photoperiod on the premolt periods of nymphs was detected. Female ticks produced a mean of 7.0 6 1.94 egg batches after a preoviposition period of 16.4 6 8.41 days for the first batch. The mean oviposition period was 61 6 20.8 days, and the duration of oviposition for each batch varied from 1 to 6 days. The mean number of eggs per batch was

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93.1 6 87.53. The minimum incubation period for the first egg batch was 13.6 6 2.77 days. The total number of eggs laid by each female was 651.6 6 288.9. Parthenogenesis was not observed. The REI (number of eggs laid/ weight of female in mg) was 5.5 6 1.26. Pearson’s correlations showed a significant direct relationship between the weight of the females and number of eggs laid (P , .01) and REI (P , .05). Engorged nymphs were collected from the ears of cattle in the United States (Oklahoma), held at 26 C 6 1 C and 85% 6 2% RH until molting to the nonfeeding adult stage. Eggs collected from these adults were held until eggs hatched, and resulting larvae were used to infest both cattle and rabbits. The mean duration of feeding (engorgement time) for larvae and nymphs on cattle were 4.2 6 2.1 and 38.2 6 7.4 days, respectively.12 The mean number of eggs laid/female (789 6 199.5) was similar to that reported by Nava et al.11 from Argentina. A debris-filtering method was developed to collect nonfeeding adult O. megnini off of ungulate hosts.13 To sample for host-seeking larvae a CO2 trap was developed using compressed CO2 released through tubing along with cotton fabric used for tick attachment. Both methods proved successful for collecting adults and larvae from animal shelters, and for larvae from oak leaf litter away from any structure. Locating both life stages off the host is the first crucial step toward the management of this tick species and facilitates collection of all stages for in vitro and in vivo testing.

In vitro method(s) No contact exposure tests of parasiticides against adult or nymphs of O. megnini were found in the literature, including the annotated bibliography of the spinose ear tick by Keirans and Pound.8 However, a filter paper method was evaluated for the repellency of test compounds by exposing stages of O. megnini.14 The movement of ticks was evaluated at intervals of 30 seconds for 5 min after introduction on the treated surfaces.

In vivo method(s) Drummond et al.15 summarized contact exposure control methods for O. megnini prior to 1967 and evaluated the efficacy of 26 insecticides applied directly into infested ears of cattle in the formulations of dusts, suspensions of wettable powders, emulsifiable concentrates, or as technical acaricides. Systemic exposure of ivermectin was ineffective in controlling nymphs of O. megnini in the ears of 12 yearling Shetland ponies.16 Similarly, a second-generation avermectin, eprinomectin, was shown to be ineffective against O. megnini when administered to cattle in a pour-on formulation at the approved label dose rate of 0.5 mg/kg bw.17

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Argas persicus—Oken, 1818—fowl or poultry tick Biology and life cycle Argas persicus is distributed worldwide with a preference for warmer climates, such as the tropics.5 It is a small soft tick that parasitizes domestic poultry (chickens, ducks, and geese) distributed along the Gulf Coast and Mexican border in the United States. The life cycle from egg to adult can be completed within 30 days, if suitable hosts are available for blood meals, or may be prolonged without suitable hosts. Larvae and nymphs can survive for months without a blood meal, and the adult for more than 2 years.6 The adult tick is yellow to reddish-brown when unfed, but this becomes blue in color (often called the blue tick) when engorged. The female is about 8
10 mm in length and larger than the male (5 mm) with leathery, mammillated, and wrinkled dorsal, and ventral surfaces. They can be found on the host or in their nests or in cracks and crevices of commercial poultry buildings and sheds. They transmit the spirochete of Borrelia anserina, the cause of avian spirochetosis. Female ticks often lay eggs in clutches of 25
100 off the host after each blood meal. Mating and the laying of eggs occur off the host, particularly in bird nests. Six-legged larvae hatch from the eggs in 1
4 weeks, seek out a host, and feed for a few hours up to 5 days. The fed larvae leave the host for a sheltered area. The larvae molt to first-instar nymphs in about a week and seek a second host (same individual as the first host and likely the same species) to feed usually at night. There is one larval and at least two nymphal stages (although from three up to seven stages may occur) with the last stage molting to the adult. The life cycle takes B30 days under optimal warm and humid conditions. Other Argas tick species that parasitize avian hosts include Argas reflexus, the pigeon tick, and Argas walkerae, the chicken tick.

Rearing method(s) An SOP has been established by Thangamani18 for the rearing and maintenance of argasid ticks in the laboratory.

In vivo tick feeding procedures Feeding procedures are conducted on a white tray with double-sided masking tape around the periphery of the tray, which serves as a containment barrier for escaping ticks. Mice are anesthetized or restrained in a small mesh cage (without anesthesia) according to an approved animal care protocol during tick feeding. Tick vials are chilled to immobilize the ticks. Larval ticks are painted onto the neck and upper back of the mouse using a fine brush.18 Nymphal and adult ticks are placed on the neck and upper back of the mouse using blunt end forceps.18 Ticks feed to repletion within 30
60 min and

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drop-off the mouse. The fed ticks are collected with blunt end forceps (without puncturing) and transferred to a fresh glass vial for housing inside an incubator.

Tick housing Ticks are housed in a sterile clear-glass vials with a plastic snap cap with several 2 mm ventilation holes and the lid secured with fine mesh. Sterile mouse bedding material is added to the vials to allow the ticks a substrate to burrow.18 The storage vials are stored inside a glass desiccator with Vaseline applied around the rim and containing a saturated salt solution in the desiccator basin (saturated potassium nitrate and potassium sulfate will provide 94%
97% RH).18 Alternatively, plaster of Paris and charcoal can be used at the vial base to maintain humidity.18 Storage vials are labeled with the tick species, sex in the case of adult ticks, the number of individuals for each life cycle stage, and the date of collection.18 Desiccators are stored in an environmental chamber at 23 C and 95% RH with 10L:14D h cycle (this information is specific for Ornithodoros turicata while some species might require different housing conditions).18 Other laboratory rearing methods of A. persicus have been published.19,20 In vitro method(s) Contact—filter paper—adult Three acaricides (permethrin, propoxur, and diazinon) were tested against A. persicus ticks in a test of susceptibility and in a multiple choice test.21 A mixture of guanine hydrochloride and diatomaceous earth in saline was used as an attractant in the bioassays, causing 53.1%
95.7% assembly. The attractant was mixed with acaricides to reduce their repellency and enhance their acaricidal efficiency in bioassays. Permethrin was the most toxic (LC95 at TD 7 5 0.5
1.4 mg/m2 depending on the developmental stage) and was the most repellent acaricide. The mortality of males in the bioassay was significantly higher (76.7%
94.3%, P , .01) when acaricide in amounts of 16 and 160 µg/filter paper disk were mixed with attractant (0.5 mg/filter paper disk) instead of acaricide alone (20%
45.7% mortality only). The mean permethrin residue on the tick body at the end of the bioassay with the acaricide
attractant mixture was significantly higher (13.62 6 11.64 ng) than in experiments without the attractant (,1 ng). Propoxur was less toxic (LC95 at TD 7 5 0.9
1.9 mg/m2) and diazinon the least toxic (LC95 at TD 7 5 2
9.4 mg/m2), both being not or only slightly repellent. Male and female ticks also assembled on filter paper disks treated with propoxur without an attractant. Diazinon displayed significant mortality only in amounts of 0.1 and 1 mg/filter paper disk with or without the attractant. Therefore the

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repellency of permethrin can be reduced and its effectiveness enhanced when used in a mixture with an attractant. No similar effect was observed with propoxur or diazinon.

In vivo method(s) The acaricidal efficacy, both in vitro and in vivo, of peracetic acid and deltamethrin was tested separately against A. persicus. The in vitro bioassay utilized a dipping technique.22 Peracetic acid (0.5%) was highly effective against larval A. persicus, resulting in 100% mortality after 2 min. The lethal concentrations LC50 and LC95 were 0.3% and 0.5%, respectively. The lethal time values LT50 and LT95 were 5.3 and 40 min, respectively, after treatment with peracetic acid (0.25%). About 2 min after exposure to deltamethrin, LC50 and LC95 values were 0.0033% and 0.0052% (33.2 and 51.5 mg/L), respectively. The LT50 and LT95 values were 27 and 305 min, respectively, after treatment with 0.0025% deltamethrin (25 mg/L). After in vivo dipping in peracetic acid (0.5%) the chickens did not show respiratory signs or inflammation of the eyes and/or skin. By contrast, temporary coughing, sneezing, and ocular inflammations without dermatitis were observed in chickens dipped in deltamethrin (0.005% or 50 mg/L). After 7 days PT, the reduction in the percentages of A. persicus infesting laying hens were 99.2% and 63.4% after dipping in peracetic acid and deltamethrin, respectively. However, a complete elimination of the number of ticks occurred after 28 days PT with deltamethrin. Peracetic acid inhibited molting effectively (28%) when compared with that of deltamethrin (52%). Results indicated that peracetic acid is a more potent and promising acaricide against A. persicus (in vitro and in vivo) than deltamethrin. The acaricidal efficacy of cypermethrin, ivermectin, trichlorphon, and Azadirachta indica (Neem) was evaluated in commercial layers (N 5 500 of 40
50-week-old), naturally infested with A. persicus.23 The number of ticks was counted on all the birds before treatment and on TD 1, 7, 14, 21, and 28 PT. The best control was 87% at TD 21 and 28 PT on cypermethrin treated birds against A. persicus infestation followed by ivermectin (83%) on TD 14 PT, 40% A. indica (52%) on TD 14 PT, and trichlorphon (42%) on TD 14 PT.

Ornithodoros moubata
Murray, 1877; Ornithodoros turicata—Duge`s, 1876 Biology and life cycles Ornithodoros species are found on all continents except Antarctica and are endophilic, often being found in loose soil or litter in the proximity of their hosts. There are three general life cycle patterns for the Ornithodorinae,

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that is, (1) all stages (larvae, nymphs, adults) feed rapidly (15
30 min) and immature stages molt after feeding (e.g., O. turicata), (2) larvae feed for an extended period of several days and nymphs and adults are rapid feeders with immature stages molting after feeding (e.g., Ornithodoros puertoricensis), and (3) larvae do not feed but molt to the nymphal stage, which feeds rapidly as does the adult (e.g., O. moubata). A number of Ornithodoros species are efficient vectors of arboviruses, including African swine fever virus,24
27 and the relapsing fever spirochete Borrelia turicata. Adult soft ticks are more globular lacking a sharp margin and not appearing distinctly ovoid. Their body is flattened when unfed becoming convex dorsally when engorged. Ornithodoros spp. have multiple gonotrophic cycles and are extremely hardy under conditions of lack of food, some species surviving from 5 to 7 years without feeding.6 The eggs are protected from the environment by an impermeable waxy layer deposited on the surface of each egg by the Gene’s organ of the female tick. The eggs incubate and hatch within 7
30 days. Larvae have six legs and may either feed and molt to the nymphal stage or not feed and molt to the nymphal stage. The nymphs feed on hosts and undergo several stages before the finally molt to an adult tick.

Rearing method(s) Collecting methods Soft tick collection methods have been summarized by Perez de Leon et al.28 and include the following: 1. Direct mechanical methods of collecting Ornithodoros spp. include scraping suspected habitats with a shovel and passing collected material through several sieves of decreasing mesh size, then manually picking up individual ticks with a forceps. 2. Vacuum collection with a gas-powered mechanical vacuum device for tick collection from burrows based on a leaf blower that aspirated the contents of burrows, passed the collected material through a series of screens, which were then removed from the machine and emptied into a tray for handpicking of individual ticks with a forceps. 3. CO2 traps. Hokama and Howarth29 devised a CO2 trap system that included a container for placing dry ice slightly above a tray buried in the soil to a level that attracted ticks could walk over the edge of the tray and fall in, thereafter being unable to climb out. Dry ice that sublimates to CO2 proved an efficient attractant for Ornithodoros coriaceus. Adeyeye and Butler30 showed a CO2 trap to be effective for the collection of O. turicata from gopher tortoise burrows and a similar CO2 trap was shown to be effective for the collection of Ornithodoros erraticus in Portugal.31

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Feeding and cultivation methods The relatively short attachment and feeding duration of soft ticks facilitates the use of membranes for blood feeding (34 C
38 C) that include animalderived membranes from mice, rabbits, chickens, pigeons, bat wings, air sac membranes from eggs, Parafilm and silicon membranes.32,33 Nymphs and adults engorge on feeding membranes within 15 min. Heparinized or defibrinated bovine, ovine, pig, and rat blood have been used with various soft tick species. Techniques have been described for efficiently producing thousands of soft ticks for experimental use.34 A general method is described for each of the following feeding categories: (1) small species with short-term-feeding larvae (O. turicata, Ornithodoros parkeri, O. erraticus); (2) small species with long-term-feeding larvae (O. puertoricensis, Ornithodoros dugesi); and (3) large species with long-term-feeding larvae (O. coriaceus). Apparatus used for tick feeding and containment are described in detail, as are special conditions required by the Ornithodoros spp. listed. In vitro method(s) Contact—immersion—nymph A fluralaner stock solution (20 mg/mL) was diluted with DI water to a test concentration of 1000 µg/mL and a series of 1:10 dilutions with DI water were prepared to obtain fluralaner test concentrations between 1000 and 1024 µg/mL (i.e., 1000, 100, 10, 1, 0.1, 0.01, 0.001, and 1024 µg/mL). Twenty unfed O. moubata nymphs were immersed in 5 mL of either test or a vehicle solution in an Erlenmeyer flask for 5 min or nymphs remained untreated. Ticks were then strained, dried on a paper towel, transferred into a Petri dish lined with a dry filter paper, and covered. The Petri dish was incubated at 20 C and 95% RH for 48 h. Thereafter, the number of live and dead ticks was counted on a heated plate.35 Contact—topical—adult Studies were done to determine the susceptibility of fasting Ornithodoros tholozani (5O. papillipes) adult ticks to DDT and benzene hexachloride.36 Pesticide tests were carried out with unfed adults comprising three different physiological states, namely, (1) unfed for 3
4 months postmolt, (2) unfed for 12
13 months postmolt, and (3) fed several times postmolt for 5
6 years, the last feeding for 2 years before the tests. Treatments were made with a self-filling capillary applicator (WHO test kit). Topical applications were made with acetone solutions of DDT (p,p0 -isomer) and benzene hexachloride (gamma isomer) applied as a single 0.38 µL drop to each tick. Each treatment was applied to 10 ticks of a given physiological age group, comprising 1 replicate with 1
3 replicates per concentration of insecticide.

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Treated ticks were held in glass ampoules at 23 C
26 C and 60%
80% RH. The photoperiod was not controlled. Observations were made at intervals of 1
2 days initially, less frequently later through a 125-day period. The responses of the treated ticks to light and mechanical stimuli were observed and recorded, in accordance with previously reported criteria for determining insecticidal poisoning.36 Dead ticks were removed and the time of death estimated. The treated ticks exhibited a gradual and progressive development of toxic symptoms, with stages similar to those observed by Uspenskiy and Levikov37 in hard ticks. Often death was delayed until several months after the application of the acaricides, in some cases up to 125 days. The toxic syndrome was always fatal if ticks reached the fourth (immobile) stage of intoxication, although differences in acaricide concentration and the physiological state of ticks could affect the time of death. Older ticks were more susceptible to DDT than younger ticks. Reports of tick resistance to acaricides may be misleading unless adequate time is allowed following the treatment for the manifestation of the characteristically slow toxic response.36

Membrane feeding—blood—nymphs Defibrinated sheep blood was added to a fluralaner stock solution (50 mg/mL) to produce a 1000 µg/mL fluralaner preparation that was then further diluted in series of 1:10 with sheep blood to obtain test concentrations between 0.1 and 10210 µg/mL (i.e., 0.1, 0.01, 0.001, 1024, 1025, 1026, 1027, 1028, 1029, and 10210 µg/mL). Twenty O. moubata nymphs were fed using artificial membranes with either a test or vehicle control preparation or an untreated control (feeding on blood only) until engorgement. Ticks were counted and placed into a plastic vial that was then closed with a stretched membrane and placed upside down on a glass dish containing 2 mL warmed test or vehicle preparation to permit feeding. Fully engorged ticks were transferred to dry filter paper in a Petri dish and covered. Ticks were incubated at 22 C and 90% RH for 48 h, then the dishes were opened and numbers of live and dead ticks counted on a heated plate.35 In this assay, dissolving potential acaricidal agents in calf blood, O. parkeri feeds through the Parafilm membrane and can rapidly demonstrate systemic acaricidal activity by killing or paralyzing the ticks after the feeding event. Citrated calf blood (5 mL) is added into a Nunclon Petri dish (40 3 12 mm). Test compound (12.5 µL) is pipetted into the calf blood (200 g sodium citrate/L to each 500 mL evacuated container). The calf blood is drawn directly into the evacuated container and mix until the volume is fully drawn. The citrated blood is mix with the pipette. Parafilm is stretched over the Petri dish to create a membrane, and the contents shaken to fully mix the compound in the blood. The Petri dish is placed on a warm flat bed.

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A small piece of gauze is placed on the Parafilm and “dampened” with untreated blood to encourage feeding. Six nymphal stage ticks are collected (that have had two molts) and placed onto the membrane and covered with the lid of the Petri dish. The nymphs are allowed to feed until engorged. Ticks must not be left to feed for too long as they can overfeed. While the ticks are feeding, a No. 3 pot is filled a quarter to a third with sand for each compound being tested. Once the ticks are engorged, they are removed from the membrane, their mouthparts blotted dry with gauze, and placed into the labeled pot. Fed ticks are observed at 24, 48 and 72 h. To assess the compound activity the ticks are physically brought to the surface of the sand by shaking the container in a back and forth movement and left for a few hours before counting. Ticks that have buried into the sand are unaffected. Those ticks that have not buried are observed for paralysis or death. Paralysis is determined if the legs twitch when touched, for dead ticks no twitching of the legs is observed. Results are recorded as the number dead per six ticks and number paralyzed. A repellent/ expellant effect of some compounds can be observed when ticks refuse to feed at certain concentrations. This should be observed and noted. A positive control for the assay is fipronil. DMSO is used for the negative control. Quality control: No more than two dead ticks in the negative control pot.

Membrane feeding—blood—adult In this assay, potential acaricidal agents are dissolved in calf blood, and O. turicata are fed through the Parafilm membrane as described for the membrane feeding assay for nymphs. This assay can rapidly demonstrate systemic acaricidal activity by killing or paralyzing the ticks after the feeding event. Citrated calf blood (5 mL) in a Nunclon Petri dish plus 12.5 µL of test compound is mixed. Parafilm is stretched over the Petri dish to create a membrane and the contents shaken to fully mix the compound in the blood. The Petri dish is placed on a warm flat bed. A small piece of gauze is placed on the Parafilm and “dampened” with untreated blood to encourage feeding. Adult ticks (5) are placed onto the membrane and covered with the lid of the Petri dish. The adults are allowed to feed until engorged. Canine hair may be used to help stimulate the ticks to feed. Ticks must not be left to feed for too long as they can overfeed. Once the ticks are engorged, they are removed from the membrane, their mouthparts blotted dry with gauze, and place into the appropriately labeled No. 3 pot just as described for the nymphs. Fed ticks are observed at 24, 48, and 72 h. Ticks are physically brought to the surface of the sand and left for a few hours before counting. Ticks that have buried into the sand are unaffected. Those ticks that have not buried are observed for paralysis or death. Paralysis is determined if the legs twitch when touched, for dead ticks no twitching of the legs is observed. Results

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are recorded as the number dead per five ticks and number paralyzed. A repellent/expellant effect of some compounds can be observed when ticks refuse to feed at certain concentrations. This should be observed and noted. A positive control is fipronil and DMSO is used for the negative control. Quality control: No more than two dead ticks in the negative control pot. In the screening of isoxazolines, those compounds that demonstrated activity in the primary in vitro flea screens were then evaluated in an in vitro soft tick screen that used a system similar to that described earlier for testing compounds against the soft tick, O. turicata.38 Five individual nymphal thirdand fourth-stage O. turicata were placed on top of the membrane covering a 50 mm Petri dish containing blood with test compound, and ticks were allowed to feed to repletion (B5
15 min). Assays were conducted in duplicate for initial screening and in triplicate for comparative assays. Ticks were removed and placed in a separate container with sand and incubated for 72 h at 25 C and 75% RH. Ticks were observed at 24 and 72 h for paralysis/mortality. End point results for the subjective visual assessment of the organism viability were recorded as EC50 and LC100 in µg/mL.38 The susceptibility of males, females, and third nymphal instars of O. erraticus and O. moubata to fluralaner was determined through in vitro feeding exposure.39 Fluralaner activity against these developmental stages and species was assessed by feeding the ticks on ovine blood medicated with decreasing fluralaner concentrations between 1 and 1028 µg/mL. Tick mortality was measured at 4, 24, and 48 h and 1, 2, and 3 weeks postfeeding. Tests included solvent-treated and untreated blood controls. Fluralaner was extremely active against O. erraticus, with mean LC50 and LC95 of 2.0 3 1028 and 5.4 3 1028 µg/mL, respectively. Fluralaner was also highly active against O. moubata, showing a mean LC50 of 1.5 3 1026 µg/mL and a mean LC95 of 1.8 3 1023 µg/mL. In the latter species the most susceptible life stages were the females (LC95 1.4 3 1024 µg/mL). Fluralaner demonstrated potent acaricidal activity against all developmental stages of O. erraticus and O. moubata tested within the first 48 h after in vitro feeding.

Membrane feeding—blood—larvae An artificial membrane system was adapted to feed O. turicata larvae from a laboratory colony using defibrinated swine blood.40 Aspects related to larval feeding and molting to the first nymphal instar were evaluated. A total of 55.6% of all larvae exposed to the artificial membrane in two experimental groups fed to repletion and 98.0% of all fed larvae molted. Mortality rates of first-instar nymphs differed significantly depending on the sorting tools used to handle engorged larvae (χ2 5 35.578, P , .0001): engorged larvae handled with featherweight forceps showed significantly higher mortality (odds ratio 5 4.441) than those handled with a camel-hair brush. Differences in the physical properties of the forceps and camel-hair brush may affect the

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viability of fragile soft tick larvae even when care and the same technique are used to sort them during experimental manipulations. The current results represent those of the first study to quantify successful feeding to repletion, molting, and postmolting mortality rates in O. turicata larvae using an artificial membrane feeding system.

Repellency A method for evaluating tick repellents on guinea pigs has been described.41 Pyrethrum was outstanding and far superior to all the other chemicals tested in protecting from tick bites. Its effect was probably due mostly to a strong irritant effect on the ticks. Experiments were carried out to determine the effect of repellents against ticks in the laboratory and in an infested natural cave.42 The materials were applied to rats by spraying them with a solution of the repellent in ethanol. Pyrethrum was outstanding and far superior to the other repellents in preventing ticks of all stages from biting; however, it did not repel ticks from a distance. Ten commercial and 11 experimental repellents were tested on white mice against the soft tick O. parkeri, a vector of the causative agent of relapsing fever in the western United States.43 The most effective commercial repellent was permethrin (EC95 5 0.14%), followed by dimethyl phthalate (EC95 5 0.23%). The most effective experimental repellent was USDA AI3
35765 [l-(3-cyclohexen-l-ylcarbonyl) piperidine, EC95 5 0.75%]. In vivo method(s) Ivermectin was injected into hosts and evaluated for effectiveness against O. parkeri and the chicken mite Dermanyssus gallinae (DeGeer).44 O. parkeri second-stage nymphs showed a marked increase in mortality when fed on mice injected IP with ivermectin at a dose between 0.1 and 0.2 mg/kg bw. For adult O. parkeri and D. gallinae, 0.4 and 0.5 mg/kg bw, respectively, were necessary for an increase in mortality over controls. These latter findings are comparable to those reported for other tick species but differ from those reported for the northern fowl mite, Ornithonyssus sylviarum. Further testing narrowed the effective dose range for O. parkeri adults to between 0.425 and 0.450 mg/kg bw. The time interval (4, 8, and 24 h) between ivermectin injection of the host and tick feeding had only a slight influence on the overall effectiveness of the drug. In O. parkeri, doses of 0.0125
0.1000 mg/kg bw did not affect fecundity, hatchability, gross morphology of the reproductive system and synganglion, or histology of the reproductive system. Contrary to the reports of irreversibility of effects of ivermectin on GABA-mediated neurotransmission, many ivermectinparalyzed ticks recovered partial mobility over time.

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Field studies were conducted to assess the reliability of CO2 baits in sampling O. turicata.30 Tick response to B50 g of dry ice placed at various distances away from tick-infested burrows was monitored over a 2-h period. In addition, tick attraction to different flow rates of CO2 was monitored. Tests were conducted over a 15-month period, during which seasonal effects on O. turicata response to CO2 were assessed. The efficacy of the baits was evaluated at night and in daytime. Ticks responded to dry ice baits placed up to 8 months away from the burrows. There was no significant difference in the total number of ticks attracted in a 1-h period using 500
2000 mL CO2/min. At night, ticks were attracted to CO2 bits throughout the years except in December and January. By contrast, ticks were attracted to CO2 baits during daytime only between May and mid-December.

References 1. Burger TD, Shao R, LaBruna MB, Barker SC. Molecular phylogeny of soft ticks (Ixodida: Argasidae) inferred from mitochrondrial genome and nuclear rRNA sequences. Ticks Tick-Borne Dis 2014;5:195
207. 2. Hoogstraal H, Aeschlimann A. Tick-host specificity. Bull Soc Entomol Suisse 1982;55:5
32. 3. Hoogstraal H. Argasids and nuttalliellid ticks as parasites and vectors. Adv Parasitol 1985;24:135
238. 4. Mehlhorn H. Argas species, leather or soft ticks. Encyclopedia of parasitology. Springer-Verlag Berlin Heidelberg; 2015. p. 1
4. 5. Urquhart GM, Armour J, Duncan JL, Dunn AM, Jennings FW. Veterinary parasitology. 2nd ed. Blackwell Science Ltd; 1996. p. 188
9. 6. Bowman DD, Lynn RC, Eberhard ML, Alcaraz A. Georgis’ parasitology for veterinarians. 8th ed St, Louis, MO: Saunders; 2003. p. 49
51. 7. Bishopp FC, Trembley HL. Distribution and hosts of certain North American ticks. J Parasitol 1945;31:1
53. 8. Keirans JE, Pound JM. An annotated bibliography of the spinose ear tick, Otobius megnini (Duge`s, 1883) (Acari: Ixodida: Argasidae) 1883
2000. Syst Appl Acarol Special Pub, 13. 2003. p. 1
68. 9. Cooley RA, Kohls GM. The Argasidae of North America, Central America and Cuba, Monograph 1. Am Midl Nat. Notre Dame, IN: University of Notre Dame; 1944. p. 21
36. 10. Dives GC, Raiakaruna RS. Life cycle of Spinose ear tick, Otobius megnini (Acari: Argasidae) infesting the race horses in Nuwara Eliya, Sri Lanka. Acta Trop 2017;166:164
76. 11. Nava S, Mangold AJ, Guglielmone AA. Field and laboratory studies in a Neotropical population of the spinose ear tick, Otobius megnini. Med Vet Entomol 2009;23(1):1
5. 12. Wanchinga DM, Barker RW. Colonization and laboratory development of Otobius megnini (Acari: Argasidae). J Econ Entomol 1986;79(4):999
1002. 13. Niebuhr CN, Breeden JB, Lambert BD, Eyres AI, Haefele HJ, Kattes DH. Off-host collection methods of the Otobius megnini (Acari: Argasidae). J Med Entomol 2013;50 (5):994
8.

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14. Niebuhr CN, Mays SE, Breeden JB, Lambert BD, Kattes DH. Efficacy of chemical repellents against Otobius megnini (Acari: Argasidae) and three species of ixodid ticks. Exp Appl Acarol 2014;64(1):99
107. 15. Drummond RO, Whetstone TM, Ernst SE. Insecticidal control of the ear tick of cattle. J Econ Entomol 1967;60(4):1021
5. 16. Craig TM, Kunde JM. Controlled evaluation of ivermectin in Shetland ponies. Am J Vet Res 1981;42:1422
4. 17. Nava S, Gugliemone AA. Difficulties to control natural infestations with Otobius megnini (Acari: Argasidae) nymphs in cattle with systemic biocides. Res Vet Sci 2009;87(2):258
9. 18. Thangamani S. Chapter 2: Maintenance of Argasid ticks in the laboratory—SOP. In: Levin ML, Schumacher L, Thangamani S, editors. Laboratory colonies of ticks. BEI Resources; 2015. 19. Micks DW. The laboratory rearing of the common fowl tick, Argas persicus (Oken). J Parasitol 1951;37(1):102
5. 20. El-Kammah KM, Abdel Wahab KS. Argas (Persicargas) persicus life cycle under controlled and outdoor conditions. Acarologia 1980;21(2):163
72. ˇ 21. Dusba’bek F, RupesSˇ V, S7breve;imek P, ZahradnicSkova H. Enhancement of permethrin efficacy in acaricide-attractant mixtures for control of the fowl tick Argas persicus (Acari: Argasidae). Exp Appl Acarol 1997;21:293 Available from: https://doi.org/10.1023/ A:1018407307532. 22. Khaler HF, Seddiek SA, El-Shorbagy MM, Ali MM. Erratum to: the acaricidal efficacy of peracetic acid and deltamethrin against Argas persicus, infesting laying hens. Parasitol Res 2013;112(10):3669
78. 23. Khan LA, Khan MN, Iqgal Z, Qudoos A. Comparative acaricidal efficacy of cypermethrin, ivermectin, trichlorphon and Azadirachta indica (neem) in layers naturally infested with Argas persicus. Pak J Agric Sci 1965;38(3/4):29
31. 24. Plowright W, Perry CT, Peirce MA, Parker J. Experimental infection of the argasid tick, Ornithodoros moubata porcinus, with African swine fever virus. Arch Gesamte Virusforsch 1970;31:33
50. 25. Hess WR, Endris RG, Lousa A, Caiado JM, Mebus CA, Monahan MJ. Clearance of African swine fever virus from infected tick populations. J Med Entomol 1989;26 (4):314
17. 26. Endris RG, Haslett TM, Monahan MJ, Hess WR. Experimental transmission of African swine fever virus by the soft tick, Ornithodoros (Alectorobius) puertoricensis (Acari: Argasidae). J Med Entomol 1991;28:854
8. 27. Endris RG, Hess WR. Experimental transmission of African swine fever virus by the Iberian soft tick, Ornithodoros (Pavlovskyella) marocanus (Acari: Argasidae). J Med Entomol 1992;29:652
6. 28. Perez De Leon AA, Showler A, Kucheryavenko RO, Li AY, Kucheryavenko V, Filatov S, et al. Soft tick sampling and collection. J Vet Med Biotech Biosafety 2015;1(2):5
11. 29. Hokama Y, Howarth JA. Dry-ice (CO2) trap for efficient field collection of Ornithodoros coriaceus (Acari: Argasidae). J Med Entomol 1977;13(4-5):627
8. 30. Adeyeye OA, Butler JF. Field evaluation of carbon dioxide baits for sampling Ornithodoros turicata (Acari: Argasidae) in gopher tortoise burrows. J Med Entomol 1991;28(1):45
8. 31. Caiado JM, Melo MA, Boinas F, Louza AC. The use of carbon dioxide insect traps for the collection of Ornithodoros erraticus on African swine fever virus infected farms. Prev Vet Med 1990;81(1):55
9.

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32. Butler JF, Hess WR, Endris RG, Holsher KH. In vitro feeding of Ornithodoros ticks for rearing and assessment of disease transmission. In: Griffith DA, Brown CE, editors. Proc VI Int Cong Acarol, Acarology VI, vol. 2; 1984. 33. Sonenshine DE, Roe RM. Biology of ticks, vol. 2. New York: Oxford University Press; 2014. p. 463
4. 34. Endris RG, Haslett TM, Monahan MJ. Rutledge. Techniques for mass rearing soft ticks (Acari: Argasidae). J Med Entomol 1986;23(3):225
9. 35. Williams H, Zoller H, Roepke RK, Zschiesche E, Heckeroth AR. Fluralaner activity against life stages of ticks using Rhipicephalus sanguineus and Ornithodorus moubata in vitro contact and feeding assays. Parasit Vectors 2015;8:90 Available from: https://doi. org/10.1186/s13071-015-0704-x. 36. Uspenskiy IV. Susceptibility to acaricides of adult Ornithodoros tholozani (Ixodoidea: Argasidae) in relation to the slow-death syndrome. J Med Entomol 1982;19:701
5. 37. Uspenskiy IV, Levikov VB. Development of effects in ixodid ticks (Ixodes persulcatus, Dermacentor silvarum, Haemaphysalis concinna) after DDT application. [In Russian with English summary]. Med Parazitol Parazit Bolezni 1974;43:411
17. 38. 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;222:3
11. 39. Pe´rez-Sa´nchez R, Oleaga A. Acaricidal activity of fluralaner against Ornithodoros moubata and Ornithodoros erraticus argasid ticks evaluated through in vitro feedings. Vet Parasitol 2017;243:119
24. 40. Kim HJ, Filatov S, Lopez JE, Perez De Leon AA, Teel PD. Blood feeding of Ornithodoros turicata larvae using an artificial membrane system. Med Vet Entomol 2017;31(2):230
3. 41. Bar-Zeev M, Gothilf S. Laboratory evaluation of tick repellents. J Med Entomol 1973;10:71
4. 42. Bar-Zeev M, Gothilf S. Field evaluation of repellents against the tick Ornithodoros tholozani Labou. & Me´gn. in Israel. J Med Entomol 1974;11:389
92. 43. Mehr ZA, Rutledge LC, Morales EL, Inase JL. Laboratory evaluation of commercial and experimental repellents against Ornithodoros parkeri (Acari: Argasidae). J Med Entomol 1986;23:136
40. 44. Ash LS, Oliver Jr. JH. Susceptibility of Ornithodoros parkeri (Cooley) (Acari: Argasidae) and Dermanyssus gallinae (DeGeer) (Acari: Dermanyssidae) to ivermectin. J Med Entomol 1989;26(3):133
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Chapter 4b

Arachnida, Metastigmata, Ixodidae (except Ixodes holocyclus) Maxime Madder, PhD, Professor1,2, Josephus J. Fourie, PhD3 and Theo P.M. Schetters, PhD, Professor1,2 1 Clinglobal, La Mivoie, Mauritius, 2Department of Veterinary Tropical Diseases, University of Pretoria, Pretoria, South Africa, 3Clinvet International, Bloemfontein, South Africa

Arachnida Metastigmata Ixodidae

Biology and life cycles The family Ixodidae or hard ticks are all ectoparasites of wild and domestic vertebrates including man. They occur worldwide and are known to be vectors of diseases or cause of toxicosis.1 Of the B900 tick species, about 700 are hard ticks that can be found in virtually all terrestrial regions of the planet. Hard ticks can easily be distinguished from soft ticks by the presence of a dorsal anteriorly positioned sclerified scutum (in larvae, nymphs, and females) or conscutum that covers the entire dorsal side in the case of males, and the presence of anteriorly projecting mouthparts that are visible from the dorsal side. As in soft ticks, larvae are six legged, whereas nymphs and the adult stages are eight legged. Depending on the species of ticks, hard ticks can display three different types of life cycles: a one-host, two-host, or three-host life cycle. The different cycles refer to the behavior of the postengorgement stages and not to the number of hosts they feed on. A three-host tick might just feed on one-host; Rhipicephalus sanguineus, the dog tick, feeds just on a dog, but is still a threehost tick because of the behavior of the engorged stages. In one-host ticks, engorged larvae and nymphs remain on the host after engorgement and feed on that same host after the molt; in two-host ticks only engorged larvae stay on the host, and in three-host ticks all postengorgement stages drop off from the host and molt to the next stage in the vegetation or in the case of females oviposit. In hard ticks, all stages of ticks need a blood meal to complete their life cycle. In contrast to soft ticks, blood feeding takes days to weeks, and females only have one gonotropic cycle but may lay up to 20,000 eggs in one egg batch of some of the Amblyomma species. Depending on the species, the life cycle can be as short as 3 months for one-host ticks, such as

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Rhipicephalus microplus, in tropical conditions or extending up to 3
4 or even more years in temperate regions in the case of the prostriate Ixodes ricinus. Synchronization of the life cycle in order to avoid mortality during periods of adverse conditions, extremely low temperatures during the winter (hibernation) or dry and hot conditions during the summer (estivation), is often regulated by a form of dormancy, a state of minimal metabolic activity, scientifically called behavioral or developmental diapause or quiescence. Knowledge of the life cycle, host preference, and also the activity patterns of tick species is essential for the successful maintenance and reproduction of tick species under laboratory conditions.

Dermacentor variabilis—Say, 1821—American dog tick or wood tick Ticks of the genus Dermacentor are ornate ticks with medium-sized mouthparts, eyes, festoons, and bifid first pair of coxae, and male ticks have no adanal plates, but enlarged fourth pair of coxae. The distribution of Dermacentor variabilis is restricted to Eastern parts of the United States up to the Atlantic coast. The American dog tick is a three-host tick and the life cycle can take up to 2 years, depending on the environmental factors such as temperature and humidity. Synchronization of the life cycle with favorable conditions for development and survival is obtained by diapausing nymphs or larvae at the end of the season.2 The larvae of D. variabilis commonly feed on mammals, such as squirrels, mice, raccoons, skunks, and deer, but also parasitize humans and birds. After the molt, nymphs feed on the same kind of hosts as the larvae. Adults, on the other hand, feed on larger animals, such as cats, dogs, opossum, deer, and humans. D. variabilis is a known vector of Rocky Mountain spotted fever (Rickettsia rickettsiae), and tularemia (Francisella tularensis). Female ticks can also cause paralysis of their host after feeding for 4
7 days.

Dermacentor andersoni—Stiles, 1908—Rocky Mountain wood tick Dermacentor andersoni is known as the Rocky Mountain wood tick and is mainly restricted to the Rocky Mountain region in the northwestern United States and southwestern Canada. D. andersoni has a light silver-gray ornamentation on the scutum in the females and conscutum of the males. The scutum is heavily and deeply punctate compared to other species of the genus. Differentiation between the different Dermacentor species relies on the size and number of goblets on the spiracular plates. D. andersoni ticks have 100
200 moderately sized goblets, while D. variabilis ticks have more than 300 small goblets.

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Like all Dermacentor ticks, D. andersoni is a three-host tick and takes a single blood meal in each life cycle stage. The entire life cycle will normally take 1
3 years under natural conditions. Unfed stages can estivate for extended periods of time or can hibernate if unsuccessful in host finding. All stages feed on mammals, although adults prefer larger mammals, such as dogs, livestock, larger wildlife, and also humans. Larvae and nymphs are mainly attracted to mustelids, hares, rabbits, and rodents, such as mice, grounds squirrels, and voles. With regards to disease transmission, D. andersoni is generally a vector of Colorado tick fever and can also be a vector of Rocky Mountain spotted fever, bovine anaplasmosis, and tularemia.3

Rhipicephalus sanguineus—Latreille, 1806—brown dog tick, kennel tick or pan-tropical dog tick Members of the genus Rhipicephalus or brown ticks, with the exception of the recently added members of the previously known Boophilus species, are almost all three-host ticks with a brown appearance. Some of the species do have colorful ornamentation, but most are brown. Ticks of this genus are of medium size with medium-sized mouthparts. Their basis capituli is hexagonal, eyes and festoons are both present, and the first pair of coxae is bifid. Males have adanal and accessory anal plates. One of the most widespread ticks is R. sanguineus, also known as the brown dog tick, kennel tick, or pan-tropical dog tick. The tick has extended its distribution range together with its principal host, the domesticated dog. The identification of R. sanguineus sensu stricto is currently not possible, as biological, morphological, and molecular studies have highlighted a complex situation involving Rhipicephalus turanicus and several other cryptic species.4 R. sanguineus is like most brown ticks a medium-sized tick usually yellowish-brown to reddish-brown. The distinguishing feature between R. sanguineus and the closely related species R. turanicus females is that the latter has a narrow U-shaped genital aperture, but broad V-shaped in R. sanguineus. Both sexes of R. sanguineus have spirical plates with narrow tails, whereas in R. turanicus the tails are broad.5 The distribution of R. sanguineus is determined by the presence of the domestic dog and has spread between 50 N and 35 S. In more northern regions, this tick is often introduced by dogs with a travel history and can survive indoors causing heavy infestations in human and animal dwellings. This tick shows a three-host life cycle where larvae, nymphs, and female ticks detach after engorgement. But in contrast to most other species that molt or lay eggs on or just beneath the soil, all engorged instars of R. sanguineus climb up the wall and hide in cracks and crevices of the human or animal dwelling where they lay eggs or molt to the next instar. Depending on the temperature, more than one life cycle/year is possible.

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Although R. sanguineus is a three-host tick, it is monotropic and mainly feeds on dogs in all life cycle stages. Adult ticks and nymphs feed on the neck, head, and shoulders, although larvae feed on the stomach and flanks of dogs. Other hosts, such as cattle or humans, can be infested, but only if dogs are present to maintain the tick population. The main pathogens transmitted by R. sanguineus to dogs are the bacterium, Ehrlichia canis, causing canine monocytic ehrlichiosis and the protozoans, Babesia vogeli and Babesia gibsoni. Hepatozoon canis is not transmitted by the ticks during feeding, but only during grooming and ingestion of the entire infected tick. When feeding on human, R. sanguineus can transmit Rickettsia conori causing tick typhus.

Ixodes pacificus—Cooley & Kohls, 1943—Western blacklegged tick Ticks of the genus Ixodes also known as prostriate ticks are characterized by an anal groove surrounding the anus anteriorly, in contrast to all other genera where the groove is posteriorly positioned to the anus or not present at all. Mouthparts of all stages of Ixodes species are long, eyes and festoons are lacking, and males have no anal plates. In contrast, males have seven ventrally positioned sclerified body plates. Ixodes pacificus or the Western blacklegged tick is best known as the main vector of Lyme disease in western North America. Being a prostriate tick, it is easily recognized by an anal groove anteriorly surrounding the anus. The ticks has no cornua, auriculae are present but small, and the scutum has uniformly small punctuations. Coxa 1 has a moderately long internal spur, partly overlapping coxa 2, and all coxae have short external spurs.6 Its main distribution range covers the entire west coast of North America from the Canadian province of British Columbia in the North to Mexico in the South.7 This tick has been found at altitudes ranging from sea level up to 2150 m.8 The primary host of the adults is the black-tailed deer and is responsible for keeping high tick densities. Other hosts include domestic animals, such as cattle, horses, goats, dogs, and cats, and a whole series of wildlife including bear, several deer species, rats, mice, squirrels, badgers, and foxes. Immature stages feed on similar hosts, excluding larger mammals, and also on a variety of bird species and lizards. All stages readily feed on humans.7 I. pacificus is the primary tick that feeds on humans and also the primary vector of Borrelia burgdorferi, the causative agent of Lyme disease. It also transmits Anaplasma phagocytophilum, the etiological agent of human granulocytic ehrlichiosis, and equine ehrlichiosis.9 I. pacificus has also been the cause of tick paralysis in dogs.10

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Ixodes scapularis—Say, 1821—blacklegged tick, deer tick Ixodes scapularis, blacklegged tick or deer tick is the main vector of B. burgdorferi, the agent of Lyme borreliosis, but only distributed in the eastern and northern Midwestern United States and southeastern parts of Canada.11 Previously Ixodes dammini was also considered a valid species, but now defined as junior subjective synonym of I. scapularis, based on Article 23 of the International Code of Zoological Nomenclature.12 Morphologically, I. scapularis is very similar to I. pacificus, but the former species has a small but distinct cornua, an almost circular scutum in the females, and punctuations that are larger peripherally. The spiracular plate in males is elongate, whereas in I. pacificus it is oval.6 The published literature shows that white-tailed deer, cattle, dogs, and other medium-to-large-sized mammals are important hosts for adults as are native mice and other small mammals, certain ground-frequenting birds, skinks, and glass lizards for nymphs and larvae.13 I. scapularis is a typical three-host tick and has a 2
4 years life cycle. They have the ability to overwinter both in unfed and engorged states and are similar in this respect to the Palearctic relative I. ricinus.11 The greatest risk of being bitten by nymphs and female ticks exists during spring, summer, and fall. Adults may be questing for a host during winter as well when no frost is observed.14 Stages most likely to bite humans are nymphs and adult females. In addition to Lyme disease caused by B. burgdorferi and also Borrelia mayonii, I. scapularis is also a primary vector of Borrelia microti, the agent of human and rodent babesiosis, A. phagocytophilum, the etiological agent of human granulocytic ehrlichiosis, and Powassan disease. Moreover, I. scapularis often reaches extremely high infestations on livestock, and females can cause tick paralysis in dogs.13

Ixodes ricinus—Linnaeus, 1758—sheep, castor bean, deer, or wood ticks The species I. ricinus is also known as the sheep tick, castor bean tick, deer tick, or wood tick. The distribution range is mainly determined by the climatic conditions rather than tick
host interactions.15 This tick is a generalist and adapted to a wide variety of environments in Alpine, Boreal, Nemoral, Continental, Atlantic, Lusitanian, and Mediterranean domains.16 It is the most common tick found in central, western, and northern Europe,17 and in the dry areas of the Mediterranean region in Europe and North Africa, this species was renamed Ixodes inopinatus.18 I. inopinatus is allopatric with I. ricinus in Spain and Portugal.

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I. ricinus is a three-host tick and all stages climb the vegetation to quest for a host. The activity of adult ticks starts in early spring and can last until late autumn when temperatures become too low. During warm summer mo, adult ticks will not be found questing as mortality would be too high. The larvae are found questing from April onward followed by the nymphs. The sheep tick is a generalist with telotropic behavior and has been collected from a variety of hosts. Immatures feed on small and larger mammals, birds, and lizards, whereas adult ticks feed on large ruminants (sheep, cattle, and deer) or dogs and wild carnivores.5,19 I. ricinus is a competent vector of a variety of pathogens with zoonotic potential including bacteria, viruses, and protozoans. The most important diseases caused by these pathogens are borreliosis, anaplasmosis (A. phagocytophilum), and rickettsiosis (Rickettsia helvetica). I. ricinus also transmits Babesia divergens, that is, the causative agent of babesiosis in cattle and a life-threatening infection in splenectomized humans, tularemia due to F. tularensis, tick-borne encephalitis, and the Louping ill virus.17

Ixodes hexagonus—Leach, 1815—hedgehog tick Ixodes hexagonus is better known as the Hedgehog tick. Morphologically, I. hexagonus and I. ricinus look very similar, but they differ in many aspects.5 Apart from the hexagonal scutum in the females, the scapular grooves and fields are absent in I. hexagonus, auriculae are indistinct, and the internal spur on coxa 1 is short, whereas in I. ricinus it is long. Furthermore, the tarsi of the first pair of legs are bluntly stepped toward the claws. I. hexagonus is distributed over western, central, and southern Europe and has been recorded from Ireland, United Kingdom, Norway, Sweden, into the northwest up to Poland and Ukraine in the east,20 and Italy, Spain, Portugal, and Greece in the south.17 I. hexagonus is a three-host tick and in contrast to I. ricinus which is an exophilic tick that can be captured by dragging or flagging, I. hexagonus shows an endophilic behavior. All life cycle stages spend their life inside burrows or nests of their hosts and are therefore difficult to collect. Female ticks are normally found on hosts from spring to autumn with a peak in April
May.5 The principal hosts are medium-sized carnivores (dogs, cats, foxes, and mustelids) and hedgehogs, all animals with a permanent dwelling. Sporadically, it can be found on birds, sheep, cattle, and horses21 and can also bite humans. I. hexagonus may be found on dogs in large numbers and cause biting stress. It is also a competent vector of B. burgdorferi, tick-borne encephalitis virus, and Babesia microti, hence it’s importance in infesting humans.

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Amblyomma americanum—Linnaeus, 1758—lone star tick Species belonging to the genus Amblyomma are one of the largest ticks in the world. The most overt characteristic for all species is the extremely large mouthparts and the ornate scutum, both in male and female specimens. They also have banded legs, flat or beady eyes, and all stages have festoons. Males do not show any anal plates. Amblyomma americanum, also known as the lone star tick, is characterized by a silver-white spot at the posterior end of the scutum in female ticks and varied white streaks or spots around the margin of the top of the body in the males. It is a three-host tick present in the eastern and southeastern United States, but it is expanding its range northward and westward. This tick is also known to be very aggressive such that it also bites humans in the nymphal and adult stages.22 Just like Hyalomma ticks, adults and also nymphs of this genus are hunters and actively run toward their hosts. The adult stages primarily feed on large- or medium-sized mammals, but also on rodents and wild turkeys.23,24 Larvae are found feeding on birds and mammals, but not on small rodents, while nymphs feed on all animals as larvae and adults.23 The lone star tick is an important vector of zoonotic pathogens affecting humans in the United States. Ehrlichia chaffeensis, Ehrlichia ewingii, Coxiella burnetii, F. tularensis, Rickettsia amblyommii, and Borrelia lonestari have been isolated or identified from A. americanum and are capable of causing human disease.25

Amblyomma maculatum—Koch, 1844—Gulf Coast tick Amblyomma maculatum, known as the Gulf Coast tick, is also an ornate tick, and males of this species may look similar to A. americanum, whereas female ticks may be confused with female American Dog ticks (D. variabilis). The silvery ornamentation on male ticks is restricted to six disassociated, symmetrical markings located on the lateral edges of the scutum, whereas, the Gulf Coast tick patterning is interconnected and covers most of the tick’s dorsum. Dermacentor ticks can be differentiated from Amblyomma ticks by looking at the mouthparts. Mouthparts of Dermacentor species are short and rectangular in relation to the long and slender mouthparts of the Amblyomma ticks. These ticks are found in grass prairies and coastal uplands throughout much of the Western Hemisphere. Originally, they occurred in the coastal areas of the southern United States, but they are now also found more inland in Kansas, Oklahoma, and some other states. It is also found in parts of several Central and South American countries that border the Gulf of Mexico and Caribbean Sea, including Mexico, Guatemala, Belize, Nicaragua, Honduras, Costa Rica, Colombia, Venezuela, and some parts of Ecuador and Peru.26

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A. maculatum is a three-host tick as engorged larvae, nymphs, and females drop from the host after engorgement. They have a ditropic behavior as larvae and nymphs feed on small animals, such as birds, rodents, and rabbits, while adults feed on larger animals including dogs, coyotes, skunks, panthers, and bears. Gulf Coast ticks favorably infest the ears of host animals. The peak activities of A. maculatum depend on the geographic origin of the ticks. Coastal populations are mainly active from May through March, whereas continental populations are active from February through October.27 Seasonal peaks in Gulf Coast tick populations vary based on their geographic location.27 Coastal populations found in Texas are active from May through March, whereas inland populations in Oklahoma and Kansas are active from February through October.27 In Texas, adult activity peaks in August, followed by larvae in December, and nymphs in January. In Oklahoma and Kansas, adult activity peaks in April, followed by larvae in June, and nymphs in July. Gulf Coast ticks in Mississippi have been collected from March through November, with adults peaking during late July to early August.28 Seasonal activity of Gulf Coast ticks in northern Florida occurs from February to September.29 A. maculatum ticks are xerophilic and are able to quest during long periods in high temperatures. Apart from severe inflammation, abscesses, lethargy, debilitation, paralysis, and altered body composition of infested animals, the Gulf Coast tick transmits a series of pathogens including Rickettsia rickettsii, the causative agent of Rocky Mountain spotted fever, and Rickettsia parkeri. Hepatozoon americanum is not transmitted by injection, but by ingestion of the tick during grooming. Ticks also transmit Leptospira pomona and Ehrlichia ruminantium, the causative agent of heartwater, which is present in sub-Sahara Africa and also in the Caribbean.

Rhipicephalus microplus—Canestrini, 1888—Asian blue tick, cattle tick or southern cattle tick R. microplus is known in many parts of the world as the Asian blue tick, the cattle tick or the southern cattle tick, the latter specifically in the New World. Although now classified into the genus Rhipicephalus, the blue ticks where previously known under the separate genus Boophilus. But molecular evidence has demonstrated a monophyletic origin of the genus that arose within Rhipicephalus.30 Basically only the name changed, not the identification. R. microplus from Australia on the other hand changed to Rhipicephalus australis, and this not because of phylogenetic results, but morphological identification.31 It was Hoogstraal32 that postulated that R. microplus was introduced into East and Southern Africa together with Rhipicephalus annulatus from Asia

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via Madagascar with cattle importations after the rinderpest epidemic in the late 19th century. The tick established itself in East and Southern Africa, but more recently it was introduced into West Africa by cattle importation from Brazil.33 The ticks are also present in South America and Asia. This tick can be considered one of the most important tick species in the world as cattle infestations with R. microplus economically impact the cattle industry significantly by reducing weight gain and milk production and by transmitting pathogens that cause babesiosis (Babesia bovis and Babesia bigemina) and anaplasmosis (Anaplasma marginale).34 R. microplus has a one-host life cycle, which in total takes about 2 months to complete. Several generations are possible, especially in those areas with a prolonged rainy season. Because of its one-host life cycle, all stages of development occur on the host at the same time and the presence on a single host of B70 engorged females can represent a total parasitic population exceeding 10,000 ticks. R. microplus may be present in variable numbers throughout the year. Large numbers of larvae are usually present on pastures in late spring, and successive waves of larvae then occur through the summer and into the cooler autumn and early winter mo. Domestic cattle are probably the only effective hosts of this tick, although several collections, including engorged females have been made from goats. There are very few records from wildlife and then only when domestic cattle were also present. The tick transmits bovine babesiosis (B. bovis and B. bigemina). B. bovis infection is acquired by the adults of one generation of ticks and transmitted transovarially by the larvae of the next generation. Because this tick transmits both B. bovis and B. bigemina it poses a greater potential threat to livestock production than Rhipicephalus decoloratus. Bovine anaplasmosis (A. marginale) and spirochaetosis (Borrelia theileri) are also transmitted by R. microplus.

Rhipicephalus annulatus—Say, 1821—Texas fever tick or cattle fever tick R. annulatus or in the United States known as the Texas fever tick or cattle fever (babesiosis) tick is also a typical one-host tick previously belonging to the genus Boophilus and has similar characteristics as R. microplus. Morphologically, females are very difficult to distinguish from R. microplus and in many areas in West Africa where R. microplus was introduced from Brazil, R. microplus was for many years identified as R. annulatus. Distinguishing morphological features for both females and males is the absence of clear external spurs on coxae 2 and 3, except males only lack a caudal process, which is far easier to identify.

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The distribution of R. annulatus is very similar to that of R. microplus, but in Africa its range extent toward North Africa and also the European Mediterranean area and south of the Sahara eastward up to southeast Sudan. In the United States, this tick has been eradicated but still occurs in Mexico35 and can occasionally be found in Texas and California in a buffer quarantine zone along the Mexican border. The life cycle of R. annulatus is very similar to that of R. microplus, being a one-host tick only larvae have to quest for a host. The parasitic phase is B3 weeks where after the engorged females will detach and drop to the ground. Oviposition and eclosion of the eggs will take in ,2 mo, depending on the prevailing temperature. Under optimal conditions of high temperature and humidity, five generations can be completed in 1 year. Activity in North Africa begins in late summer (September) up to January with a peak in October.36 R. annulatus prefers to feed on cattle, but occasional hosts as sheep, goats, and wild ungulates are recorded if cattle are present in the area. In the absence of cattle, this tick can normally not maintain itself in the area. The predilection sites of R. annulatus are legs, belly, neck, and dewlap in the case of heavy infestations the back and shoulders are also infested.5 The main parasites transmitted by this tick are B. bovis and B. bigemina, both causing bovine babesiosis or redwater in cattle. Second, A. marginale responsible for causing bovine anaplasmosis or gall sickness is also transmitted. Heavy infestations can lead to production losses and could also cause damage to hides.

Haemaphysalis longicornis—Neumann, 1901—cattle tick or bush or shrub ticks The genus Haemaphysalis is characterized by small ticks with short mouthparts. Some species show lateral extending posterolateral angles on the second segment of the palps, giving the palps a triangular appearance. They can easily be distinguished from the other genera harboring small tick species by the following characteristics: absence of eyes and presence of festoons in all stages, rectangular basis capituli, and absence of adanal plates on the males. Only few species infest domestic livestock and are economically important. Haemaphysalis longicornis is also known as the cattle tick, shrub tick, or bush tick and its distribution ranges from temperate regions of East and Central Asia including China, Korea, and Japan and was introduced into Australia and New Zealand at the end of the 19th and beginning of 20th century.35,37 This tick is a typical three-host tick with larvae, nymphs, and adults questing for a host, feeding to repletion, detaching, and developing in the vegetation to the next stage. However, it is an unusual tick being represented by various genetic races: bisexual diploid, aneuploidy parthenogenetic

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(thelytokous), and obligate-parthenogenetic triploid.37 In the southern hemisphere, only the triploid races occur, whereas in Japan diploid and triploid races are found. The aneuploidy race is found on an island off Korea. Parthenogenetic races normally do not produce any male ticks. Known hosts of adult and nymphs of H. longicornis are domestic cattle, horses, pigs, sheep, goats, cats and dogs, and wildlife hosts include several deer species, badgers, wild cats, foxes, raccoons, brown hares, and rabbits. Larvae are primarily found on birds and smaller mammals.38 H. longicornis is a competent vector of Theileria orientalis causing bovine theileriosis in dairy and beef cattle, and It has also been associated with Theileria sergenti and Theileria buffeli.39 B. gibsoni is primarily transmitted by H. longicornis and Haemaphysalis bispinosa to dogs, but direct transmission from dog to dog has been observed in fighting breeds.40 Also Babesia ovata, Babesia major, as well as Babesia equi can be transmitted by this tick.39 Furthermore, the flavivirus causing Russian spring
summer encephalitis and Powassan encephalitis virus have been isolated from H. longicornis.41,42

Haemaphysalis leachi—Audouin, 1826—yellow dog tick Haemaphysalis leachi is also known as the yellow dog tick. Together with R. sanguineus, the brown dog tick, this tick parasitizes domestic dogs and are the main ticks found on dogs in the Afrotropical region. The H. leachi is part of the H. leachi group, and many stages have been misdetermined in the literature.4 In southern Africa (including all of South Africa, Zimbabwe, and southern Mozambique) Haemaphysalis elliptica, the southern African yellow dog tick, has been misdetermined for years as H. leachi. H. leachi is a three-host tick and the females feed for about 1
2 weeks. As most of the hard ticks, the body expands slowly for most of the feeding time, but engorges rapidly on the last day after copulation. After detaching and dropping to the ground, female ticks start laying 5000 eggs within 7
28 days. The eggs hatch in 2
9 weeks, depending on the temperature. Engorged larvae molt within 2
26 weeks and the nymphs within 2
7 weeks. H. leachi adults are active throughout the year. The larvae and nymphs of this tick usually infest common murid rodents but may also be found on dogs. Adults prefer dogs or wild carnivore, such as cats, foxes, jackals, and wild dogs. Cattle are also recorded as hosts, but probably due to the close association between domestic dogs and cattle. Adult ticks attach to the anterior part of the body, mainly head, neck, and shoulders. Being a typical dog tick, H. leachi transmits the protozoan Babesia rossi causing canine babesiosis, both transstadially as transovarially. The tick is also a competent vector of R. conori to humans, causing tick typhus.36

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Hyalomma marginatum—Koch, 1844—Mediterranean Hyalomma tick Members of the genus Hyalomma are all characterized as being large ticks with a fairly long hypostome, beady eyes, banded legs, and in male ticks the presence of three pair of anal plates. The species Hyalomma marginatum is also known as the Mediterranean Hyalomma tick and is one of the most important vectors of Crimean
Congo hemorrhagic fever in humans in North Africa. The identification of Hyalomma species is far from easy and distinguishing characteristics for H. marginatum from Hyalomma rufipes is the lack of dense setae around the spiracles and relatively smooth scutum compared to a heavily punctated scutum in H. rufipes. As indicated by its common name, H. marginatum occurs in the Mediterranean both in Europa and North Africa and steppe climates further eastward, but mainly in humid climates. It has been recorded in Morocco, Tunisia, Algeria, and on the European side in Portugal, Spain, France, Italy, Greece, Albania, former Yugoslavia, and up to India.5,36 Most of the Hyalomma ticks have a two-host cycle where only larvae and adults need to find a host. Engorged larvae remain on the host and molt directly into nymphs that easily attach and drop off the host after completion of the blood meal. Adult Hyalomma ticks show a hunter strategy compared to the questing strategy in most of the tick genera except for Amblyomma species. Ticks actively run toward a suitable host, whereas questing ticks await a passing host. The adults become active in March and have a peak activity in April
May and stay active until November. Larvae are active in summer months (May
September). Adults of this species feed on cattle, horses, sheep, goats and camels, whereas preferred hosts for the immature stages are small mammals, such as rabbits and hares, hedgehogs, and also births. Since immature Hyalomma ticks feed on nonmigratory as well as migratory birds, Hyalomma ticks including pathogens they might harbor are often introduced into areas along the migration routes of their hosts.43 Like most of the Hyalomma species the predilection sites for attachment of H. marginatum are the perineum, genital area, and the udder. Apart from Crimean
Congo hemorrhagic fever, H. marginatum transmits Babesia caballi to horses and under laboratory conditions can also transmit Theileria annulata that is normally transmitted by Hyalomma scupense in the same area.

Hyalomma scupense—Schultze, 1919 H. scupense was formally known as Hyalomma detritum, but Guglielmone et al.44 proposed the name to be annotated “H. scupense (5H. detritum).”

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It is a domestic endophilic tick of cattle and is of great economic importance to cattle farmers. In comparison to most other Hyalomma ticks, H. scupense is a relatively small species lacking pronounced and heavy punctuations. White bands on the legs, a genus-distinguishing feature for Hyalomma and Amblyomma species, are lacking. H. scupense is one of the most widespread Hyalomma species in the world. It has been collected in the Palearctic and Oriental zoogeographic region in several countries in northern Africa, southeastern European countries up to Russia and many Asiatic countries.45 It is mainly present in humid to arid regions and as many other Hyalomma species, H. scupense has a two-host life cycle. Adults show a hunter strategy. Because of its domestic endophilic behavior, this tick has a similar behavior as the three-host tick R. sanguineus. Engorged nymphs and adults climb up the walls in stables and hide in cracks and crevices where nymphs molt to adults or were female ticks oviposit. Larvae quest in autumn and engorged nymphs will enter diapause for several months before molting to adults. Adults become active in the spring
summer. Ticks mainly feed on cattle, horses, and donkeys but are also found on dromedaries and occasionally humans.45 The tick is the major vector of Theileria annulata causing tropical theileriosis in cattle, and this disease is often present on small farms with barns and stables. This tick also transmits Theileria equi to horses and donkeys causing equine piroplasmosis. Q fever caused by the bacterium C. burnetii is also transmitted by this tick to livestock and humans.5

Rearing method(s) The rearing methods of hard ticks are found in the Manual for maintenance of multihost ticks in the laboratory by Levin and Schumacher.46, Appendix B

In vitro method(s) Evaluation of antitick compounds Introduction In the light of the continuous emerging resistance against antitick treatments, an active screening and selection process for new products is required for sustained control of ticks and tick-borne diseases. Presently, tick infestations are controlled by a combination of farm management practices, treatment of animals with chemicals, and/or vaccination (R. microplus). Each of these methods has its specific effect on tick populations on the animals and in the environment. Research has been directed toward the development of assays that allow measuring the effect of the different intervention practices.

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Antitick products Different products for the control of tick infestation have been developed. These can be divided into chemical products and immunological products. The chemical products (either directly or indirectly through the activity of the parent molecule or derived metabolite) interfere with the physiology and development of the tick, which reduces viability and may lead to death. The immunological products are biological molecules, mostly proteins, that are used as antigens in vaccine formulations. The antigens themselves do not affect the ticks but induce immune responses that counteract tick infestation. Chemical products—acaricides A thorough understanding of the biology of the tick led to the development of products that interfered at different phases of the tick life cycle. When adult female ticks have completed a blood meal, they produce a high number of eggs that are deposited in the environment after the ticks have dropped off the infested host. Spraying of tick-infested areas with an acaricide is used to reduce tick infestation and to create a barrier for tick-free areas. In traditional farming practices in Africa, certain shrubs and plants that are avoided by ticks are planted around pastures where cattle are being kept. Extracts of some of these plant parts are traditionally used as insect repellents on humans and livestock. Fractionation of such plant extracts has led to the identification and purification of an array of molecules with antitick activity. Some compounds were shown to kill ticks by interfering with normal tick physiology (e.g., blocking of acetylcholinesterase). This group of compounds (acaricides) is mainly used to protect animals from tick infestation and is usually administered by plunge dip, pour-on, drop-on, or spray. Antitick compounds display different modes of action varying from tick repellence and reduction of fertility to killing of ticks. In vivo infestation experiments using suitable animal host species have been used to determine the protective activity of antitick compounds. A major advantage is that such experiments allow determining the combined effect of each of the antitick activities. Although these experiments are perfectly suited for testing final product formulations, they are less appropriate for the screening of new compounds with possible antitick activity. Preferably, an in vitro system is used that allows HTS of entire libraries of putative compounds. Such systems have their limitations, however, since they usually measure only one property of active compounds. Tick vaccines More recently, vaccination against ticks is being used to protect animals against tick infestation (reviewed in Schetters47). The composition of the different vaccines that are available is highly variable ranging from crude

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extracts from dead R. microplus larvae (TickVac, Technochimica) to relatively pure recombinant proteins, such as recombinant Bm86 protein, produced by Pichia pastoris (Gavac, Heber Biotech). Although the targets of protective immune reactions underlying the efficacy of these vaccines have not been elucidated, it is likely that these are as variable as the vaccine compositions. For instance, an experimental vaccine against R. microplus based on subolesin (a homolog of akirin, which is involved in intracellular signal transduction in arthropods), was shown to more specifically affect oocyte production. As a result, the protective effect was mainly reflected in a reduction of viable progeny. In contrast, vaccination against Bm86, which is a molecule that is located at the surface of the cells lining the midgut of R. microplus, affects the integrity of the midgut. This ultimately leads to death. Similar to chemical compounds, the protective activity of tick vaccines can be measured by the infestation of susceptible hosts. The main efficacy parameters are the reduction in engorged ticks and the size and viability of the progeny. The latter is usually determined after feeding by further incubation of engorged females in vitro to allow oviposition, maturation, and hatching. The screening of candidate vaccine antigens by immunization of susceptible hosts followed by infestation (either whole body or local infestations), as practiced to date, hampers progress in this research area. Especially now that new (combinations of) antigens need to be evaluated, an in vitro efficacy assay would be advantageous. Although the contribution of cellular immune reactions to vaccine-induced tick immunity cannot be excluded, antibodies against tick antigens play a major role in the expression of immunity. This opens the possibility to evaluate the combinations of specific antisera and monoclonal antibodies directed against tick antigens.48 In vitro hard tick assays Historically, compounds against insect and other arthropods were developed to protect crops from pests. Some of these pesticides were further developed to protect animals and humans. Products were formulated such that they could be sprayed on the plants, hence, they were applied on the outside of the arthropods. For screening purposes, assays were developed that determined the effect of exogenously applied compounds as opposed to assays in which compounds were actively taken up by the arthropods (feeding). With the development of tick vaccines, in vitro feeding assays have been developed to study the effect of antitick antibodies that are taken up with the blood meal. Feeding assays could also be used to study the effect of acaricides that are administered systemically and are taken up by ticks during feeding (for instance the isoxazolines). In order to determine the repellent activity of putative antitick compounds, some in vitro assays were developed in which the behavior of tick is measured when they approach an area with a putative repellent. It is of

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importance to realize that in these assays, ticks are not brought into physical contact with the antitick compounds, but that the ticks sense the compound with their chemosensory systems.

Biological assays All of the biological assays are difficult to standardize because of biological variability, such as the origin, species, strain and age of the ticks. Because of the complicated life cycle of ticks, in vitro assays using live ticks are lengthy and may take 6 weeks to complete. This is mainly due to the fact that it is preferred to work with relatively young larvae that first need to be hatched in the laboratory. External application of test compounds Historically, most of the insecticides used to protect crops were administered by direct spray. This is reflected in the application of some of the acaricides that were developed to protect cattle from tick infestation, for example, the use of a nap sack sprayer or the spray race. In the latter system, cattle are routed through a multihead shower that sprays the acaricide onto the animals when they pass. Alternatively, animals are routed through a bath with acaricide solution (dipping). The early in vitro assays that were developed relied on the immersion of ticks in a test solution.49,50 Later, variations of this assay were developed by different research groups. In order to come to some harmonization the FAO is advocating the larval packet test (LPT) for the screening for acaricide resistance (FAO 2004).51 Larval ticks Larval packet test In the LPT, larvae are wrapped in filter paper that has been impregnated with test compound as originally described by Stone and Haydock (1962).49 After incubation the viability of the larvae is assessed. For the preparation of the filter paper, serial dilutions of test compounds are made in an appropriate carrier solution. Starting concentrations vary with the acaricide that is tested. As an example, Lovis et al.52 used fourfold serial dilutions of test compounds ranging from 53 to 0.05 mg/m2. A volume of 0.7 mL of each dilution was then applied onto a 7.5
10 cm filter paper (Whatman No. 1). The impregnated paper is folded to create a packet for incubation of larvae. Larvae are hatched in the laboratory and used at 7
21 days old. Larvae (B100) are inserted in a packet. The packet is sealed and subsequently incubated for 24 h at 28 C 6 1 C and 70%
80% RH. After incubation the proportion of larvae that is not able to walk is determined and counted as being dead. Tests with over 10% mortality in the controls are rejected.

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Larval immersion test Instead of incubating larvae with impregnated filter paper, larvae can be submerged in acaricide solutions to study the effect of acaricides on viability (Shaw’s LIT, Shaw et al.53). This assay is more generally referred to as the larval immersion test (LIT). Immersion solutions are prepared in an appropriate diluent. Starting concentrations vary depending on the test compound. As an example, ivermectin was tested in twofold dilutions in distilled water with 1% (v/v) ethanol and 0.02% (v/v) Triton X-100, starting at a concentration of 0.01% (w/v).54 Next, larvae are incubated in the test solution for B10 min, and subsequently air-dried on filter paper. As in the LPT, 100 larvae are inserted in a packet and incubated for 24 h at 28 C 6 1 C and 70%
80% RH. After incubation the proportion of larvae unable to walk is determined and counted as being dead. A variant of this assay called the larval immersion microassay was developed by White et al.55 Larval tarsal test The manipulation of live larvae makes the LPT and LIT tedious assays. Lovis et al.52 have developed an assay in which eggs are allowed to hatch within the test system. In addition, microtiter plates are used instead of test tubes, which allows for easier and faster screening of test articles. Standard, flat-bottom 96-well microtiter plates are pretreated with serial dilutions of test compounds to obtain concentrations of 100
0.05 mg/ m2. After drying of the plates, 50 eggs are added to each well and incubated (uncovered) for 1
3 days at B95% RH and 28 C 6 1 C. Next, plates are sealed and further incubated at 28 C 6 1 C and 70%
80% RH. Two weeks after egg hatching, larval mortality is determined by the examination of each well using a dissecting microscope. Hard tick larval contact (glass vials) test—Rhipicephalus sanguineus The objective of the assay is to determine the contact activity of compounds against the brown dog tick larvae, R. sanguineus.56 The test can also be conducted with other hard tick species (D. variabilis, A. americanum, I. scapularis, etc.). The sensitivity of the test may differ between different species of ticks. Test compounds are dissolved in IPA and aliquots (500 µL) added to 4 dram glass vials (inner surface area 34.5 cm2, 345 µg/tube/500 µL 5 690 µg/ mL for 10 µg/cm2) placed on a roller for at least 2 h to allow the IPA to evaporate. IPA alone is used as a negative control and fipronil is used as a positive control. A vial containing tick larvae is opened carefully within an area B8 3 10 in. that has been taped off with duct tape. The tape is carefully rolled so that the sticky side is facing outward. The tape is then applied to the bottom of a disposal spill tray. Any tick larvae that escape from the vial without being captured will get trapped in the tape. Tick larvae (B50
200)

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are added to the vials using a swab and the vials are closed with silk screened ventilated caps. At B24 and 48 h the vials are examined and knockdown [Y 5 active, N 5 not active; knockdown (Y) can infer paralysis or death at 48 h] is recorded as active. Vials showing knockdown are examined for tick paralysis and/or death at B48 h. End point data are recorded as an effective dose 100% (ED100), and a lethal dose 100% (MLD100) in µg/cm2. Adult ticks Adult immersion test Instead of larvae, adult ticks can be used to determine the effects of acaricide treatment.56 Usually, engorged female ticks are used to determine the effect on egg production, and size and viability of progeny. Because of the size of adult ticks, test systems with relatively high volume are used. Serial dilutions of test articles are made in appropriate diluent. Subsequently, the required number of adult female ticks is immersed in the diluted test articles for 30 min. After immersion, ticks are taken from the solutions, dried, and fixed on Petri dishes. Next, ticks are incubated at 28 C 6 2 C and 80%
90% RH. Seven days later the proportion of ticks that have laid eggs is determined. This is referred to as then egg laying test (ELT). The reproductive estimate (RE) test (RET) is an extension of the ELT: Ticks are incubated for another week to determine mortality and determine the egg mass.57 At the completion of hatching, which may take 42 days for Rhipicephalus (Boophilus) ticks, the RE is calculated.

Feeding of test compounds The use of tick vaccines and of acaricides that are administered PO or systemically calls for in vitro systems that allow measuring the effects of compounds that are present in the plasma of treated animals. A number of feeding assays have been developed for studying the effects of antibodies against tick molecules. It is expected that similar assays will be developed for studying systemic acaricides. Larval engorgement test48 Critical for the feeding of tick larvae is the preparation of a feeding membrane that is thin enough for larvae to reach the blood and resilient enough to prevent leakage through punctures created by larvae. Gold beater’s skin membranes with a thickness .30 µm are treated with silicone are used.48 To further stimulate feeding the membrane is treated with a cow hair extract.58 The system uses a standard 24-well set up with the blood and/or serum in the upper compartment that is separated by the siliconized membrane from the compartment with the larvae underneath. The feeding unit is placed in an incubator at 37 C, 90% RH and 5% CO2. After incubation for 48 h

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the unit is frozen to stop feeding, and the number of larvae that has fed is determined. Adult feeding assays58 Two different systems have been developed for feeding of adult ticks: membrane feeding and capillary feeding. The feeding assays can be extended to measure ELT and/or RET as described earlier. Membrane feeding (Ixodes ricinus) Pieces of siliconized lens cleaning paper (Kodak, 70 3 120 mm) of 50
100 µm are used as feeding membranes.58 Feeding membranes are mounted onto the tubular of the feeding unit with silicon glue (internal diameter of B25 mm). Feeding units are placed in 6-well cell culture plates that contain 3.1 mL blood with test compounds. A ring that is fitted around the feeding unit ensures the unit rests on the rim of the well with the blood, ensuring free flow of blood at the membrane surface. To further stimulate feeding, a piece of glass fiber mosquito netting (1.4 mm mesh) is glued to the membrane in the feeding unit with silicone glue (Wacker Elastosil E4), and a plastic cross is placed on the membrane. In addition, each unit is pretreated with a cow hair extract. Next, 10 unfed female and 5 unfed male ticks are placed in each unit. The ticks are covered with short-cut cow hair and a brass grid to prevent ticks from escaping. Finally, the unit is closed with a stopper. Plates are incubated at 37 C, .95% RH. Blood with test compounds is refreshed at 12 h intervals. Mortality is evaluated daily, and dead ticks are removed. Capillary feeding59 In order to feed adult female ticks with capillary tubes, it appears of importance that these have been partially engorged on a suitable host. For in vitro feeding experiments, these ticks have to be collected by hand from the host that is used for partial feeding.59 Ticks are washed with warm (37 C) distilled water to remove animal tissue and/or tick cement. Ticks with intact mouthparts, devoid of any remaining tissue or cement cones, are selected. Ticks are mounted with their back onto glass microscope slides using double-sided tape. Slides are weighed prior to and after mounting of the ticks to determine tick weight prefeeding. Glass tubes with a diameter of 0.8
1 mm60 are carefully positioned over the ticks’ mouthparts under magnification using a dissection microscope. Tubes are filled with 200 µL of blood for overnight feeding (B18 h) in a 30 C chamber at 80% RH. After completion of feeding, tubes are removed and the slides with the fed ticks are each reweighed.

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Evaluation of test results Determination of acaricide activity In each of these assays, ticks from laboratory strains that are not resistant to acaricides are used. The effect of treatment with test article is compared with the effect of treatment with diluent only. Assays are performed at least in triplicate to allow calculating the average treatment effect, and statistical significance of difference with control groups. Depending on the viability of the ticks that were used in the assay, there may be variable mortality upon treatment with diluent only (background mortality). In general, assays in which background mortality surmounts 10% are rejected and must be repeated. Abbott’s formula is used to calculate the net effect of acaricide treatment by correcting for background mortality61: ð% live ticks in control groupÞ 2 ð% live ticks in treated groupÞ 3 100 5 % control % live ticks in the control group

Adjustments for changes in the control groups can also be made for other parameters, such as ELT and RE, using essentially the same formula. Determination of resistance to acaricides The same assays can also be used to determine the sensitivity of a tick isolate to acaricides. In such case the effect of the acaricide treatment on the tick isolate is compared with the effect of acaricide treatment on a tick strain with known sensitivity. For each of the tick strains the concentration that induces 50% mortality (LC50) is calculated. From these values the ratio LC50test/LC50control is calculated. For the different acaricides, defined cutoff values have been determined to classify tick strains as being Sensitive, Emerging-Resistant or Resistant.57

Deterrent, repellent, and behavioral assays For clarity, we adopt here the following definitions of repellents and deterrents as suggested by McMahon et al.62: A repellent is a compound whose vapor inhibits the response to an attractant, and a deterrent is a compound that inhibits the response to an arrestment stimulus (e.g., tick feces). These compounds do not necessarily kill the ticks, however, some compounds that kill ticks have repellent activity. The assays described later are only used for evaluating the effects of chemical compounds in a scientific laboratory setting. Deterrent assay62 In the deterrent assay the behavior of ticks is studied in Petri dishes (80 mm diameter and 15 mm high) in which strips of filter paper with test compounds are placed. Filter paper strips are first treated with an arrestment

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stimulus such as feces of newly molted male ticks and/or synthetic arrestment stimuli (e.g., a solution of 0.3 mM xanthine, 0.03 mM uric acid, and 0.03 mM 8-azaguanine in nanopure water).63 After drying, serially diluted test compounds are pipetted onto the paper strips. The paper strips are dried and placed at equal distances on the bottom of the Petri dish. In each Petri dish an adult tick is placed and left overnight in the dark. The next morning, the position of each tick is recorded. The sum of ticks on the treated filter paper strips is compared with the sum of ticks on the positive control strips. Ticks occurring elsewhere in the Petri dishes are ignored. Repellent assays Tick motion tracking A highly sophisticated apparatus was developed by Kramer,64 which allowed recording the route and speed at which a walking insect moves relative to a wind stream with or without odors. Briefly, the insect is placed on top of a Perspex ball (33 cm diameter) in a closed circular area through which a wind stream flows. The position/orientation of the insect is detected by a sensor placed right above it. Any change in the orientation and movement of the insect is compensated for by reorienting the Perspex ball such that the insect is maintained in its starting position; hence, if the insect walks forward, then the ball is rotated backward. The virtual track that the insect follows is deduced from the activity from the two servomotors that are needed to readjust the Perspex ball continuously (one motor adjusts forward
backward movements, the other one left
right movements). This system has been used to study tick attractants and repellents by McMahon et al.62 To test the response of ticks to attractants (e.g., pheromones) or repellents, test compounds are put on filter paper that is placed in a 500 mL gas-wash bottle. After 5 min of equilibration, the air from the gaswash bottle is added at a speed of 150 mL/min to the wind stream (3.5 L/ min) serving the area where the tick is placed. Each experiment comprises three distinct phases during which the movement of the tick is recorded: 1 min mix with air from solvent control bottle, immediately followed by 1 min presentation of the test stimulus, and finally 1 min representation of the solvent control. The system allows determining the combined effect of repellents that are mixed with attractants. In such case, two gas-wash bottles are used (one with attractant and one with repellent), and defined mixtures of the air from those bottles are fed to the main wind stream. The system allows determining the effect of defined compounds but is not suited for HTS. Olfactometer trials Attraction or avoidance of specific odors can also be measured using a system that presents a choice to the ticks (olfactometer). There is little or no

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standardization of olfactometers and in general researchers use their own home-made devices. Bissinger et al.65 used a four-choice olfactometer that consisted of four glass bulbs (19.0 3 89 cm) that are each mounted to an arm of a central cross-like piece in which ticks can be placed. The distance from the center of the device to the bulbs is 25 cm. Air with test compounds is fed through the glass bulbs in the direction of the central piece (50 mL/min), where excess of air is pumped off (200 mL/min).65 Test compounds are pipetted onto semicircular pieces of filter paper (31.8 cm2, Whatman No. 1) and allowed to dry. Afterward, the filter paper is folded to fit into the bulb. In each experiment, only one of the four bulbs contains filter paper with test compound, the others contain paper with water only. Tests are conducted at 25.4 C 6 0.1 C. For each experiment, eight unfed ticks (of either sex) are placed in the device, and after 15 min the number of ticks in each of the glass bulbs is recorded. Ticks that do not move in one of the four bulbs are recorded as nonresponders. Tick climbing assays Especially for tick species that climb onto vegetation waiting for a suitable host to pass, experimental setups in which ticks can climb onto rods have been developed. Test compounds are applied on the rods, and the number of ticks climbing up the rods is monitored. The assay is considered extremely time consuming if ticks are to be monitored continuously. It has been suggested that alternatively the position of ticks may be observed after fixed time intervals, but contacts between tick and repellent may be easily missed.66 A variant of this assay has been developed for Rhipicephalus appendiculatus by Madder and Berkvens.67 Instead of using rods, a gauze cylinder is used (35 cm long, 3.5 cm diameter). At the bottom of the cylinder a glass bottle with ticks can be fitted. At the top of the column an attractant is added (bovine ear grease and/or tick feces). In addition, the top end of the cylinder is warmed to around 38 C, the ear surface temperature of cattle, and air with 5% CO2 is passed over the column. When a bottle with ticks (10 males, 10 females) is mounted at the lower end of the cylinder, ticks climb up in the cylinder. The number of ticks in the cylinder is counted at various times until 1 h after release of the ticks. In order to study the effect of repellents, filter paper with dried test articles can be mounted in the top end of the column. Moving-object bioassay The tick climbing assay as described earlier studies the highly complex interaction between ticks and their environment, mimicking as much as possible the tick
host encounter. Most of the variables, however, form a continuous gradient (temperature, attractants, and repellents), and the system lacks the

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discontinuous event when a tick translocates from the surrounding vegetation to the vertebrate host. Dautel et al.68 developed a laboratory test system in which the ticks can display the natural behavior of clinging to a passing host. Briefly, ticks are placed at the end of a glass rod with its tip positioned closely to a slowly rotating vertical drum/cylinder. To attract ticks the temperature of the system is 35 C
36 C. An elevated part on the drum, which serves as a tick attachment site, passes the tip of the glass rod with every rotation at a distance that allows the tick to reach the elevation but not the surface of the drum/ cylinder. Filter paper with test substances can be applied to the tick attachment site. During a test, records are made of whether or not the tick moves toward the drum/cylinder (distance effect), clings to the attachment site, and if so, whether it holds on to the attachment site or drops off (contact repellence). Results obtained with this moving-object bioassay closely mimicked those obtained with static test systems using human volunteers.69 Evaluation of results The repellent/deterrent/behavioral assays are labor intensive and cannot be used for HTS of libraries of chemical compounds. In addition, there is little or no harmonization between the different research groups, and essentially individual laboratories rely on their own test system. This implies that the results obtained with different components should be regarded as scientific data that allow inference as to the repellent/deterrent/behavioral activity of a particular compound, but not about its relative activity as compared to other components. More standardization is required.

In vivo method(s) Introduction The use of animals as experimental animals to evaluate the efficacy of acaricides in vivo is a necessary requirement for establishing the full efficacy profile of a biological or pharmaceutic test substance. Current in vitro models cannot simulate actual on-host use for evaluating the effectiveness of acaricides in, for example, (1) killing the ticks already on the animal at the time of treatment (therapeutic efficacy); (2) repelling new host-seeking ticks and/or prevent/reduce biting; (3) killing ticks newly acquired by the host for a period of time after treatment [persistent (or residual) activity]; and (4) progressively reducing or eliminating off-host life cycle stages in the environment (see Appendix A of Ref. [70]). Consequently various scientific guidances have been published aiming at (1) aiding regulatory officials responsible for developing meaningful efficacy registration requirements within their countries, (2) assisting researchers in preparing basic plans for effective definition of the efficacy of an IVP, (3) reducing the number of study animals used in testing

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TABLE 4B.1 Applicable scientific guidance documents for evaluation of acaricidal efficacy on dogs, cats and ruminants addressing various aspects of trial design, biosafety and animal welfare considerations. Type

Document reference

Scientific guidance

Ref. [71], Appendix A

Scientific guidance

Ref. [70], Appendix A

Biosafety guideline

Ref. [72]

Biosafety guideline

Ref. [73]

Animal welfare

Ref. [74]

Animal welfare

Ref. [75]

for cost saving and animal welfare considerations, and (4) ensuring sufficient biosafety measures taken to protect the environment and the health and safety of research staff. Table 4B.1 summarizes the applicable scientific guidance documents that may be of interest to the reader, this excludes regulatory guidance’s specific for countries or regions. The published scientific guidance documents for tick evaluations on dogs, cats, and ruminants address the various aspects of trial design, biosafety, and animal welfare considerations as well as techniques to be used for tick challenges and assessments (see Appendix A of Refs. [70,71]). Since these techniques will form the basis of any good trial design, the purpose of this sections is to provide a more detailed overview of some techniques that can be used for infesting and counting ticks. Admittedly, numerous tick infesting and counting methods are published in the scientific literature. Techniques will be described in detail for use on cattle, dogs, cats, rabbits, and rats. These species represent the hosts most frequently used in vivo and the techniques described for these species can be adapted to other similar species if needed.

Infestation and count techniques for cattle Whole body infestations and counts suitable for one-host ticks (e.g., Rhipicephalus microplus, Rhipicephalus decoloratus, Rhipicephalus annulatus, and Rhipicephalus australis) Counting of larvae ticks needed for infestation The use of whole body infestation techniques on cattle are generally used for larvae of one-host tick species (e.g., R. microplus). In order to infest the experimental animals, samples for infestation must be prepared by the accurate weighing of a predefined weight of eggs that will result in, if

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successfully hatched, the required number of larvae. Alternatively, the required number of larvae must be counted or estimated without mechanically damaging the ticks. The weighing of eggs is a cumbersome approach with the accuracy dependent on the subset of eggs weighed fully hatching, which for many samples normally is not the case. Enumerating the required number of tick larvae prior to an infestation is a much more flexible approach and can easily be done using the following technique: Equipment: Graded glass or plastic vials (B10 and 20 mL) Tap water Soap dilution (10 parts tap water, 1 part commercial dishwashing soap) Pipette 500 mL wash bottle 500 mL wash bottle with tap water Filter paper (150 mm) Funnels Funnel stands Larval ticks of specific tick species Bulldog clamps Drying rack Masking tape Procedure: Place the funnels in the funnel stands and line the funnels with the filter paper. Place the required amount of filter paper and clips for closure on the workspace to ensure enough equipment is available before starting the counting process (number should equal the required number of samples to be prepared.). Add 5 drops of soap solution (dishwashing liquid) to 500 mL of water in a wash bottle (Fig. 4b.1). Screw the cap and shake the bottle, if there is excessive foam, redo using less soap. As a rough estimate a fully engorged blue tick may produce about 2000
4500 eggs.3 To estimate how many larvae there is in any given flask, it is advised to take a more conservative approach and base the estimate on the assumption that 2000 larvae successfully hatched per female used for oviposition. For example, if the flask contained 30 females, the assumption will be that there are at least 60,000 larvae in that specific flask. Transfer through the use of a spatula the estimated number of required larvae (contained in the flask), to the wash bottle containing the water (Fig. 4b.2). Larvae tend to stick to the sides, so start by scraping them from the sides. It is advisable to work on a white surface when manipulating the larvae as this will allow for easy monitoring for any accidental escapes during the transfer process.

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FIGURE 4B.1 A 500 mL wash bottle with water and flask with larvae.

FIGURE 4B.2 Transfer of larvae from flask to wash bottle.

Try to avoid transferring dead females, as they tend to get stuck in the spout of the wash bottle. Once the larvae are successfully transferred from the flask to the wash bottle, shake vigorously until the larval ticks are dispersed evenly within the solution. Ensure that no clumping is observed. While keeping the larvae mixture in motion by frequent light shaking, squirt 10 mL of the mixture into a 10 mL vial (Fig. 4b.3). Decant the vial into a funnel lined with filter paper (Fig. 4b.4). By using clean tap water,

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FIGURE 4B.3 Larvae squirted into graded vial.

FIGURE 4B.4 Decanting of vial into funnel.

ensure that all larvae are washed out of the vial. Repeat five times. Five packets will be needed to calculate the average per milliliter. Once all larvae are successfully transferred from the vial to the funnel line with filter paper, ensure that all larvae are washed from the sides of the filter paper to the bottom of the funnel. This can be done using the wash bottle with clean tap water prepared earlier in the procedure.

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FIGURE 4B.5 Process of folding the larval packet. (A) Collecting of larval ticks on filter paper. (B)
(E) Folding of filter paper containing ticks into a larval packet. (F) Crimping of larval packet containing ticks.

After all the water is filtered out, remove the filter paper containing the larvae, fold the top close, and clamp it so the larvae cannot escape (Fig. 4b.5). Dry the five packets by hanging them on the drying rack. A hairdryer can be used to speed up the drying process, just ensure that the heated air blown from the hairdryer is not used (Fig. 4b.6). Once the packets are dry, count the number of larvae in each of the five packets. Any method of counting may be used, but the following two examples are provided: 1. Tick larvae may be spread out evenly onto masking tape to simplify the counting procedure. Care must be taken to only count larvae, and not eggs, that may be present. Take a piece (B1 mm) of masking tape and stick it to the workspace. Then take the filter paper between the thumb and forefinger, and sprinkle larvae onto the masking tape ensuring to

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FIGURE 4B.6 Drying of larval packets.

FIGURE 4B.7 Counting of tick larvae by the spreading method. (A) Strip of tape with sticky end up. (B). Removing ticks from filter paper. (C) Placing ticks on tape strip. (D) Counting the number of ticks on tape strip.

spread them evenly, as this makes the counting process easier and more accurate (Fig. 4b.7). Once the larvae are sprinkled onto the masking tape, count the larvae. 2. Alternatively, the dried larval packet can be opened, and the larvae counted and collected by sticking them to a small piece (B5 cm long) of masking tape while being counted. Continue this process until you have 100 larvae counted and collected and then fold the masking tape into a packet to prevent larvae from escaping. Repeat until no larvae are left on the filter paper. The number of packets can then be counted to determine the number of larvae in the filter paper packet. Using this method, speedy counting is of the essence since larvae will start running off the filter paper if counting is to slow (Fig. 4b.8).

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FIGURE 4B.8 Counting larvae directly from filter paper. (A)
(C) legends.

When all five larval packets are counted, add all counts together to get a total of the five. Divide that by 50 (this is the amount in milliliter of the larval mixture used), this will be the average number of larvae per milliliter. Once the number of larvae per milliliter of the suspension in the wash bottle has been determined, the volume of suspension needed to be filtered to achieve the desired number of larvae needed for an infestation can be calculated. For example, if 3000 larvae are needed for infestation: Total counted in the five 10 mL aliquots/50 5 Bnumber of larvae/mL 3000/average number of larvae 5 mL needed to average 3000 larvae Packet No.

Amount per packet

1 2 3 4 5

1257 1301 1198 1214 1312

Total

6282

6282 larvae/50 5 B125 larvae/mL. 3000/125 5 24 mL needed/packet.

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Once the required volume to supply the correct number of ticks has been determined, the same procedure can be followed as described earlier to prepare the desired number of larval packets each containing the required number of larvae to be used for infestation. Note: Infestation must take place on the same day to ensure all larvae are still live and viable. Infesting cattle with tick larvae Equipment: Suitable retraining device, for example, neck clamp Prepared packets with larval ticks Procedure: Prior to infestation, the animal may be placed in a neck restraint to prevent the animal from grooming. Alternatively, an Elizabethan collar may be fitted to each calf prior to tick infestation. Neck restraints include neck clamps, crushes, and a chain secured around the neck of the animal to restrict movement. The filtration packet containing the larval ticks (B3000
5000, depending on study plan/protocol specifications) must be opened. Check the viability of the larvae prior to infestation by observing the movement of the larvae. Viable larvae will move over the filtration paper. If ticks are not viable (no movement or fall off the filtration paper), do not proceed with the tick infestation. Take care in windy conditions to prevent the ticks from blowing off of the filtration paper, and, if possible, postpone the infestation until conditions are more favorable. It is advised not to perform whole body tick infestations using larval ticks in windy conditions (this method is only applicable to outdoor housing facilities). Once the filtration paper is opened completely, hold the filter paper on the animal for a few seconds to allow the ticks to attach to the hair of the animal. Then brush it gently over the coat of the animal to remove the rest of the ticks from the filter paper. Ensure that all the larval ticks are removed from the filtration paper. Larval ticks must preferably be placed on the lateral aspects of the neck or as specified in the study plan/protocol. When multiple infestations are performed, alternate the side of infestation between subsequent infestations to spread the tick load evenly over the body of the animals. Animals may remain restrained (if applicable) for a minimum period of 1 h after infestation. During the period of restraint, animals should be monitored consistently for any signs of asphyxia or trauma. Gloves and aprons must be changed between groups. Always handle untreated control animals before treated groups (only applicable in nonblinded studies).

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FIGURE 4B.9 Cattle stanchion for collecting detached engorged ticks.

Counting of engorged one-host ticks The accurate enumeration of one-host ticks following whole body infestations of cattle with larvae is dependent on the collection of detached engorged female ticks. To be able to effectively collect the detached engorged female ticks undamaged, cattle need to be housed in stanchions specifically designed for this purpose (Fig. 4b.9). Depending on the stanchion design (e.g., collection tray versus flushing system), the following method may be used to collect and count ticks: Equipment: Wide-mouth plastic bottle (B100 mL) with lid Water hose with pressure nozzle Stanchions Tick collection pan Paper towels Ethyl alcohol (70%) Forceps Low-viscosity grease Procedure: A separate uniquely identified container must be used for the collection of ticks from each animal for each day. The containers should be identified with at least the study/project number, animal identification, and date. Slide out the tick collection tray from underneath the stanchion holding the animal. Visually examine the tray contents, feed bucket and water bucket for any ticks, whether engorged or not. Use forceps to collect all ticks and place them into a collection bottle. After collection of all visible ticks, spread any feces evenly in the collection tray in order to expose ticks possibly concealed by the feces. A water jet, spray, or suitable tool may be used for this purpose.

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Remove feces and clean the collection tray. Apply a thin layer of grease (which acts as a tick barrier) to the edges of the tray and place the clean collection tray back underneath the stanchion of the animal. If further assessments are required on the collected ticks (e.g., oviposition assessments), the collected ticks must be washed to remove any debris and then dried with a paper towel after they have been counted.

Localized infestations and counts suitable for one-host and multihost ticks (e.g., species of Rhipicephalus, Dermacentor, Ixodes, Amblyomma, and Hyalomma) Localized infestation techniques are suitable for one-host ticks as well as multihost ticks. This infestation method is well suited for evaluating the effect of systemically acting compounds as well as when ticks needs to be confined to specific predilection site such as the ears (e.g., R. appendiculatus). Larvae from one-host ticks and adult multihost ticks to be used for infestations can be counted as described previously. Patch infestations on cattle Equipment: Suitable retraining device, for example, neck clamp Prepared vials with required number of ticks Elizabethan collars Patches Contact adhesive Procedure: To prevent injury during the procedure, animals should be suitably restrained. Additional restraint for a minimum period of 1 h after tick infestation will also be required as well as fitting Elizabethan collars after patch attachment until the end of the tick collection period to prevent the animals from grooming/damaging the patches. This specific infestation method requires animals to be fitted with a tick infestation patch confining the ticks to a specified, limited area on the host animal (Fig. 4b.10). Patches are constructed from a suitable dense (to preclude tick escape) and durable (resistant to tearing) cloth material with a base flange (e.g., artificial leather) that can be glued to a shaven area on the host animal. The patch effectively creates a “sock” effect with the open end then knotted and/or secured by rubber bands to prevent escape of the ticks placed within (Fig. 4b.11). Depending on the size of patch used, a parting must be shaved on each animal the day prior to infestation. The patch should be glued to the shaved parting with preferably a medical grade glue (preferably a contact adhesive), or any other adhesive approved by an IACUC, 24 h prior to the tick

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FIGURE 4B.10 Patches are constructed from a suitable dense (to preclude tick escape) and durable (resistant to tearing) cloth material with a base flange (e.g., artificial leather).

FIGURE 4B.11 A patch glued to the shaved parting 24 h prior to the tick infestation.

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infestation (Fig. 4b.11). Follow the exact instructions of the glue to ensure the desired results. Once patches are securely fixed to the animals and the adhesive properly cured, the desired number of ticks can be infested in the patch. Secure the open end of the patch sock by tying a knot at the end of the sock, and/or secure the knot with a rubber band to prevent escape of the ticks placed within.

Infesting ticks in ear bags Equipment: Suitable retraining device, for example, neck clamp Prepared vials with required number of ticks Elizabethan collars Ear bags Contact adhesive Procedure: This method is most suitable for use when infesting ticks with specific predilection for the ears (e.g., R. appendiculatus). To prevent injury during the procedure, animals should be suitable restrained. Various different methods exist for infesting ticks in the ears of a variety of host species. The approach described here has however been found to provide one of the most secure fitment of the ear bags with very little irritation caused to the animal. The method relies on a “cap” designed with two ear socks securely fastened to the animal’s head as follows: Place a halter over the animal’s head so that it fits securely. Place the ear bags over the ears of the animal and tie them to the halter and around the head of the animal to prevent the ear bags from coming off of the ears (Fig. 4b.12). Open each ear bag and place the required number of ticks from the vial into each ear bag. Adult ticks will be stimulated to show an appetence response by tapping the vial and breathing on the ticks. Close the ear bags securely with the Velcro strips to preclude escape of the ticks placed within. It is advisable to fit animals with Elizabethan collars after tick infestation and until the tick collection to prevent the animals from grooming/ damaging the ear bags (Fig. 4b.13).

Infestation and count techniques for dogs (species of Amblyomma, Rhipicephalus, Haemaphysalis, Dermacentor, and Ixodes) Infesting dogs using a sedative Depending on the specific research objectives, sedating dogs prior to infestation may be required to limit confounding variables, for example, a dog’s

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FIGURE 4B.12 Ear bags are fitted over the ears of the animal and tied to the halter to prevent the ear bags from coming sliding back of the ears.

FIGURE 4B.13 A halter and ear bags tied together to prevent the ear bag from sliding of the animals head and ears. An Elizabethan collar is fitted to prevent the animal from grooming/damaging the ear bags.

behavioral response to infestation. The following procedure can be used for infesting sedated dogs with ticks. Equipment: Vials with ticks Infestation crate Masking tape/double-sided tape Suitable sedative for dogs

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Procedure: Infestations of dogs with ticks are generally performed in an infestation crate. Infestation crates are “tub” like structures designed with suitable dimensions to accommodate a dog and functions as the primary containment barrier for the ticks. Prior to the infestation procedure, crates should be prepared by placing double-sided tape around the top of the infestation crate or along the inside wall to prevent vertically migrating ticks from escaping. Once the crates have been prepared, dogs can be sedated with an IACUC-approved sedative (e.g., medetomidine hydrochloride at 0.06 mL/kg as IM injection) by a veterinarian or suitably qualified person. Ideally, sedation should be done B60 min prior to infestation to ensure that the dogs to be infested have been sedated sufficiently, that is, the dog should not be able to scratch or actively move around. Sedated animals should be monitored consistently throughout the duration of the sedation period for any adverse reactions to the sedatives. Once it has been established that the sedative has the desired effect, place the sedated dog in an infestation crate so that the dog is lying on its side. Stimulate the ticks to show an appetence response by tapping the vial and breathing on them. The behavior of the ticks in the vial should immediately change from docile to active, if they are viable. Release the ticks on the lateral aspects of the sedated dog, from the shoulder blades to the lateral stomach. Ticks may be dispersed using gloved fingers to prevent all the ticks from being released in one area on the dog. Check the vial that all ticks have been released and your hands to ensure that no ticks are clinging to them. Depending on the research objectives, a protocol may specify to infest the ticks next to the dog in the infestation crate and not directly on the dog. The same procedures should be followed, except the ticks will be released next to the back or the stomach of the sedated dog. Keep the dogs in the infestation crate for B1 h following tick infestation. Dogs that do not show signs of recovery from sedation following the 1 h tick infestation period should be administered a suitable antidote to reverse the effects of the sedative. Dogs should be removed from the crates as soon as the effects of the sedative wear off to prevent injury. Once the dog has been removed from the crate, collect all remaining ticks in the infestation crate.

Infesting nonsedated dogs Depending on the specific research objectives, it may be required to infest dogs without the use of sedation. This approach may require manually restraining the dog by hand in an infestation crate during the procedure or confining the dog to an infestation cage. This approach may be considered if (1) dogs need to be infested on multiple occasions and sedating repeatedly

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for infestation impacts the welfare of the animal, (2) a more natural host/tick interaction is required, and (3) sedation may potentially impact on the pharmacokinetics of a test substance. The following procedure can be used for infesting nonsedated dogs with ticks. Equipment: Vials with ticks Infestation crate or cage Masking tape/double-sided tape Procedure: Infestations of dogs with ticks may be performed in an infestation crate as previously described. This approach will require a dog to be manually restrained to allow the ticks a sufficient amount of time to penetrate the dog’s hair coat and start the process of finding a suitable attachment site. Prior to infestation, the dog may be placed in an infestation crate standing upright. It is advisable to have a second operator restraining the dog during the procedure. Stimulate the ticks to show an appetence response by tapping the vial and breathing on them. The behavior of the ticks in the vial should immediately change from docile to active, if they are viable. Release the ticks on the dorsal areas of the dog, from the neck downward toward the lower back. If the dogs are treated with a topical test substance, avoid placing the ticks directly onto the treatment area. Ticks may be dispersed using gloved fingers to prevent all the ticks from being released in one area on the dog. Once the ticks are released, check the vial for any remaining ticks as well as your hands to ensure that no ticks are clinging to them. Manually restrain the dogs for at least 15 min, during this period all ticks that fall off the dog can be placed back. This period may be extended based on the research objectives. If infestations are not directly on the dog, but obtained from the immediate environment or ticks are collected from the environment for extended periods of time, infestations should be performed in an infestation cage. The basic design principles of these cages are similar to that of an infestation crate with the added functionality that a dog may be enclosed safely in it while still preventing ticks from escaping. It should however be noted that the dimensions of these cages are greater than that of a crate since dogs may be enclosed in these cages for extended periods of time. The dimensions of the cages should be approved by the IACUC.

Counting and removing ticks on dogs Equipment: Forceps suitable for tick removal or a tick removing device Fine-toothed comb (B11
13 teeth/cm)

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Container with suitable solution to kill and preserve removed ticks if required Procedure: The counting of ticks on dogs relies on the methodical examination of the skin by palpation and parting of the hair coat. To ensure all ticks are found the whole body must be examined, specifically the following areas: outside hind legs, including feet tail and anal areas lateral area, not including shoulders abdominal area, from chest to inside hind legs forelegs and shoulders, including feet all neck and head areas dorsal strip from shoulder blades to base of tail

G G G G G G G

Depending on the specific research protocol, it may be required to count the ticks in situ (thumb count) or removing them. To remove ticks, extraction must be done using forceps or a tick removing device by grasping the tick as close as possible to the skin and carefully twisting it so that the rostrum does not remain in the skin of the dog. Once removed, ticks can be categorized according to Marchiondo et al.71 Appendix A as follows in Table 4b.2: To ensure all ticks are removed, the dog should be combed with a finetoothed comb for a set time period after the whole body is examined.

Infestation and count techniques for cats (species of Amblyomma, Rhipicephalus, Haemaphysalis, Dermacentor, and Ixodes) Infesting cats with ticks In order to achieve any reasonable infestation rate on cats, they need to be sedated during the procedure and grooming as much as possible limited PI. The following procedure can be used for infesting sedated cats with ticks.

TABLE 4B.2 Tick category assignments to assess the parasiticidal effect of treatment (see Appendix A of Ref. [71]). Category

General findings

Attachment status

Acaricidal effect

1

Live

Free

No

2

Live

Attached

No

3

Dead

Free

Yes

4

Dead

Attached

Yes

a

a

If justified, only live attached tick counts may be used to assess efficacy for systemically acting acaricides.

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Equipment: Vials with ticks Infestation crate Masking tape/double-sided tape Suitable sedative for cats Elizabethan collar for cats Procedure: Prepare infestation crates as described for dogs. Cats must be sedated by an IACUC-approved sedative suitable for cats by a veterinarian or suitably qualified person. Prior to placing the cat in the infestation crate, ensure that the sedative has taken full effect, the cat should not be able to scratch or actively move around. Sedated animals should be monitored consistently throughout the duration of the sedation period for any adverse reactions to the sedatives. Once it has been confirmed that the sedative has reach full effect, place the sedated cat in an infestation crate so that the cat is lying on its side and fit an Elizabethan collar. Care should be taken not to fasten the Elizabethan collar excessively, the fit should be loose enough to not cause any breathing restrictions. Stimulate the ticks to show an appetence response by tapping the vial and breathing on them. The behavior of the ticks in the vial should immediately change from docile to active if they are viable. Release the ticks on the lateral aspects of the sedated cat, from the shoulder blades to the lateral stomach. Ticks may be dispersed using gloved fingers to prevent all the ticks from being released in one area on the cat. Check the vial that all ticks have been released and your hands to ensure that no ticks are clinging to them. Cats should remain in the infestation crate for as long as possible while the effect of the sedative still lasts. During this period the crates should be routinely inspected and all ticks not on the cat placed back. Cats that do not show signs of recovery from sedation should be administered a suitable antidote (e.g., yohimbine) to reverse the effects of the sedative. Cats should be removed from the crates as soon as the effects of the sedative wear off to prevent injury. Once the cat has been removed from the crate, collect all remaining ticks in the infestation crate.

Counting and removing ticks on cats The method for counting and removing ticks from cats is similar to that described for dogs.

Infestation and count techniques for rabbits (species of Amblyomma, Rhipicephalus, Haemaphysalis, Dermacentor, and Ixodes) The methods for infesting, collecting, and counting of ticks on rabbits are described in detail by Levin and Schumacher,46, Appendix B included as a supplement to this chapter.

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Rabbit repellency assay An in vivo screen has been developed to determine tick repellency on rabbits. On TD 21, each rabbit (New Zealand White, B1.5
3 kg) is clipped close to the skin and a sleeve attached. A collar is fitted onto each rabbit following the attachment of the sleeve. The animals are housed in individual cages at 18 C, 55 6 10% RH with a 12L:12D hour cycle. On TD 0, placebo or test compounds are applied (total volume of 1 µg/mL (microgram/mL)) in even streaks to the tick feeding site, an area enclosed by the sleeve outlet, using a Gilson pipette. A challenge of 10 R. sanguineus adults (5 of each sex) is added to each treated area 1 h following application of the compound. Observations are made at 6, 12, and 24 h PT in order to establish the time of attachment, repellence, or toxicity of ticks. Repellence is assessed by comparing the percent successful attachment of adults on treated areas with those on the placebo-treated rabbits. Treatment groups that repel 9 or more of the 10 challenge ticks are judged to have good repellence activity. Tick mortality, expellency, hyperactivity, and behavioral effects differing from the placebo-treated rabbits are recorded. All treatments and the positive control (amitraz) are compared to the placebo. This assay can be conducted with R. appendiculatus adults, nymphs or larvae with evaluation 14 days PT. Infestation and count techniques for rats (species of Amblyomma, Rhipicephalus, Haemaphysalis, Dermacentor, and Ixodes) Rats can be successfully used as experimental animals for the in vivo evaluation of acaricidal activity. The tick life stages that can be used are usually limited to larvae and nymphs. In order to achieve any reasonable infestation rate on rats, they need to be sedated during the procedure and grooming as much as possible limited PI. The following method can be used for infesting and counting ticks on rats.

Infesting rats with ticks Equipment: Vials with ticks Heated bench top Ophthalmic solution for rats Suitable sedative for rats Elizabethan collar for rats Procedure: Rats must be sedated by an IACUC-approved sedative administered by a veterinarian or suitably qualified person. Always work with sedated rats on a

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heated surface to limit body temperature loss of the animal. Once the rat is sedated, an ophthalmic solution (eye drops or eye gel, e.g., Refresh Liquigel) should be applied to each eye and an Elizabethan collar fitted. Collar fit should not constrict breathing but should be tight enough to prevent removal over the head (Fig. 4b.14). Once the preparation of the rat is complete, the required number of tick nymphs or larvae can be infested. It is advisable to dislodge nymph or larval ticks from the vial wall by tapping on the vial and have ticks grouped at the bottom immediately prior to infestation. This will greatly simplify the action of placing the tick on the rat. Ticks can be released on the lower back of the rat if the rat is resting flat on its stomach or along the flank if the rat is rested on its side. Avoid clusters of ticks holding onto each other and rolling off from the rat by applying ticks to these areas (Fig. 4b.15). Once released, observe the vial that contained the ticks to ensure that all ticks were released.

FIGURE 4B.14 Attaching and adjusting the Elizabethan collar. (A) Rat held in cupped hands with neck extended. (B) and (C) An Elizabethan collar fitted comfortably around the neck and tested via (B) rotation and (C) linear movement of the collar along the neck.

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FIGURE 4B.15 Dislodging ticks from vial sides and infesting on rat. (A) Plastic vial containing live ticks. (B) Ticks gathered at the bottom of the vial after dislodging ticks from vial cap and wall. (C) Vial cap visually inspected for ticks. (D) Ticks released from vial along the lower dorsal side of the rat.

Counting ticks on rats Equipment: Analog tally counters Heated bench top Ophthalmic solution for rats Suitable sedative for rats Procedure: To effectively count and remove ticks, rats must be sedated by an IACUC-approved sedative administered by a veterinarian or suitably qualified person. Always work with sedated rats on a heated surface to limit body temperature loss of the animal. Once the rat is sedated, an ophthalmic solution (eye drops or eye gel, e.g., Refresh Liquigel) should be applied to each eye and the Elizabethan collar removed. Ticks can be found by a methodical search of the body using a pair of forceps to create partings in the rat’s fur coat and searching for ticks within the partings. To remove ticks, extraction must be done using forceps or a tick removing device by grasping the tick as close as possible to the skin and carefully twisting it so that the rostrum does

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not remain in the skin of the dog. Once removed, ticks can be categorized according to Marchiondo et al.71 Appendix A (Table 4b.2).

Rodent acaricidal assay An in vivo surrogate animal bioassay has been developed capable of rapidly and accurately predicting the topical activity of acaricides emerging from in vitro acaricide bioassays.76 The rat acaricide test (RAT) requires adult rats, Rattus norvegicus, a flexible tick containment device fastened to their dorsal-thoracic region, and the nymphal stage of the lone star tick, A. americanum. The feeding kinetics of A. americanum nymphs on rats was assessed, and compound efficacies were determined by measuring tick survivorship and engorgement weight on acaricide-treated animals. Results from this bioassay demonstrated efficacy with fipronil, ivermectin, permethrin, and chlorpyrifos, and dose
response relationships for each acaricide were determined. The rank order of potencies was fipronil . ivermectin . chlorpyrifos 5 permethrin for nymphal mortality and fipronil . ivermectin . chlorpyrifos . permethrin for inhibition of nymphal engorgement. The activity of permethrin against nymphs in the RAT was positively correlated with potency values for technical and commercial permethrin formulations against adult A. americanum infestations on cattle. The RAT proved to be an economical, rapid surrogate animal bioassay that together with the in vitro acaricide bioassay can be used for the rapid identification, characterization, and prioritization of candidate acaricides.

References 1. Coetzer JAW, Tustin RC. Infectious diseases of livestock. Cape Town: Oxford University Press; 2004. 2. McEnroe WD. A determination of tick population size within an area of migration [Dermacentor variabilis (Say) (Acari, Ixodidae)]. J Appl Entomol 1985;99(1-5):422
5. 3. Wall R, Shearer D. Veterinary ectoparasites. biology, pathology & control. 2nd ed Oxford: Blackwell Scientific Ltd; 2001. p. 262. 4. Guglielmone AA, Robbins RG, Apanaskevich DA, Petney TN, Estrada-Pen˜a A, Horak I. The hard ticks of the world., 10. Dordrecht: Springer; 2014. p. 978
94 (1007). 5. Estrada-Pen˜a A, Bouattour A, Camicas JL, Walker AR. Ticks of domestic animals in the Mediterranean region. A guide to identification of species. In: Int consortium ticks and tick borne dis, European Union, vol. 131. 2004. p. 137. 6. Guzma´n-Cornejo C, Robbins RG. The genus Ixodes (Acari: Ixodidae) in Mexico: adult identification keys, diagnoses, hosts, and distribution. Revista mexicana de biodiversidad 2010;81(2):289
98. 7. Castro MB, Wright SA. Vertebrate hosts of Ixodes pacificus (Acari: Ixodidae) in California. J Vector Ecol 2007;32(1):140
9. 8. Furman DP, Loomis EC. Bull CA Insect Survey The ticks of California (Acari: Ixodida), vol. 25. Berkeley and Los Angeles, CA: University of California Press; 1984. p. 239.

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9. Barlough JE, Madigan JE, Kramer VL, Clover JR, Hui LT, Webb JP, et al. Ehrlichia phagocytophila genogroup rickettsiae in ixodid ticks from California collected in 1995 and 1996. JCM 1997;35(8):2018
21. 10. Lane RS, Peek J, Donaghey PJ. Tick (Acari: Ixodidae) paralysis in dogs from northern California: acarological and clinical findings. J Med Entomol 1984;21(3):321
6. 11. Belozerov VN, Naumov RL. Nymphal diapause and its photoperiodic control in the tick Ixodes scapularis (Acari: Ixodidae). Folia Parasit 2002;49(4):314
18. 12. Oliver Jr JH, Owsley MR, Hutcheson HJ, James AM, Chen C, Irby WS, et al. Conspecificity of the ticks Ixodes scapularis and I. dammini (Acari: Ixodidae). J Med Entomol 1993;30(1):54
63. 13. Keirans JE, Hutcheson HJ, Durden LA, Klompen JSH. Ixodes (Ixodes) scapularis (Acari: Ixodidae): redescription of all active stages, distribution, hosts, geographical variation, and medical and veterinary importance. J Med Entomol 1996;33(3):297
318. 14. Carroll JF, Kramer M. Winter activity of Ixodes scapularis (Acari: Ixodidae) and the operation of deer-targeted tick control devices in Maryland. J Med Entomol 2003;40 (2):238
44. 15. Randolph SE, Green RM, Hoodless AN, Peacey MF. An empirical quantitative framework for the seasonal population dynamics of the tick Ixodes ricinus. Int J Parasitol 2002;32:979
89. 16. Estrada-Pen˜a A, Venzal JM, Sanchez Acedo C. The tick Ixodes ricinus: distribution and climate preferences in the western Palaearctic. Med Vet Entomol 2006;20:189
97. 17. Salman M, Tarres-Call J, editors. Ticks and tick-borne diseases, geographical distribution and control strategies in the Euro-Asia region. Wallingford: CABI; 2013. 18. Estrada-Pen˜a A, Nava S, Petney T. Description of all the stages of Ixodes inopinatus n. sp. (Acari: Ixodidae). Ticks Tick-Borne Dis 2014;5(6):734
43. 19. Jaenson TG, T¨aLleklint L, Lundquist L, Olsen B, Chirico J, Mejlon H. Geographical distribution, host associations, and vector roles of ticks (Acari: Ixodidae, Argasidae) in Sweden. J Med Entomol 1994;31(2):240
56. 20. Kolonin GV. Fauna of ixodid ticks of the world. ,http://www.kolonin.org/.; 2009 [last accessed 08.06.10]. 21. Hillyard P. Ticks of North-West Europe. Natural History Museum London; 1996. 22. Goddard J, Varela-Stokes AS. Role of the lone star tick, Amblyomma americanum (L.), in human and animal diseases. Vet Parasitol 2009;160:1
12. 23. Kollars Jr TM, Oliver Jr JH, Durden LA, Kollars PG. Host associations and seasonal activity of Amblyomma americanum (Acari: Ixodidae) in Missouri. J Parasitol 2000;86:1156
9. 24. Mock DE, Applegate RD, Fox LB. Preliminary survey of ticks (Acari: Ixodidae) parasitizing wild turkeys (Aves: Phasianidae) in Eastern Kansas. J Med Entomol 2001;38:118
21. 25. Childs JE, Paddock CD. The ascendancy of Amblyomma americanum as a vector of pathogens affecting humans in the United States. Ann Rev Entomol 2003;48:307
37. 26. Sumner JW, Durden LA, Goddard J, Stromdahl EY, Clark KL, Reeves WK, et al. Gulf coast ticks (Amblyomma maculatum) and Rickettsia parkeri, United States. Emerg Infect Dis 2007;13(5):751. 27. Teel PD, Ketchum HR, Mock DE, Wright RE, Strey OF. The Gulf Coast tick: a review of the life history, ecology, distribution, and emergence as an arthropod of medical and veterinary importance. J Med Entomol 2010;47:707
22. 28. Goddard J, Paddock CD. Observations on distribution and seasonal activity of the Gulf Coast Tick in Mississippi. J Med Entomol 2005;42:176
9.

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29. Cilek JE, Olsen MA. Seasonal distribution and abundance of ticks (Acari: Ixodidae) in northwestern Florida. J Med Entomol 2000;37(3):439
44. 30. Beati L, Keirans JE. Analysis of the systematic relationships among ticks of the genera Rhipicephalus and Boophilus (Acari: Ixodidae) based on mitochondrial 12S ribosomal DNA gene sequences and morphological characters. J Parasitol 2001;87(1):32
48. 31. Labruna MB, Naranjo V, Mangold AJ, Thompson C, Estrada-Pen˜a A, Guglielmone AA, et al. Allopatric speciation in ticks: genetic and reproductive divergence between geographic strains of Rhipicephalus (Boophilus) microplus. BMC Evol Biol 2009;9(1):46. 32. Hoogstraal H. Changing patterns of tickborne diseases in modern society. Annu Rev Entomol 1981;26(1):75
99. 33. Madder M, Thys E, Geysen D, Baudoux C, Horak I. Boophilus microplus ticks found in West Africa. Exp Appl Acarol 2007;43(3):233
4. 34. de la Fuente J, Estrada-Pena A, Venzal JM, Kocan KM, Sonenshine DE. Overview: ticks as vectors of pathogens that cause disease in humans and animals. Front Biosci 2008;13 (13):6938
46. 35. Jongejan F, Uilenberg G. The global importance of ticks. Parasitology 2004;129(S1): S3
14. 36. Walker AR, Bouattour A, Camicas JL, Estrada-Pena A, Horak IG, Latif AA, et al. Ticks of domestic animals in Africa: a guide to identification of species. Biosci Rep 2003;217. 37. Heath ACG. Biology, ecology and distribution of the tick, Haemaphysalis longicornis Neumann (Acari: Ixodidae) in New Zealand. NZ Vet J 2016;64(1):10
20. 38. Hoogstraal H, Roberts FH, Kohls GM, Tipton VJ. Review of Haemaphysalis (Kaiseriana) longicornis Neumann (resurrected) of Australia, New Zealand, New Caledonia, Fiji, Japan, Korea, and northeastern China and USSR, and its parthenogenetic and bisexual populations (Ixodoidea, Ixodidae). J Parasitol 1968;1197
213. 39. Heath ACG. Vector competence of Haemaphysalis longicornis with particular reference to blood parasites. Surveillance 2002;29(4):12
14. 40. Birkenheuer AJ, Correa MT, Levy MG, Breitschwerdt EB. Geographic distribution of babesiosis among dogs in the United States and association with dog bites: 150 cases (2000
2003). J Am Vet Med Assoc 2005;227(6):942
7. 41. Hoogstraal H, Wassef HY, B¨uttiker W. Ticks (Acarina) of Saudi Arabia. Fam. Argasidae, Ixodidae. Fauna Saudi Arabia 1981;3:25
110. 42. Mackereth G, Cane RP, Snell-Wakefield A, Slaney D, Tompkins D, Jakob-Hoff R, et al. Vectors and vector borne diseases: ecological research and surveillance development in New Zealand
risk assessment. Wellington, New Zealand: Ministry of Agriculture and Forestry; 2007. p. 64. 43. Jameson LJ, Morgan PJ, Medlock JM, Watola G, Vaux AG. Importation of Hyalomma marginatum, vector of Crimean-Congo haemorrhagic fever virus, into the United Kingdom by migratory birds. Ticks Tick-Borne Dis 2012;3(2):95
9. 44. Guglielmone AA, Robbins RG, Apanaskevich DA, Petney TN, Estrada-Pen˜a A, Horak IG. Comments on controversial tick (Acari: Ixodida) species names and species described or resurrected from 2003 to 2008. Exp Appl Acarol 2009;48(4):311. 45. Gharbi M, Darghouth MA. A review of Hyalomma scupense (Acari, Ixodidae) in the Maghreb region: from biology to control. Parasite 2014;21:2. 46. Levin ML, Schumacher LBM. Manual for maintenance of multi-host ixodid ticks in the laboratory. Exp Appl Acarol 2016;70:343
67. 47. Schetters T. Vaccination against ticks. In: Meng C, Sluder A, editors. Drug discovery in infectious diseases. Wiley-Blackwell; 2018 (in print).

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48. Trentelman J, Kleuskens J, Van de Crommert J, Schetters TPM. A new method for in vitro feeding of Rhipicephalus australis (formerly Rhipicephalus microplus) larvae: a valuable tool for tick vaccine development. Parasit Vectors 2017;10:153. 49. Stone BF, Haydock P. A method for measuring the acaricide susceptibility of the cattle tick Boophilus microplus. Bull Entomol Res 1962;53:563
78. 50. Shaw RD. Culture of an organophosphorus resistant strain of Boophilus microplus (Can.) and an assessment of its resistance spectrum. Bull Entomol Res 1966;56:389
405. 51. FAO Working Group on Parasite Resistance. Resistance management and integrated parasite control. In: Ruminants: guidelines. Module 1. Ticks: acaricide resistance: diagnosis, management and prevention. 2004. p. 25
77. 52. Lovis L, Perret J-L, Bouvier J, Fellay J-M, Kaminsky R, Betschart B, et al. A new in vitro test to evaluate the resistance level against acaricides of the cattle tick, Rhipicephalus (Boophilus) microplus. Vet Parasitol 2011;182:269
80. 53. Shaw RD, Cook M, Carson RE. Developments in the resistance status of the southern cattle tick to organophosphorus and carbamate insecticides. J Econ Entomol 1968;61:1590
4. 54. Klafke GM, Sabatini GA, de Albuquerque TA, Martins JR, Kemp DH, Miller RJ, et al. Larval immersion tests with ivermectin in populations of the cattle tick Rhipicephalus (Boophilus) microplus (Acari: Ixodidae) from State of Sao Paulo, Brazil. Vet Parasitol 2006;142(3-4):388
90. 55. White WH, Plummer PR, Kemper CJ, Miller RJ, Davey RB, Kemp DH, et al. An in vitro larval immersion microassay for identifying and characterizing candidate acaricides. J Med Entomol 2004;41(6):1034
42. 56. Drummond RO, Ernst SE, Trevino JL, Gladney WJ, Graham OH. Boophilus annulatus and B. microplus: laboratory tests of insecticides. J Econ Entomol 1973;66:130
3. 57. Mekonnen S, Bryson NR, Fourie LJ, Peter RJ, Spickett AM, Taylor RJ, et al. Comparison of 3 tests to detect acaricide resistance in Boophilus decoloratus on dairy farms in the Eastern Cape Province, South Africa. J S Afr Vet Assoc 2003;74:41
4. 58. Kr¨ober T, Guerin PM. In vitro feeding assay for hard ticks. Trends Parasitol 2007;23 (9):445
9. 59. Lew-Tabor AE, Bruyeres AG, Zhang B, Rodriguez Valle M. Rhipicephalus (Boophilus) microplus tick in vitro feeding methods for functional (dsRNA) and vaccine candidate (antibody) screening. Ticks Tick-Borne Dis 2014;5:500
10. 60. Willadsen P, Kemp DH, McKenna RV. Bloodmeal ingestion and utilization as a component of host specificity in the tick, Boophilus microplus. Z Parasitenkd 1984;70:415
20. 61. Abbott WS. A method of computing the effectiveness of an insecticide. J Econ Entomol 1925;18:265
7. 62. McMahon C, Kr¨ober T, Guerin PM. In vitro assays for repellents and deterrents for ticks: differing effects of products when tested with attractant or arrestment stimuli. Med Vet Entomol 2003;17(370):378. 63. Grenacher S, Kr¨ober T, Guerin PM, Vlimant M. Behavioural and chemoreceptor cell responses of the tick, Ixodes ricinus, to its own faeces and faecal constituents. Exp Appl Acarol 2001;25:641
60. 64. Kramer E. The orientation of walking honeybees in odor fields with small concentration gradients. Physiol Entomol 1976;1:27
37. 65. Bissinger BW, Apperson CS, Watson DW, Arellano C, Sonenshine DE, Roe RM. Novel assays and the comparative repellency of BioUDs, DEET and permethrin against Amblyomma americanum. Med Vet Entomol 2011;25:217
26.

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66. Dautel H. Mini review. Test systems for tick repellents. Int J Med Microbiol 2004;293:182
8. 67. Madder M, Berkvens DL. Evaluation of an in vitro method to measure behavioural diapause in the tick Rhipicephalus appendiculatus (Acarina: Ixodidae) in the laboratory. Parasitology 1997;115:97
100. 68. Dautel H, Kahl O, Siems K, Oppenrieder M, M¨uller-Kuhrt L, Hilker M. A novel test system for detection of tick repellents. Entomol Exp Appl 1999;91:431
41. 69. Dautel H, Dippel C, Werkhausen A, Diller R. Efficacy testing of several Ixodes ricinus tick repellents: different results with different assays. Ticks Tick-Borne Dis 2013;4:256
63. 70. Marchiondo AA, Holdsworth PA, Fourie LJ, Rugg D, Hellman K, Snyder DE, et al. World Association for the Advancement of Veterinary Parasitology (W.A.A.V.P.) second edition: guidelines for evaluating the efficacy of parasiticides for the treatment, prevention and control of flea and tick infestations on dogs and cats. Vet Parasitol 2013;94:84
97. 71. Holdsworth PA, Kemp D, Green P, Peter RJ, De Bruin C, Jonsson NN, et al. World Association for the Advancement of Veterinary Parasitology (W.A.A.V.P.) guidelines for evaluating the efficacy of acaricides against ticks (Ixodidae) on ruminants. Vet Parasitol 2006;136(1):29
43. 72. Arthropod Containment Guidelines. A project of the American Committee of Medical Entomology and American Society of Tropical Medicine and Hygiene. Vector-Borne Zoonot Dis 2003;3(2):61
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32.

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Chapter 4c

Arachnida, Metastigmata, Ixodidae, Ixodes holocyclus Maurice C. Webster, BVSc, MACVSc1, Ian S. Ridley, BRurSc1 and Alan A. Marchiondo, MS, PhD2 1 Invetus Pty Ltd., Armidale, NSW, Australia, 2Adobe Veterinary Parasitology Consulting LLC, Santa Fe, NM, United States

Arachnida Metastigmata Ixodidae

Ixodes holocyclus—Neumann, 1899—the Australian paralysis tick Biology and life cycle The most comprehensive review of the biology, life cycle, and pathogenesis of Ixodes holocyclus was published by Barker and Walker.1 Ixodes holocyclus is considered the most important medical and veterinary tick in Australia and is responsible for most of tick paralysis cases reported in humans, domestic animals, and wildlife. The tick has a geographic distribution that is restricted to a strip on the east coast that extends from north of Cairns in far northern Queensland to the south of the New South Wales (NSW)—Victoria state border.2 I. holocyclus is nonhost specific and has been recorded on 34 species of mammals, including humans, and 7 species of birds.1 Bandicoots (Isoodon macrourus and Perameles nasuta) are considered the principal hosts and are likely to be required for the populations of I. holocyclus to persist between seasons, at least in southeastern Queensland.3 I. holocyclus is a three-host tick.4 Although all stages have been reported at most times during the year5; one main generation per year. The tick undergoes four life cycle stages, egg, larva, nymph, and adults, and requires three-hosts for blood meals. Six-legged larvae hatch from the eggs after an incubation period of 50
110 days. Larvae quest for mammalian, avian or reptilian hosts and takes a blood meal for about 4
6 days before dropping from the host and molting to an eight-legged nymph. Nymphs require a blood meal to molt to the adult stage in B3
11 weeks. The unengorged female tick is yellow in color and becomes gray after feeding on a host for about 6
30 days. The female tick begins to produce

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toxin in B72 h that causes paralysis, if the host is not immune. Adult male ticks seek out a female to mate and do not engorge. Female ticks detach from the host with oviposition beginning 11
20 days at a rate of 20
200 eggs/day, but they die a few days after oviposition ceases. The life cycle can be rapid at 27 C with all stages requiring humid conditions. Temperatures .30 C or ,7 C can be fatal within a few days.6,7 The life cycle can be as long as 379 days.8 Larvae are more common between January and March, nymphs between April and September, and adults between October and December. The cases of canine paralysis have been reported to peak in September, before the peak numbers of adults are found3,9 and may reflect a seasonal decline in acquired immunity in the hosts. Dogs, cats, sheep, goats, and some wildlife are particularly susceptible to holocyclotoxin, but calves and foals also succumb.10 Holocyclotoxicosis typically presents as a rapid ascending flaccid paralysis. Death is caused by the paralysis of respiratory muscles. Other clinical signs include changes in vocalization, vomiting, inappetence, and unequal pupil size.11,12 This tick injects a neurotoxin (holocyclotoxin)13 into its host while feeding, releasing increasing quantities B72 h after tick attachment. The most prominent feature of toxin intoxication in the dog is dysfunction of the efferent motor system, although some disturbance of the afferent pathways and involvement of the autonomic nervous system can occur.14 The period elapsing between the attachment of female I. holocyclus and onset of signs varied from 5.5 to 7 days, while the mean duration of the disease is 23.3 h.14 I. holocyclus causes paralysis, which can be rarely fatal, and hypersensitivity, including acute anaphylactic shock in humans.15 The tick acts as a vector for Rickettsia australis, the cause of Queensland tick typhus.

Rearing method(s) The life cycle of I. holocyclus under laboratory conditions has been published by Goodrich et al.16 Guinea pigs were used for larval feeding, while bandicoots were used for nymph and adult tick feeding.16 The laboratory colony maintained the life cycle for 3 years producing 3000 female I. holocyclus annually.16 However, the use of native animals to feed I. holocyclus ticks under laboratory conditions is now strictly regulated, and no laboratory rearing of I. holocyclus is currently conducted in Australia to the authors’ knowledge. Levin and Schumacher17, APPENDIX B have published a manual of step-bystep recommendation for various procedures used in the maintenance of ixodid tick colonies. Procedures and methods of this manual with respect to facilities, personal protective equipment, workstation, housing ticks, feeding ticks on animals, and storing ticks might be useful for colony maintenance of I. holocyclus. Wild-caught unfed, female ticks of I. holocyclus are collected from vegetation during October to December each year from localities within the Northern Rivers region of NSW or from far north Queensland, Australia.

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Ticks are stored in a dark incubator at 12 C
15 C and high humidity before being placed on dogs or cats. Ticks will remain viable for 12 months under these conditions, but mortality does increase with time, often due to fungal overgrowth in the storage containers. Low humidity results in desiccation of the ticks. Adult unengorged female I. holocyclus are used to infest dogs and cats to increase their antibody titer against holocyclotoxin prior to collecting and processing blood to produce tick serum antitoxin used in the treatment of animals with holocyclotoxicosis. Dogs, cats, and calves are also infested to assess the efficacy of acaricides.

In vitro method(s) Artificial feeding of unfed or partially fed female I. holocyclus has been accomplished using the tissue culture medium TC199 plus additives covered by a silicone rubber membrane.18 Ticks have also been maintained for in vitro testing using fresh cattle blood (Jackson C, Pers. Comm.). The feeding pattern of artificially fed I. holocyclus was similar to that of naturally fed ticks. This method could be used to test the systemic activity of parasiticides against I. holocyclus by adding test compounds to the medium wells. An in vitro method of testing acaricides against ticks, which may be applicable to I. holocyclus in certain circumstances, is provided in the WAAVP guidelines for evaluating the efficacy of parasiticides for the treatment, prevention, and control of fleas and ticks on dogs and cats (see Appendix A of Refs. [19,20]).

In vivo method(s) Parasiticide efficacy studies against I. holocyclus should be conducted in compliance with the WAAVP guidelines for evaluating the efficacy of parasiticides for the treatment, prevention, and control of fleas and ticks on dogs and cats (see Appendix A of Refs. [19,20]), the Australian Pesticides and Veterinary Medicines Authority (APVMA) Preamble for the WAAVP guideline for fleas and ticks on dogs and cats (https://apvma.gov.au/node/1040)21 and the VICH GL9 Good Clinical Practices.22 WAAVP has also published guidelines for evaluating the efficacy of acaricides against ticks on ruminants (see Appendix A of Ref. [23]). APVMA has published a preamble (https:// apvma.gov.au/node/904) that notes Australia’s unique environmental and geographic parameters, parasite burdens and their population dynamics, and farm management practices and animal breeds, which means there are some differences between the WAAVP guidelines and the APVMA’s recommendations for the efficacy trials for products to be registered in Australia. Applicants are advised to conduct efficacy trials within Australia under typical farm management practices covering relevant geographical regions and to contact APVMA for additional guidance.

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Various aspects of parasiticide efficacy testing against I. holocyclus are not specifically covered in the WAAVP or APVMA guidelines.19
21,23 In addition, some aspects the current guidelines require further examination.

Efficacy threshold and confidence limits: dogs and cats APVMA requires an efficacy of .95% for the registration of a claim against I. holocyclus, presumably because of the pathogenicity of intoxication. However, point estimates of efficacy derived from a sample provide a relatively uninformative measure of the true population parameter. Using a confidence interval (95% or 99%) provides additional useful limits, which aids in the interpretation of data that may encompass the true but unknown population mean. This issue is exacerbated by the pathogenicity of the ticks as untreated na¨ıve animals are exposed to the real risk of potentially fatal paralysis. APVMA therefore requires a minimum of only five ticks per animal in trials with at least eight animals per group. Using these criteria could allow a product to be registered if only one tick was found on one treated dog. The mean reduction is 97.5% even though the lower 95% confidence interval is 87.1%. Registration is also possible if one tick was found on two treated dogs, resulting in a mean efficacy of 95% and a lower 95% confidence interval of 83.4%. The use of immunized animals provides a superior model that allows much higher levels of infestation while at the same time reducing the risk of paralysis. The higher infestation rate provides more robust data, which could in turn allow a reduction in the number of animals required for testing, but at the same time providing a tighter estimate of the true but unknowable efficacy of the test product. It has been estimated that more than 60 dogs infested with 5 ticks would be required to attain equivalent confidence intervals to a group of 8 dogs with a mean of 30 ticks per untreated animal. There are therefore regulatory and animal welfare advantages to using immune animals to determine efficacy. There is no evidence that immunity against holocyclotoxin impacts tick attachment, feeding, or survival. It is therefore suggested that efficacy be tested on immune animals as this will reduce the risk of paralysis while at the same time generating more robust data for review by the APVMA. It is noted that the WAAVP recommends infestation with B50 ticks per animal and that at least 20% should become attached. This attachment rate may be too low to generate narrow confidence intervals, and consideration should be given to defining challenges based on being able to attain efficacy in excess of a threshold lower 95% confidence interval rather than the mean. The author (M.W.) does not consider it possible to distribute five ticks over an animal and reflect the natural distribution of 80% of ticks attaching to the head and neck, this leaves only one tick to be placed elsewhere.

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Efficacy threshold and confidence limits: calves WAAVP guidelines for evaluating the efficacy of acaricides against ticks on ruminants (see Appendix A of Ref. [23]) in respect of paralysis ticks need revising. The guidelines suggest mean efficacy should be .95% and that no individual calf should have ,90% reduction in tick burden after treatment. It is not possible to immunize calves as they develop local inflammatory reactions after repeated infestations, therefore the number of ticks must be kept low to avoid paralysis. Typically, calves are infested by attaching 10
15 ticks. A single live tick found on a treated calf infested with 10 ticks could render a claim inadmissible for the regulator, which essentially increases the threshold to 100%. Similarly, persistent activity is defined as the period following treatment for which no attached tick survives for longer than 3 days on any individual treated calf, that is, the period for which efficacy is maintained at 100%. There is a need for a statistically valid definition of efficacy to be developed, which recognizes the interaction between tick numbers and animal numbers in determining the confidence interval around the estimated mean. There is also a need for a valid risk
benefit analysis that evaluates the benefits that might accrue from using less efficacious products that are prevented from being marketed due to the very high barriers that have been put in place. Dose-confirmation studies: dogs and cats APVMA requires two dose-confirmation studies be conducted and that the “studies should include groups of animals infested with ticks collected both from Far Northern Queensland and from regions representative of the rest of the geographic range of Ixodes holocyclus.” There is no evidence that I. holocyclus from different localities have differences in susceptibility to acaricides. Currently, the only place where large numbers of ticks are collected is the Northern Rivers region of NSW. This is because this is the site of tick serum producers and the contract research organization that conducts tick efficacy research. Note that .10,000 ticks may be required for a single study. The author (M.W.) has attempted to establish infrastructure required to collect ticks from Far Northern Queensland since 2003, including commissioning an entomologist from James Cook University to find ticks around Cairns, but has not been able to collect ticks in sufficient numbers to allow comparison of efficacy. APVMA does not require definition of comparative efficacy, rather that ticks be included in the challenge. The value of such an approach is questionable as there is no way of determining the source of any surviving ticks. An alternative approach to assessing possible geographic variation in acaricidal susceptibility is discussed later. I. holocyclus is found in a range along the eastern seaboard from Port Douglas to Victoria, with unconfirmed reports from Tasmania. It is

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submitted that it is simply not possible to collect ticks from such disparate locations in sufficient numbers to conduct efficacy trials. Confirmation that efficacy is maintained (or more correctly that inefficacy does not occur) under field conditions is better assessed using a well-designed multisite clinical (field) study with cases enrolled from representative geographic locations from the parasite’s ecological range. Due to obvious ethical constraints, such a study would need to have a positive control and no negative control and could only be considered after efficacy had been confirmed in a rigorous pen trial(s).

Acaricidal evaluation: in vivo testing of Ixodes holocyclus on dogs Dogs are infested by either inducing the ticks to embed into the host (attachment method) or releasing ticks onto the head and neck and along dorsal spine of dogs allowing the ticks to embed naturally (release method). The attachment method is used when the IVP has no repellency or when it acts systemically in comparison to being absorbed through the tick’s cuticle. The release method is used in cases where the IVP is repellent or must be absorbed from residual chemical on the animal’s skin. Foxhound and foxhound cross dogs (1
10 years old) are immunized against holocyclotoxin using a modification of the method of Stone et al.24 After immunization, dogs can tolerate a challenge of 30
50 attached ticks with no evidence of intoxication. Dogs are infested by attaching or releasing 30
50 female adult unfed I. holocyclus ticks. Ticks are individually attached to each dog predominately on the head, shoulders, and dorsal midline to simulate the natural distribution of attached ticks. If the release method is being used, the ticks are released on the head, neck, and dorsal midline. A tick carrying capacity test is conducted prior to treatment to select dogs that can carry high numbers of ticks without being affected. Dogs are ranked by their tick carrying capacity and randomly allocated to control or treatment groups. Ticks are attached 24 h prior to treatment and assessed at intervals of up to 72 h PT or reinfestation to determine efficacy and persistence. It is important that ticks are killed well within the critical period of 72 h after attachment before paralysis begins to set in. Dogs are housed individually during tick challenges. Trained personnel conducts all tick counts to ensure a standardized technique during assessments. Tick counting personnel wears disposable overalls during infestation and counting to reduce the chance of being infested and to eliminate chemical transfer between dogs. Working with I. holocyclus poses a significant health and safety risk for study personnel. Staff can become infected with rickettsia and other vector-borne disease and develop hypersensitivity to components of tick salvia or mammalian meat

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TABLE 4C.1 Parameters used to describe observed Ixodes holocyclus ticks at each assessment for efficacy of treatment.25 Parameter

Choices

Comments

Viability

Live, dead

Live ticks show leg movement, no crenation, possible engorgement, presence of tick feces, and/or attachment site inflammation. Dead ticks have no leg movement, no reaction when stimulated, and possible crenation.

Attachment

Attached, free

Attached ticks have their hypostome embedded into the dog’s skin and are not easily dislodged. Free ticks are either live and moving through the coat, or dead and sitting in the hair.

Feeding

Unengorged, partially engorged, or fully engorged

Unengorged ticks show no swelling or evidence of blood ingestion; engorged ticks are 12
15 mm long, 8
10 mm wide and have a turgid appearance. Engorged ticks are typically not seen before 6 days postinfestation and are not observed in efficacy trials because ticks were removed at 72 h PI. Engorging ticks show a conspicuous swelling of the alloscutum and appear wider and longer than unengorged ticks. These ticks contain blood or digested blood.

allergy. Chemical transfer is controlled by washing hands and changing overalls between dogs. The whole body of the dog is searched visually and by palpation until no more ticks can be found. The 24 and 48 h assessments are conducted without removing ticks, while at the 72 h assessments, any remaining ticks are assessed, removed, and discarded. All dogs are searched prior to each infestation to identify and remove ticks that had migrated from the environment and attached subsequent to the previous infestation period or that had been missed during the previous tick assessment. Each tick observed during the count is described according to selected parameters (Table 4c.1).25 Some ticks are classified as “dead” when examined in situ but display uncoordinated agonal leg movement after removal at 72 h and are then reclassified as “moribund.” These ticks typically exhibit reduced engorgement and are smaller than live ticks of the same age with evidence of mild crenation (slightly shriveled appearance). These ticks should be included in the total dead tick count as they are not capable of

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TABLE 4C.2 Ixodes holocyclus tick category assignments used to assess the parasiticidal effect of treatment. Category

Observation/survival and attachment status

Parasiticide effect

1

Live, free, attached

Not demonstrated

2

Live, attached, unengorged

Not demonstrated

3

Live, attached, and engorging or engorged

Not demonstrated

4

Dead, free

Yes

5

Dead, attached, unengorged

Yes

6

Dead, attached, engorging

Yes

7

Dead, attached, engorged

Not demonstrated

Source: Adapted from Marchiondo AA, Holdsworth PA, Green P, Blagburn BL, Jacobs DE. World Association for the Advancement of Veterinary Parasitology (WAAVP): guidelines for evaluating the efficacy of parasiticides for the treatment, prevention and control of flea and tick infestation on dogs and cats. Vet Parasitol 2007;145:332
344.

causing paralysis, but the APVMA requires such ticks to be included in the total live count. The recorded description of each tick is then used to place it in one of seven categories as adapted from Marchiondo et al.19 and utilized by Smith et al.,26 and Fisara and Webster25 (Table 4c.2). Category 6 (dead, attached, engorging) was added to the original list of Marchiondo et al.19 At each counting period the total number of ticks counted on each dog that were assigned to categories 1, 2, 3, and 7 were used in the calculation of the results. Marchiondo et al.20 have dropped the use of engorgement in the categorization of ticks by thumb or comb counts. Refer to this guideline for the survival status, attachment status and interpretation for tick efficacy assessed against topically, nonsystemic, and systemically active IVPs. Treatment efficacy is calculated using arithmetic and geometric means of log transformed counts 1 1 as follows: Treatment efficacy 5 100 3

Control group mean
Treated group mean Control group mean

Speed of kill studies have also been conducted with adult female I. holocyclus.27 Dogs are infested 24 h prior to treatment, examined with ticks classified and counted 8, 12, 24, and 48 h PT, and at 12, 24, and 48 h after subsequent reinfestations. Efficacy is determined at each time point relative to counts for placebo dogs based on mean counts.

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Acaricidal evaluation: in vivo testing of Ixodes holocyclus on cats Conducting trials in cats is more difficult that in dogs. Cats remove ticks by grooming, are smaller therefore not amenable to being infested with large numbers of ticks, and are more difficult to immunize than dogs. Finding ticks is more difficult in cats due to the fine hair masking the tick’s presence during palpation and visual searches. Cats may be more vulnerable to holocyclotoxin than dogs due to their smaller size so careful monitoring is required to minimize losses from paralysis. Immunized cats can host 10 ticks for 72 h with no ill effect and appear to respond better to treatment with tick antitoxin serum, if they are paralyzed, than na¨ıve cats. Efficacy studies of I. holocyclus infested cats are conducted in a similar way to canine studies. Cats are infested by manually attaching 10 ticks predominantly on the head, shoulders, and dorsal midline, to simulate the ticks’ natural predisposition for these areas and to prevent the cats from grooming ticks off. Cats are infested 1 week prior to treatment to conduct a tick carrying capacity test for allocation them 24 h prior to treatment to determine against preattached ticks and at weekly intervals to determine the test items persistency. Tick counts begin with a search of the areas to which the ticks were attached at the time of infestation. This is followed by a whole-body search to locate any ticks that may have migrated away from the tick application site. For all study procedures the control cats are handled first to avoid any potential transfer of product from treated to untreated cats. Ticks are counted and assessed at 24 and 48 h PT and at 24, 48, and 72 h after each subsequent infestation. The 24 h PT and 24 and 48 h post reinfestation assessments are completed without removing the ticks. Following the 48 h PT and the 72-h postinfestation assessments all ticks are removed and discarded. The ticks are classified as adapted from Marchiondo et al.19 and Fisara and Webster.25 The assessment of efficacy is based on the number of category 1, 2, 3, and 7 ticks found on each cat at 48 h PT and at 72 h after each tick reinfestation. Repellency testing of Ixodes holocyclus on dogs Repellency of I. holocyclus is a highly desirable attribute for any parasiticide as a bite from a single tick can be fatal. Ixodid ticks (such as I. holocyclus) acquire their hosts by climbing vegetation and carrying out questing behavior. They detect the host by a range of stimuli including vibrations, CO2 concentrations, and host odor.28 Due to this complex parasite
host interaction and vector role of ticks, the total benefit to the host of an acaricide cannot be completely described by a single efficacy value.20 Acaricides are effective in controlling ticks in two main ways, either solely, or in combination. Repellency sensu stricto refers to an irritant effect that causes the tick to move away from the treated animal, leading to

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avoidance, or to fall off the animal soon after contact with the hair coat.29 This is true repellency, as ticks are removed/killed before they can bite. Repellency sensu lato, or “expellency” refers to the combined effect of disruption of attachment and causing the tick to fall off the host within 24 h.29 The term “killing effect” is used to describe the ability of an acaricide to induce death of the tick. This term can also be interchanged with overall efficacy of an acaricide. To test the repellency of parasiticides against I. holocyclus in vivo, hyperimmunized dogs are sedated with medetomidine hydrochloride and placed in large plastic, uncovered infestation chambers with double-sided tape placed around the top perimeter of the walls to prevent ticks from escaping.30 The temperature of the pens containing the chambers is kept at 18 C. Each dog is placed centrally, in lateral or sternal recumbency in the infestation chamber. Ticks (50) are released alongside the dog within 20 cm of the dog, 10 cm either side of the head, 10 cm either side of the forequarters, and 5 cm either side of the hindquarters. Dogs remain in the chambers for 1 h 6 5 min, before removing them to their pens being careful not to dislodge any ticks. Ticks remaining in the infestation chambers are counted and classified after the dogs are removed following the 1 h tick challenges. These counts are used to calculate the repellency effect sensu stricto.20 Ticks on the dogs are counted at 6 h 6 30 min and 24 6 1 h postchallenge and again 72 h postchallenge as a safety check for previous undiscovered ticks. Dogs are systematically examined for ticks using digital palpation and visual inspection. Ticks are classified as adapted from Marchiondo et al.19 (Table 4C.2). Classification is a subjective process undertaken by experienced tick assessors. The total attached tick (ToA) count consists of all attached live and dead ticks found on the dogs at 6 and 24 h postchallenge.30 The repellency effect sensu lato (or expellency) is calculated by comparison of treated and untreated group mean ToA at 6 and 24 h postchallenge.20 The total live tick (ToL) count was also performed at 6 and 24 h postchallenge. This count includes all live attached or unattached ticks and gives an estimate of the killing effect, or overall efficacy, of the treatment. In order to determine the repellency effect, group arithmetic mean tick counts for residual ticks in the infestation chambers (sensu stricto), and group geometric mean tick counts of ToA at the postchallenge counts at 6 and 24 h (sensu lato or expellency), are calculated.29 Repellency effect sensu stricto (%) 5 [(Mean Residual Tick Treatment Count)/(Mean Placebo Count)]/Mean Treatment Count Repellency effect sensu lato (%) 5 Mean ToA Tick Placebo Count 2 Mean Treatment Count/Mean Placebo Count Overall efficacy was calculated using geometric mean ToL counts at 6 and 24 h after each tick challenge.

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Further methods on testing repellents and attractants on tick species, which may be applicable to I. holocyclus, are provided by Bissinger and Roe.31

References 1. Barker SC, Walker AR. Ticks of Australia. The species that infest domestic animals and humans. Zootaxa 2014;3816:100
12. 2. Seddon HR. Diseases of domestic animals in Australia, Part 3. Arthropod infestations (ticks and mites): together with a section on animals, insects and other agents harmful to stock. 2nd ed. Canberra: Commonwealth of Australia Department of Health; 1968. p. 170. 3. Doube BM. Cattle and the paralysis tick Ixodes holocyclus. Aust Vet J 1975;51(11):511
15. 4. Ross IC. The bionomics of Ixodes holocyclus Neumann, with a redescription of the adult and nymphal stages and a description of the larvae. Parasitology 1924;16(4):365
81. 5. Doube BM. Seasonal patterns of abundance and host relationships of the Australian paralysis tick, Ixodes holocyclus Neumann (Acarina: Ixodidae), in southeastern Queensland. Aust J Ecol 1979;4(4):345
60. 6. Bagnall BG, Doube BM. The Australian paralysis tick Ixodes holocyclus. Aus Vet J 1975;51(3):159
60. 7. Wall R, Shearer D. Veterinary ectoparasites. Biology, pathology and control Chapter 3 Ticks (Acari). 2nd ed Oxford, UK: Blackwell Scientific; 1997. p. 70. 8. Oxer DT, Ricardo CL. Notes on the biology, toxicity and breeding of Ixodes holocyclus (Neumann). Aus Vet J 1942;18(5):194
9. 9. Doube BM, Kemp DH, Bird PE. Paralysis of calves by the tick, Ixodes holocyclus. Aust Vet J 1977;53(1):39
43. 10. Bootes BW. A fatal paralysis in foals from Ixodes holocyclus Neumann infestation. Aust Vet J 1963;38(2):68
9. 11. Stone BF, Binnington KC, Gauci M, Aylward JH. Tick/host interactions for Ixodes holocyclus: role, effects, biosynthesis and nature of its toxic and allergenic oral secretions. Exp Appl Acarol 1989;7(1):59
69. 12. Eppleston KR, Kelman M, Ward MP. Distribution, seasonality and risk factors for tick paralysis in Australian dogs and cats. Vet Parasitol 2013;196:460
8. 13. Kaire GH. Isolation of tick paralysis toxin from Ixodes holocyclus. Toxicon 1966;4 (2):91
7. 14. Iikiw JE, Turner DM, Howlett CR. Infestation in the dog by the paralysis tick Ixodes holocyclus. 1. Clinical and histological findings. Aust Vet J 1987;64(5):137
9. 15. Sutherland SK, Tibballs J. Australian animal toxins. The creatures, their toxins and care of the poisoned patient. Melbourne: Oxford University Press; 2001. p. 467
88. 16. Goodrich BS, Murray AJ, Holmes PR. The establishment of a laboratory colony of Ixodes holocyclus. Aust Vet J 1991;54:490
3. 17. Levin ML, Schumacher LBM. Manual for maintenance of multi-host ixodid ticks in the laboratory. Exp Appl Acarol 2016;70:343
67. 18. Stone BF, Commins MA, Kemp DH. Artificial feeding of the Australian paralysis tick, Ixodes holocyclus and collection of paralyzing toxin. Int J Parasitol 1983;13 (5):47
454. 19. Marchiondo AA, Holdsworth PA, Green P, Blagburn BL, Jacobs DE. World Association for the Advancement of Veterinary Parasitology (WAAVP): guidelines for evaluating the

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20.

21.

22.

23.

24. 25.

26.

27.

28.

29. 30.

31.

Parasiticide Screening, Vol 1 efficacy of parasiticides for the treatment, prevention and control of flea and tick infestation on dogs and cats. Vet Parasitol 2007;145:332
44. Marchiondo AA, Holdsworth PA, Fourie LJ, Rugg D, Hellmann K, Snyder DE, et al. World Association for the Advancement of Veterinary Parasitology (W.A.A.V.P.) second edition: guidelines for evaluating the efficacy of parasiticides for the treatment, prevention and control of flea and tick infestations on dogs and cats. Vet Parasitol 2013;194 (1):84
97. Australian Pesticides & Veterinary Medicines Authority (APVMA). Preamble for the WAAVP guideline for fleas and ticks on dogs and cats. ,https://apvma.gov.au/node/ 1040.. International Cooperation on Harmonisation of Technical Requirements for Registration of Veterinary Medicinal Products (VICH), June 2000, VICH Guideline 9: Good Clinical Practice (GCP). https://vichsec.org/guidelines/pharmaceuticals/pharma-efficacy/good-clinical-practice.html. Holdsworth PA, Kemp D, Green P, Peter RJ, De Bruin C, Jonsson NN, et al. World Association for the Advancement of Veterinary Parasitology (W.A.A.V.P.) guidelines for evaluating the efficacy of acaricides against ticks (Ixodidae) on ruminants. Vet Parasitol 2006;136:29
43. Stone BF, Neish AL, Wright IG. Tick (Ixodes holocyclus) paralysis in the dog
quantitative studies on immunity following artificial infestation with the tick. Aust Vet J 1983;60:6
68. Fisara P, Webster M. A randomized controlled trial of the efficacy of orally administrated fluralaner (Bravectot) against induced Ixodes holocyclus (Australian paralysis tick) infestations on dogs. Parasit Vectors 2015;8:257. Smith WM, Ahlstrom LA, Rees R. Long-term efficacy of an imidacloprid 10%/Flumethrin 4.5% polymer matrix collar (Serestos, Bayer) against the Australian paralysis tick (Ixodes holocyclus) on dogs. Parasitol Res 2013;112:S1
10. Packianathan R, Hodge A, Brueeke N, Davies K, Maeder S. Comparative speed of kill of sarolaner (Simparicas) and afoxolaner (NexGards) against induced infestations of Ixodes holocyclus on dogs. Parasit Vectors 2017;10:98 Available from: https://doi.org/10.1186/ s13071-017-2024-9. Padula AM. Tick paralysis of animals in Australia. In: Gopalakrishnakone P, Faiz SMA, Gnanathasan CA, Habib AG, Fernando R, Yang C, et al., editors. Clinical toxicology. 2015. p. 1
20. 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. de Burgh S, Hunter K, Jackson C, Chambers M, Klupiec C, Smith V. Repellency effect of an imidacloprid/flumethrin (Serestos) controlled release polymer matrix collar against the Australian paralysis tick (Ixodes holocyclus) in dogs. Parasitol Res 2017;116:145
56. Bissinger BW, Roe EM. Tick repellents: past, present, and future. Pestic Biochem Phys 2010;96:63
79.

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Chapter 4d

Arachnida, Mesostigmata, Dermanyssidae, Macronyssidae, Varroidae Alan A. Marchiondo, MS, PhD Adobe Veterinary Parasitology Consulting LLC, Santa Fe, NM, United States

Arachnida Mesostigmata Dermanyssidae

Dermanyssidae Dermanyssus gallinae—De Geer, 1778—red poultry mite or chicken mite Biology and life cycle Dermanyssus gallinae is a nonburrowing, bloodsucking mite that parasitizes poultry and other bird species.1 It occasionally infests cats2 and can bite humans, dogs, rodents, rabbits, and horses. Adult mites feed at night, particularly around the breast and legs of hens, and can remove enough blood to cause anemia, kill nestlings, and reduce egg production. They hide in nests, roosts, and other concealed locations in poultry houses. These mites have a single dorsal plate, a sternal plate with two pair of setae, and an anal plate with the anus positioned in the posterior half. The chelicerae of adult mites are long, slender, and whip like. Chelae are very small, minute in size. The mite develops through five life stages. Female mites can oviposit within 24 h of their first blood meal and measure about 1 mm after engorgement.3 Eggs are deposited in the diurnal hiding places that hatch to six-legged, nonfeeding larvae. These develop to the blood-feeding protonymph and deutonymph, and finally to adult male and female mites. The life cycle can be completed within 1 week or up to 5 months. Adult mites can survive starvation for up to 34 weeks in an empty poultry house. Rearing method(s) Laboratory colonies of D. gallinae are started by collecting mites (several hundred to several thousand) from a poultry breeding farm. The colony is maintained according to the methods of Chamberlain and Sikes4 in tall metal popcorn cans surrounded by creosote. Well-fitted lids, with centers cutout

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and covered with fine mesh cloth, are sealed onto the cans with Parafilm. Two or three pieces of corrugated cardboard (4 3 6 in.) is wedged between dried grass (Bermuda or Johnson grass autoclaved and packed in the bottom of the can to a depth of B3 in.) and the side of the can to provide suitable cracks for mite to hide and oviposit. A sheet metal tray is attached on each end of the can with adhesive tape at a height to allow its bottom to rest on the grass. The tray is 8.5 in. long 3 1.75 in. deep 3 1.5 in. wide at the bottom and 1.25 in. wide at the top (that successfully discourages perching by young chicks) with the back of each container extending 1.5 in. higher to facilitate taping to the end of the can. One tray is used for water and the other for sterilized chick mash. A chick introduced into a can one to two nights a week maintain the colony with the mites feeding rapidly and then hiding under a small wooden platform on the bottom. When mites are needed for study purposes, they are brought out of hiding by removing the chick for 3
4 days. They are then aspirated from the corners and rim of the can, or one of the cardboard pieces can be moved and the mites resting on it collected. The colony is kept at 26.7 C
28.3 C. D. gallinae mites have been feed in vitro using bird skin and blood at 40 C
41 C with molting and oviposition of the mites, but the completion of the life cycle was not maintained in vitro.5 Mites can also be fed using an artificial feeding device with laboratory maintenance of a colony in vitro.6 Mites are collected from litter of infested poultry houses and separated from the litter with a suction pump and counter. After identification, mites are stored in glass vials at 30 C and 60%
95% RH in total darkness for at least 1 week prior to studies to eliminate the effects of any previous blood meals and egg laying. An in vitro feeding device consists of a Pasteur pipette and a blood reservoir separated by an artificial membrane. The main chamber of the pipette containing the mites is fitted with a folded piece of filter paper, utilizing an artificial feeding device. Mites are sucked into the chamber by fitting the larger diameter of the pipette with a filter and tubing attached to a suction pump. Once mites are in the chamber, the larger opening is covered with a filter tip and the smaller opening is sealed with Parafilm M. During blood meals, a 2 3 2 cm2 membrane is stretched over the large opening and held tightly in place with a 1 mL pipette tip serving as the blood reservoir. Biological membranes are prepared from the skin of adult chickens, 1-dayold chicks, turkey hens, or nude mice. Skin samples are scraped free of fatty SC tissue, rinsed in 0.09% saline solution, dried with paper towels, and either stored at 4 C or frozen at 220 C. Biological membranes are placed on the feeding device so that the external side faces the mite chamber. Parafilm membranes are stretched to 10 times their original size. Red mite feeding preferences of the synthetic membranes are Parafilm M and a commercial skin substitute (pansement hydrocolloide 0.3 mm). Thirty mites are placed in the feeding chamber. Blood is collected from the occipital sinus of live chickens, mixed with an anticoagulant (EDTA, heparin, or trisodium citrate),

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and either stored at 4 C or frozen at 220 C. A colony of D. gallinae mites was maintained in vitro for seven generations using the artificial feeding device with 1-day-old chick skin and heparinized (0.02 mM/mL) chicken blood at 30 C and 60%
95% RH in total darkness. Another in vitro feeding device has been developed and tested against an in vivo feeding device for the rapid screening of D. gallinae antigens with comparable results.7

In vitro method(s) Contact—filter paper—toxicity A precise determination of acaricidal toxicity can be obtained by the residue contact method8 wherein 10 female mites are placed between two filter paper circles with dried residues of a test compound (µg/cm2) in each of two Petri dish halves.8 Serial dilutions of the test compound(s) are prepared on filter paper disks, and each concentration is tested in 3
4 replicates. The dishes, containing the exposed mites are placed on wet paper towels to provide humidity and held at 27 C for 24 h, at which time mortality counts are determined. Lethal dose concentrations are calculated for the test compound(s). An in vitro filter paper assay has been developed wherein mites are confined on treated, molded filer papers held in position by two glass sheets to form an escape-proof exposure chamber.9,10 Contact—filter paper—fumigation A filter paper contact bioassay can be used to evaluate the toxicity of test compounds.11 Serial dilutions of test compounds in ethanol or a suitable solvent are applied in a volume of 50 µL onto Whatman No. 2 filter papers (4.25 cm diameter) and allowed to dry in a fume hood for 3 min prior to use. The treated and untreated filter papers are added to 28 mL glass vials with screw caps and 23 mites are transferred to the vials using a pooter (aspirator). The vials are sealed with a layer of cling film and a further layer of Parafilm before adding the screw cap. The vials are stored for 24 h at 22 C with a 16L:8D hour photoperiod and assesses for mite mortality with LD values determined for the test compounds. A test for fumigant toxicity consists of treated filter papers with the required amount of test compound to treat a 1400 mL glass vessel at the mg/cm2 concentration equal to the LD50 value of the test compound in the contact toxicity assay. Treated and untreated filter papers are added to 28 mL glass vials along with 23 mites. The vials are either closed or left open with a 0.2 mm2 mesh for air exchange and stored for 24 h at 22 C with a 16L:8D hour photoperiod. Percentage mite mortality was determined with a higher mortality of mites in the closed vials versus the open vials indicating a fumigant- or vapor-phase effect. A fumigant bioassay has been developed using treated filter paper placed on the lid of a disposable plastic Petri dish that has a center hole (3.0 cm diameter) screened with metal mesh.12
14 Groups of 20
30 adult mites are

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put into the Petri dish and covered with the lid with the treated filter paper. The dishes are then sealed with another lid and wrapped with Parafilm and incubated for 24 h at 26 C 6 1 C and 55% 6 5% RH under a 16L:8D hour cycle. This system prevents direct contact of adult mites with the test compound. Mite mortality is determined at 24 h postincubation. Contact—filter paper—susceptibility and resistance The resistance of D. gallinae to acaricides is said to resemble the resistance of ticks to acaricides, particularly pyrethroids resistance in Boophilus microplus, with LD50 values of susceptible mites and the cattle tick and resistance factors for pyrethroids being similar.15
17 Therefore an in vitro modified LPT (FAO No. 7) designed for tick resistance testing18 can be used to test acaricidal resistance in D. gallinae. Whatman filter papers (80 3 75 mm) are impregnated with 0.3 mL of dilutions of acaricides in 0.3 mL trichloroethylene in olive oil. Two replicates of treated and control papers (solvent only) are air-dried for 2 h. Adult or nymph mites (50
100) are placed on the papers that are folded over and closed with clips. The papers are incubated at 25 C and 85% RH for 24 h. Mite mortality is determined within 7 days and lethal concentrations of acaricides determined. Filter paper contact bioassays are typically used to evaluate the toxicity and persistence of toxic effects of test compounds against D. gallinae. Persistency is tested after 15 and 30 days exposure of mites on treated filter paper introduced into Pasteur pipettes with one end covered with Parafilm and the other end plugged with cotton.19 Mite mortality is determined after the exposure period, and the percent efficacy of test compounds are determined versus the control. In vitro barrier treatments applied to the center of a 90 mm Petri dish divided into three areas: prebarrier, barrier, and postbarrier and allow for the evaluation of commercial barrier products, double-sided sticky tape, acaricidal oils, insect barrier glue, petroleum jelly, and detergent.20 Mites are placed in barrier dishes that are placed in 250 mL square weigh boats filled with oil to prevent mite escape. Dishes are left at RT for B24 or B72 h under a 6L:18D hour photo cycle. After exposure the dishes at placed at 220 C for 3 h and the mites in each on the plate are counted. Contact—glass surface—toxicity and persistency A self-dosing glass contact method to screening acaricides against D. gallinae involves the use of straight thin tubes with an internal surface area of 10 cm2 on which a defined concentration (µg/cm2) is applied.21
25 Dilutions of acaricides in acetone are pipetted into tubes that are automatically rotated in a horizontal position in a flow of warm air to yield an evenly distributed deposit. Twenty female adult mites are inspirated into the treated and control tubes with a vacuum pump. Each tube is sealed with nylon tissue, and the mites

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exposed for 24 h at 25 C and 90% RH. After the 24 h exposure the mites are transferred to clean recovery tubes and stored under the same conditions. Percentage mortality is recorded after 24 h. Mites are considered dead if they neither responded to touch nor displayed normal movement. Contact—feather surface—toxicity A feather test has been used to determine effective concentrations of parasiticides rather the pipette technique.25,26 Feathers containing mites and their excrement are removed from infested hens, held over a laboratory sink, and sprayed individually with 2.5 mL of dilute (0.005% or 0.05%) water emulsion of parasiticide or water as a control using a DeVilbiss atomizer. The treated feathers are dried for 15 min and placed in narrow-gauge, screen-capped plastic containers held in a desiccator at RT and 100% RH. Treated and control feathers are examined before and after treatment at 0, 12, 24, 48, 72, and 96 h. The number of live mites (typical movement on feathers) is counted and the percent efficacy determined. Repellency Filter paper contact bioassays can be used to evaluate the repellent effect of test compounds against D. gallinae using a video camera movement of mites exposed to test compounds for 30 min.19 A circular rubber rings (4.5 cm diameter) is placed onto a filter paper cut to the same size, and both are placed between two pieces of glass. On each filter paper, two circles (0.81 cm diameter) are previously drawn at two opposite points on the ring. A dilution of test compound or a negative control (nothing or 2 µL of commercial alimentary oil) is placed in the center of one of these circles, and 2 µL of commercial alimentary oil placed in the center of the remaining circle as a control. Prior to mounting the apparatus, filter papers are dried under a fume hood, and a single mite is then inserted just before the test. The video camera records the trajectory of mite movement for 30 min, and the area avoided by the mite is measured. Five replicates of each test compound are performed. Mites that rested for 15 min are excluded. The repellent effect is calculated as a percentage of the area avoided based on the measured area and surface of the ring. Repellency assays of test compounds against D. gallinae can be conducted on a system of four olfactometers (10 mm internal diameter, two main arms each 100 mm, and third arm of 10 mm).11 Experimenters are advised to refer to the design provided in the publication. Air pumped through each arm of the Y-tube of the olfactometer at 1 L/min is passed through a two-stage filtration process (activated charcoal and particulate filter in-line PTFE 50 mm diameter 3 0.2 µm) prior to being humidified by bubbling through distilled water. Humidified air flow is divided equally by a custom-made glass Y-tube allowing each air stream to be passed through a 250 mL conical flask. Into one of these flasks, 19.21 µL of the pure test

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compound (or distilled water for the control) is added at an initial concentration (mg oil/cm3) of flask volume. Test compounds are added to flasks on a Whatman No. 2 filter paper (4.25 cm diameter). Into the other flask, distilled water is added at the same concentration as a control. Air from these flasks is then passed down opposing arms of a further custom-made glass Y-tube, in which B40 adult female mites are confined during the duration of the experiment by securing 0.2 mm mesh over all openings. The distribution of mites in the two opposing arms of the Y-tube is then recorded at this time (t0 1 15 min) and again at t0 1 30 min and t0 1 45 min. Following this assessment, the test compounds are added to the flasks as necessary, and the distribution of mites recorded again 15, 30, and 45 min later and every 24 h for a period of 13 days. The proportion of D. gallinae in the untreated arm of the Y-tube of olfactometers (number in untreated arm/total mites) is calculated for each test compound treatment on each sampling occasion. Biological control—predator bioassay Two or no predators (soil mite species likely to prey on D. gallinae in a commercial setting) are placed in separate glass Petri dishes (48 mm diameter 3 12 mm height) along with 10 adult mites.27 Dishes with the addition of a moistened tissue paper to maintain the inner humidity at 80%
90% RH are sealed with a thin plastic film and incubated at 22 C for 48 h in complete darkness. Mite mortality is then determined where mites are deemed dead if they exhibit no movement following agitation with an entomological pin. Predation of D. gallinae of each treatment is expressed as the corrected % mortality.

In vivo method(s) Individual bird treatment for D. gallinae infestations are conducted using hand and power sprayers.26 A volume of 18 mL/s at 30 psi is applied to the vent for 2.5 s (B50 mL/hen, 3.8 mL/hen is the recommended normal volume). The effectiveness of treatments (mite infestation ratings) is determined using the 0
4 index system28; a 5 rating (200,000 mites) may be included for high-density mite infestations on untreated hens. Mite infestation ratings are conducted by examining the vent feathers for 1
17 weeks PT. Direct bird treatments against D. gallinae can also be conducted in community-type cages with wooden frames by means of a power-driven sprayer. Treatment typically involves the premise rather than individual birds. Spraying with acaricides and dusting with silicas are the primary methods of red mite control.20 However, acaricidal resistance against D. gallinae has been reported.15,29 Acaricidal resistance studies have been conducted using selected poultry farms experiencing problems with the control of D. gallinae.15 Engorged mites (2000 females, males, and nymphs) are collected from each

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of the problem farms and one control farm with no mite control problems. The mites are tested in a modified LPT to determine resistance.15 There is no established guideline for the assessment of mite population density in a poultry house. However, traps made of rectangular pieces of 3 mm thick corrugated cardboard or Bristol board can be used to as a sampling device in poultry house to evaluate D. gallinae.30,31 Trap sizes of 140 3 100 or 110 3 70 mm used for 6
10 days or 2 days, respectively, collected significantly more mites per day than other size traps and sampling duration periods. These boards can be used to assess mite density in a poultry house. This method of assessing mite populations serves as an effective method to evaluate acaricidal treatments of poultry facilities.32 A poultry house can be artificially infested with D. gallinae by seeding a group of stocked cages in a house empty for some time.33 The mites (B2000) are collected from a free-range unit and placed in a 250 mL plastic screw-cap chambers with a mesh lid for air flow. The mites are released into stocked cages B72 h after being collected by securing one chamber to the egg tray with the mesh severed to release the mites. Weekly monitoring of mite populations are conducted using traps. After about 28 days with the establishment of the mite population, the cages can be treated with a test acaricide by a pressure handheld lance sprayer to the point of runoff. The mite populations are then monitored on TD 2, 7, 14, 21, and 28 PT using traps. Permethrin-impregnated plastic strips34 attached in the hen housing system whereby birds rub against the strips or attached out of reach of birds (perches and egg-belt lids) have been tested and monitored using trapes or by collecting mites with a semitransparent plastic tape.35 An automated mite counter36,37 has been developed to monitor D. gallinae populations in layer production systems. The device consists of an opening tube (B1.0
1.5 mm diameter) in a body container comprising a casing and a lid closing the casing, a receiving section, a sensor device for counting passing mites, an electronic processor connected to a tube leading to a filter, a removal device using air suction, and a power and data cable. Mites enter the counter and are detected by the senor device. The in vitro sensitivity of the device is the detection of 100% of adults, 97% of nymphs and larvae. The device is able to track and detect D. gallinae population growth in layer facilities. Fungal spore suspensions and other biological control methods can be tested in commercial layer farms with infestations of D. gallinae.38 Oil suspensions of fungal concentrations with affinity to feather and cage surfaces are sprayed on hens and cages using portable sprayers with coarse spray droplets. Distilled water is sprayed on hens and cages in the control group. Mite populations are counted using traps35 prior to spraying and at 1
4 weeks postspraying. Mite populations are determined for the treated and control groups.39

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Macronyssidae Ornithonyssus sylviarum—Canestrini & Fanzago, 1877—the northern fowl mite Biology and life cycle The northern fowl mite parasitizes domestic and wild birds throughout the temperate regions of the world causing economic damage to poultry production in the form of anemia, decreased egg production by 10%
15%, negative effects on interior egg quality and hen integument, reduced weight gain, reduced feed conversion efficiency of 5%
7%, and even death.40
43 Newly infested chickens can have mite populations in excess of 20,000 mites/bird in 9
10 weeks. Infestations don’t usually occur until the bird reaches sexual maturity and birds older than 40 weeks usually don’t support infestations. Humans can also be bitten when handling eggs from infested laying hens. The octopod adult mites measure 0.5 mm and appear reddish brown in color after taking a blood meal. Adult mites congregate on the vent, tail, back, and legs of birds. Female mites lay eggs (2
5/female) after each blood meal. Eggs hatch in 1
2 days, but the six-legged hatched larvae do not feed and molt in B8
9 h to an octopod nymphal stage. Nymphs develop in 1
2 days and take a blood meal from birds. The second nymphal stage does not feed and molts to an adult mite in ,1 day. The life cycle from egg to adult is 2
3 weeks depending on temperature and humidity. Rearing method(s) A device for artificial feeding of Ornithonyssus sylviarum has been developed consisting of a glass cylinder (25 mm long and outside diameter), capped with a 1-week-old white leghorn or broiler chicken skin membrane at one end and a snap cap with a wire window at the other end.44 Warmed (36 C
42 C) heparinized chicken blood is used with about 80% of the mites feeding. O. sylviarum can be feed and reared in the laboratory using Parafilm membranes stretched in two perpendicular directions, folded, heat sealed, inflated, and twisted at the ends to form a sphere allowing a high percentage ( . 80%) of surviving adult females to feed on whole chicken blood.45 Protonymphs feed through the membrane, and mites were able to feed from the same sac for 3 days with oviposited on the sac. Ornithonyssus bacoti is a pest of laboratory rodents and serves as an intermediate host for Litomosoides carinii, a filarid parasite of the cotton rat used as a laboratory model for testing antifilarial drugs.1

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In vitro method(s) Contact—filter paper—susceptibility and resistance Mites can be collected from feathers of mite-infested birds with a fine brush into a Petri dish and then placed in labeled plastic bags before use. The toxicity of acaricides against O. sylviarum can be tested by exposing adult mites to residues on filter paper for 24 h.46 Filter paper triangle packets (Whatman No. 1, 10 3 8 cm) are impregnated with test compounds dissolved and diluted in butanol:olive oil (1:1). Thirty adult mites, freshly collected by aspiration from vent feathers into glass Pasteur pipettes from untreated chickens, are inserted into each packet for continuous compound exposure. Mites are tapped from the pipette onto a 15 3 15 cm metal sheet placed on an electronic chill table (Bioquip model 1431) until immobilized and then counted into the packets. Mite mortality is assessed after incubation of the test packets for 48 h at 22 C and 75% RH with LD50 and LD99 values determined.47 Contact—glass surface—susceptibility and resistance Disposable Pasteur pipettes immersed in acetone solutions of acaricide concentrations (w/v) with the addition of O. sylviarum mites can be used to test the toxicity of acaricides.22 Straight thin tubes with an internal surface area of 10 cm2 on which a defined concentration (µg/cm2) is applied.23,24 Dilutions of acaricides in acetone are pipetted into tubes that are automatically rotated in a horizontal position in a flow of warm air to yield an evenly distributed deposit. Twenty female adult mites are inspirated into the treated and control tubes with a vacuum pump. Each tube is sealed with nylon tissue and the mites exposed for 24 h at 25 C and 90% RH. After the 24 h exposure the mites are transferred to clean recovery tubes and stored under the same conditions. Percent mortality is recorded after 24 h. Mites are considered dead if they neither responded to touch nor displayed normal movement.

In vivo method(s) Experimental infections can be induced in chickens. Mite-infested feathers are cut from mite-infested source hens and put in a plastic bag. Prior to infestation (week 21), each experimental hen is removed from her cage, and the feathers and skin of her vent area are checked to verify that they did not harbor any mites. One week later (week 0), when the hens were 25 weeks of age, B35 northern fowl mites are placed on the abdominal feathers of each hen. A glass Pasteur pipette is used to aspirate48 and transfer the mites from the plastic bag to the vent feathers. Each hen is removed weekly postinfestation from her cage; the feathers of the lower abdomen are sorted in a circle B8 cm in diameter anterior to the vent, and the number of mites are visually counted.43

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For the evaluation of parasiticide treatments, beak-trimmed caged laying hens (25
45 weeks old) are experimentally infested with 20
30 adult mites 3
5 weeks before treatment. Hens are housed in experimental, open-sided poultry houses in single suspended wire cages (31 3 31 cm of floor space), with continuous access to water and commercial layer mash and with a 16L:8D hour photoperiod. Hens are matched by infestation level and assigned to treated and untreated groups in equal numbers. Hens are treated using 8 mL of parasiticide solution applied to infested vent-region feathers and skin using a calibrated hand pump sprayer. Hens are scored for infestation level in the vent-region feathers (area B6 3 8 cm) using a visual scoring method, where 0 5 uninfested, 1 5 1
10 mites, 2 5 11
50, 3 5 51
100, 4 5 101
500, 5 5 501
1000, 6 5 1001
10,000, and 7 5 $ 10,000 mites.49 This method is increasingly conservative at high infestation levels but is highly correlated with actual mite numbers. Hens are scored on TD 3, 7, 10, 17, 24, and 31 PT.50 Larger numbers of test materials can be tested simultaneously using four singly held, caged, mite-infested brown hens per treatment per trial. Hens are treated with 15
20 mL of test solution applied to the vent feathers and underlying skin almost to runoff. Hens are evaluated using the visual scoring system above at the same PT intervals.50

Varroidae Varroa jacobsoni Oudemans, 1904 and Varroa destructor Anderson & Trueman, 2000—honey bee mite or Varroa mite Biology and life cycle Varroa mites are parasites of honey bees originally only occurring in Asia, but they have been introduced into Japan, Russia, Eastern Europe, Brazil and South America, Canada, United Kingdom, the United States including Hawaii, New Zealand, and Australia. Varroa jacobsoni attacks the Asian honey bee, Apis cerana. However, it has now adapted to the western or European honey bee, Apis mellifera. Adult mites are circular with females measuring 1
1.5 mm in diameter and reddish to dark brown in color.1 The mites suck hemolymph from adult bees and the brood, preferring drones that take three extra days to emerge compared to worker bees. A female mite enters a brood cell being sealed with the larval bee in the brood capsule and lays eggs after 60 h. The hatched larval mites feed on the developing bee to become adults and mate.51 Once emerged from the brood capsule along with the adult bee, the mite can enter a new capsule or attach to worker bees in order to spread the disease between hives. Varroa destructor attacks A. s cerana and the western or European honey bee, Apis melifera. The adult female are oval measuring

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1
1.8 3 1.5
2.0 mm with eight legs and reddish brown in color. Male mites are white. Varroa destructor mites leave the brood capsule with the merging young bee but then transfer to nurse bees that spend more time near the broad. The mites reproduce on a 10 day cycle.52

Rearing method(s) No specific publications on the rearing of Varroa mites were found. There are a number of methods for rearing and caging adult bees to test miticides against Varroa mites.53,54 In vitro method(s) Contact—direct application or fumigation (evaporation) Potential parasiticides against Varroa mites can be evaluated by evaporation or topical application tests.55 In the evaporation test, mites are fixed dorsally on a small glass plate and placed in a 37 mL glass exposed to 0.5 µL of test compound on a filter paper. The topical application test is conducted by fixing mites dorsally on a small glass plate and applying 0.2 µL of test solution to the ventral side of the mites. Mite mortality is evaluated after 24, 48, and 72 h PT. Contact—glass surface—susceptibility and resistance A bioassay was developed to evaluate miticides to control V. jacobsoni by exposing bees and mites to test compounds in Petri dishes (60 3 20 mm).56 Mortality was assessed at 24, 43, and 67 h PT. Lethal concentration values were determined for the test compounds. Mites collected from young bees are tested against potential miticides using wax tubes impregnated with test substance.57 Mites can remain in contact with the treated wax to determine attractiveness and toxicity of the test substance or move from it into a tube of pure wax (repellency test). Resistance testing of miticides against Varroa mites can be conducted using a discriminating dose to a specific compound or related class of compound.58 Glass vials (20 mL) are treated with dilutions of standard commercial test substances in acetone and rolled on their sides until dry leaving a uniform film of compound on the inner surface of the vial. Susceptible V. jacobsoni are removed from individual bee brood cells and placed in the treated vials. Three adult female mites, removed from nonpigmented bee larvae/pupae, are placed in each vial and held for 24 h at 27 C. A single bee pupa is added to each capped vial as food 6 h after initial exposure. An individual is considered dead PT if it showed no leg movement when gently prodded with a probe. Mite mortality and a discriminating dose are determined. Field collected mites are tested against the discriminating dose to determine resistance.

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Contact—wax—susceptibility and resistance A miticide resistance laboratory assay has been developed for pyrethroids.59 Shallow capsules are prepared of two glass disks (62 mm diameter) and one or two stainless steel rings (56 mm inner diameter, 3
5 mm total height). The interior of these capsules (including the rings) is coated with paraffin wax (Merck 7151, melting point 46 C
48 C) containing a known concentration of the active ingredient. Four grams of paraffin wax is melted in a Petri dish kept in a water bath heated to 60 C, and then the required amount of pyrethroid, dissolved in 2 mL hexane, is added. Hexane only is added as the control. The mixture is stirred for 1 min, and the hexane allowed to evaporate for at least 10 min. The steel rings are immersed into the molten paraffin wax, and one side of the glass disks is coated by lowering the disk onto the molten paraffin; in a series of 14 capsules, the total weight of paraffin is in the range 1.6
2.0 g. The capsules are then kept open for at least 24 h at RT to allow hexane residues to evaporate. The capsules are used for 1 month after being prepared; when not in use, they are kept at 23 C
29 C. Some assays, however, are carried out using capsules prepared about 2 months earlier, to assess the effects of aging. Ten or 15 Varroa females are introduced into each capsule; after 6 h they are transferred to a clean glass Petri dish (60 mm diameter) with respectively two or three worker larvae taken from cells 0
24 h after capping. The mites are observed under a dissecting microscope (when transferred to the Petri dish), 24 and 48 h after the introduction into the capsule and classified in the following categories: (1) mobile mites: when they could move when put on their legs and stimulated if necessary, though sometimes they were affected by the treatment to a varying degree, and their movements were more or less uncoordinated; (2) paralyzed mites: when they could move one or more appendages, but they could not progress; and (3) dead mites: when they did not react to stimulation repeated three times. The mites inside the capsules coated with paraffin and in the Petri dishes are kept in an incubator at 32.5 C and 70% RH. As a rule, the assay is repeated two or three times on mites from a given origin and a given brood stage, until 30 mites per concentration are assayed. However, assays were not repeated at concentrations higher than those which are expected to give 100% mortality, or when mortality not exceeding that of the controls is expected at a higher concentration, since these data give little information. In a few experiments, when the number of mites was limited, more mites were assayed at concentrations around the median lethal concentration.60

In vivo method(s) Apiary trials are conducted by selecting mite-infested bee colonies. Prior to the trial, mite infestation levels and the size of the colonies are monitored to obtain three homogeneous experimental groups (two treated and one control)

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of five hives each.61 Mite infestation levels of sealed worker brood and adult bees are estimated in all hives by inspecting three combs per hive62 and by brushing B300 bees per hive from at least three combs.63 The size of the colonies is estimated by the surface of sealed brood using one-sixth of a Dadant-Blatt frame (188 cm2) as a unit of measurement.64 Formulation treatments are applied according to label directions. Fallen mites and dead bees are counted in control and treated hives twice a week by using a white petroleum jelly-coated plastic sheet inserted at the bottom of each hive to count mites and Gray traps to count dead bees.65 Efficacy of the treatments are determined as the percentage of mite mortality.66,67 Honey and wax samples were analyzed for treatment residues before, during and after the treatments. Test compounds can also be tested by applying the test compound to four pieces of florist block material placed on the top bars of the upper hive body of each colony.68

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34. Nordenfors H, H¨oglund J, Tauson R, Chirico J. Effect of permethrin impregnated plastic strips on Dermanyssus gallinae in loose-housing systems for laying hens. Vet Parasitol 2001;102(1-2):121
31. 35. Nordenfors H, H¨oglund J, Uggla A. Effects of temperature and humidity on oviposition, molting and longevity of Dermanyssus gallinae (Acari: Dermanyssidae). J Med Entomol 1999;36(1):68
72. 36. Mul MF, Ploegaert JPM. Crawling insect counting device, system and method for indicating crawling insect infestation and determining a moment for treatment and/or control of said insects. Int Publ Number WO2014(098579):A1; 2014. 37. Mul MF, van Riel JW, Meerburg BG, Dicke M, George DR, Groot Koerkamp PWH. Validation of an automatic mite counter for Dermanyssus gallinae in experimental laying hen cages. Exp Appl Acarol 2015;66(4):589
603. 38. Tavassoli M, Allymehr M, Pourseyed SH, Owang A, Bernousi I, Mardani K, et al. Field bioassay of Metarhizium anisopliae strains to control the poultry red mite Dermanyssus gallinae. Vet Parasitol 2011;178:374
8. 39. Nordenfors H, H¨oglund J. Long term dynamics of Dermanyssus gallinae in relation to mite control measures in aviary systems for layers. Br Poult Sci 2000;41(5):533
40. 40. Axtell R, Arends JJ. Ecology and management of arthropod pests of poultry. Annu Rev Entomol 1990;35:101
26. 41. Ruff MD. Important parasites in poultry production systems. Vet Parasitol 1999;84:337
47. 42. Mullens BA, Owen JP, Kuney DR, Szijj CE, Klingler KA. Temporal changes in distribution, prevalence and intensity of northern fowl mite (Ornithonyssus sylviarum) parasitism in commercial caged laying hens, with a comprehensive economic analysis of parasite impact. Vet Parasitol 2009;160:116
33. 43. Vezzoli G, King AJ, Mench JA. The effect of northern fowl mite (Ornithonyssus sylviarum) infestation on hen physiology, physical condition, and egg quality. Poultry Sci 2016;95(5):1042
9. 44. Crystal MM. Artificial feeding of northern fowl mites, Ornithonyssus sylviarum (Canestrini) and Fanzago (Acari: Macronyssidae), through membranes. J Parasitol 1986;72 (4):550
4. 45. Carrol JF, Young KW, Bruce WA. Simple in-vitro feeding system for northern fowl mites (Acari: Macronyssidae). J Econ Entomol 1992;85(3):848
52. 46. Crystal MM, Demilo AB. Susceptibility of laboratory-reared northern fowl mites, Ornithonyssus sylviarum (Acari: Macronyssidae), to selected acaricides. Exp Appl Acarol 1988;4(4):353
8. 47. Mullens BA, Murillo AC, Heckeroth AR, Jirjis F, Flochlay-Sigognault A. Comparative in vitro evaluation of contact activity of fluralaner, spinosad, phoxim, propoxur, permethrin and deltamethrin against the northern fowl mite, Ornithonyssus sylviarum. Parasit Vectors 2017;10:358. 48. Owen JP, Delany ME, Mullens BA. MHC haplotype involvement in avian resistance to an ectoparasite. Immunogenetics 2008;60:621
31. 49. Owen JP, Delany ME, Cardona CC, Bickford AA, Mullens BA. Host inflammatory response governs fitness in an avian ectoparasite, the northern fowl mite (Ornithonyssus sylviarum). Int J Parasitol 2009;39:789
99. 50. Mullens BA, Soto D, Martin CD, Callaham BL, Gerry AC. Northern fowl mite (Ornithonyssus sylviarum) control evaluations using liquid formulations of diatomaceous

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58. 59. 60. 61.

62.

63. 64. 65. 66. 67.

68.

Parasiticide Screening, Vol 1 earth, kaolin, sulfur, azadirachtin, and Beauveria bassiana on caged laying hens. J Appl Poult Res 2012;21:111
16. Martin SJ. Ontogenesis of the mite Varroa jacobsoni Oud. In worker brood of the honeybee Apis mellifera L. under natural conditions. Exp Appl Acarol 1994;18:87
100. Rosenkranz P, Aumeier P, Ziegelmann B. Biology and control of Varroa destructor. J Invert Pathol 2010;103:S96
119. Kulincevic JM, Rothenbuhler WC. Selection for resistance and susceptibility to in the honey bee. J Invert Pathol 1975;25:289
95. Evans JD, Chen YP, di Prisco G, Pettis J, Williams V. Bee cups: single-use cages for honey bee experiments. J Agric Res Bee World 2009;48(4):300
2. Imdorf A, Bogdanov S, Ochoa RI, Calderone NW. Use of essential oils for the control of Varroa jacobsoni Oud. In honeybee colonies. Apidologie 1999;30:209
28. Lindberg CM, Melathopoulos AP, Winston ML. Laboratory evaluation of miticides to control Varroa jacobsoni (Acari: Varroidae), a honeybee (Hymenoptera: Apidae) parasite. J Econ Entomol 2000;93(2):189
98. Kraus B, Koeniger N, Fuchs S. Screening of substances for their effect on Varroa jacobsoni: attractiveness, repellency, toxicity and masking effects of ethereal oils. J Apicultural Res 1994;33(1):34
43. Elzen PJ, Baxter JR, Spivak M, Wilson WT. Control of Varroa jacobsoni Oud. Resistant to fluvalinate and amitraz using coumaphos. Apidologie 2000;31:437
41. Milani N. The resistance of Varroa jacobsoni Oud to pyrethroids: a laboratory assay. Apodologie 1995;26:415
29. Finney DJ. The estimation of the parameters of tolerance distributions. Biometrika 1949;36:139
256. Floris I, Satta A, Cabras P, Garau VL, Angioni A. Comparison between two thymol formulations in the control of Varroa destructor: effectiveness, persistence, and residues. J Econ Entomol 2004;97(2):187
91. Floris I. Osservazioni sull’infestazione da Varroa jacobsoni Oud di covata femminile opercolata di Apis mellifera ligustica Spin. In: Atti Convegno Stato Attuale e Sviluppo della Ricerca in Apicoltura, Sassari, Italy, 25
26 October 1991. p. 87
98. Ritter W, Ruttner R. Die Varroatose de Honigbiene. 3. Diagnoseverfahren. Allg Dtsch Imkerztg 1980;14:548
51. Marchetti S. Il metodo dei sesti per la valutazione numerica degli adulti in famiglie di Apis mellifera L. Apicoltura 1985;1:41
61. Gary NE. A trap to quantitatively recover dead and abnormal honeybees from the hives. J Econ Entomol 1960;53:782
5. Henderson CF, Tilton EW. Tests with acaricides against brown wheat mite. J Econ Entomol 1955;48:157
61. Floris I, Cabras P, Garau VL, Minelli EV, Satta A, Troullier J. Persistence and effectiveness of pyrethroids in plastic strips Against Varroa jacobsoni (Acari: Varroidae) and mite resistance in a Mediterranean Area. J Econ Entomol 2001;94:806
10. Calderone NW, Spivak M. Plant extracts for control of the parasitic mite Varroa jacobsoni (Acari: Varroidae) in colonies of the western honey bee (Hymenopter: Apidae). J Econ Entomol 1995;88(5):1211
15.

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Chapter 4e

Arachnida, Astigmata Josephus J. Fourie, PhD1, Maxime Madder, PhD, Professor2,3, Theo P.M. Schetters, PhD, Professor2,3 and Alan A. Marchiondo, MS, PhD4 1

Clinvet International, Bloemfontein, South Africa, 2Clinglobal, La Mivoie, Mauritius, Department of Veterinary Tropical Diseases, University of Pretoria, Pretoria, South Africa, 4 Adobe Veterinary Parasitology Consulting LLC, Santa Fe, NM, United States 3

Arachnida Astigmata Sarcoptidae

Sarcoptidae Sarcoptes scabiei (Linnaeus, 1758) Latreille, 1802—sarcoptic mange mite Biology and life cycle Sarcoptes scabiei occurs worldwide and causes sarcoptic mange or scabies of swine, bovids, dogs, foxes, great apes, horses, humans, koalas, and wombats.1 Sarcoptic mange of cattle and sheep is a national and local reportable animal disease in many countries. Scabies mites undergo four life cycle stages. Female mites burrow into the upper layer of the skin, but never below the stratum corneum, where she deposits two to three oval eggs (0.1
0.15 mm)/day. Eggs hatch in 3
4 days and the six-legged larvae exit the tiny serpentine tunnels of the wandering female and burrow into the skin and hair follicles (short burrows called molting pouches) to molt in 3
4 days into eight-legged nymphs. This stage then molts twice into adult mites. Female mites are oval shaped, ventrally flattened with a dorsally convex body covered with short measure setae and measure 0.3
0.45 3 0.25
0.35 mm. There are four pairs of legs with the two most anterior pairs having cushion-like sucker pads (pulvilli). Male mites are smaller than females (about half the size) and have suckers on all legs except the third pair. Males develop in about 9
11 days, while females may take up to 17
25 days. Mating occurs after the male mite enters the molting pouch of the adult female. Transmission occurs by transfer of impregnated mites during skin-to-skin contact or fomites.

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Rearing method(s) Mass cultivation of S. scabiei has been established in the laboratory on rabbits and pigs.2,3 Infested rabbits are maintained for a minimum of 30 days at 21
26 6 2 C and 65
71 6 5% RH. Rabbit skin lesions are scrapped with a scalpel and placed in Petri dishes containing 5 mL of 10% solution of potassium hydroxide and examined microscopically. Skin lesions are mixed with sodium chloride solution and centrifuges at 1500 rpm. A sample of resuspended sediment is examined on a counting chamber and the number of mites determined per drop of solution.4 Mites are also collected from infested rabbits by confining the animal in linen bags (2 3 2 ft2) covering all body surfaces except the mouth, eyes, and nose. Mites are collected every 24 h from the linen bags and transferred to cells (3 3 4 in.) containing a medium of fish flakes, multivitamins, amino acids, glucose, skin shavings, and horse serum semisolid globules along with live dust mites, Dermatophagoides farina. Sealed plastic cells are held at 18
24 6 2 C and 65
71 6 5% RH.4 S. scabiei var. suis can also be maintained indefinitely providing large numbers of mites ( . 6000 mites/g skin) using the continuous passage protocol described by Mounsey et al.3 This mite rearing method on pigs increases the intensity and duration of mite infestations using an optimized immunosuppression regimen of daily PO treatment with 0.2 mg/kg dexamethasone. To passage infestations, naive piglets are placed into pens adjacent to naturally infested pigs. A heater placed on the fence line encouraged pigs to congregate, thus enhancing the potential for mite transfer. Mite transmission can additionally be “boosted” in recipient pigs by the direct transplant of mite infested skin crusts harvested from infested pigs and dissected into small pieces (B0.5 cm2). Transplantation to establish infestations is done by inserting several crusts into both ear canals of naıve piglets. In vitro method(s) Contact—filter paper—susceptibility and resistance tests The sensitivity of S. scabiei to ivermectin was assessed by Currie et al.5 using a simple contact test. In this test the sensitivity of mite populations to ivermectin was determined at a concentration of 100 mg/g of ivermectin diluted in an emulsifying ointment. Test product (B0.1 g) was applied in a thin layer with cotton swabs to the bottom, top, and sides of 36-mm Petri dishes. Emulsifying ointment-coated Petri dishes were used as controls. Mite survival was assessed as described by Walton et al.6 through observation with a dissection microscope (160 3 magnification), for the following parameters: walking/active, slow movement, no movement/dead, and missing. Death was recorded when all movement and peristalsis of the gut had ceased, and the results analyzed using a log-rank test for survival.

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The acaricidal properties of an Ailanthus altissima bark extract against S. scabiei var. cuniculi were tested in vitro.8 Mites were collected from naturally infested New Zealand white rabbits according to the methods described by Walton and Currie.9 Scabs isolated and removed from the legs and ear cerumen of the rabbits were placed in Petri dishes and transported to the laboratory within 30 min and incubated at 35 C for 30 min. Ten mites were placed in each of several Petri dishes (10 cm in diameter, 2 cm deep) along with filter-paper chips to absorb any excess solution once the test solution was applied. A 2 mL solution of each concentration was directly added to the experimental treatment Petri dish. Negative control mites were treated with distilled water and glycerin, while positive control mites were treated with fenvalerate. All treatments were replicated six times. All dishes were incubated at 25 C at 75% RH, and observed under a stereomicroscope at 30 min PT, then every hour to 7 h PT. Mites were considered dead when they did not respond to stimulation with a needle. Contact test using medium The acaricidal effect of parathion, phosmet, and phoxim against S. scabiei was assessed in vitro by evaluating of the migration ability of mites in test dishes.2 Sterile polystyrene Petri dishes (internal diameter 8.5 cm) were filled with 20 mL of sterile agar solution containing the preselected concentrations of parathion, phoxim, or phosmet. The test compounds were added to the fluid (50 C) agar solution just prior to the casting of the dishes by first dissolving it either in ethanol (parathion and phoxim) or DMSO (phosmet). Ten mites were subsequently deposited on each test dish equally distributed along the diameter line and incubated at RT for 18 h. Following incubation, the dishes were arranged in a circle below a normal tungsten bulb (60 W), with a vertical and horizontal distance to the bulb of 25 cm and 40 cm, respectively, to stimulate migration toward the light. The migration ability of the mites on a given dish was evaluated by assessing (1) the number of mites, out of 10, having demonstrated the ability to migrate on the surface of the agar gel, as indicated by footprints and/or bacteria colonies, (2) the length of the migration track was estimated for each mite according to the following scale: ,0.3 cm 5 0; 0.3
1 cm 5 1; 1
3 cm 5 1 1 ; .3 cm 5 111. The mean migration for each plate was calculated on the basis of the migrating mites only, thus giving the mean migration score. In vitro experiments evaluating the acaricidal effect of different test extracts against S. scabiei was conducted using 96-well tissue culture plates containing 3.5 mL medium plus 30 µL of serial dilutions of test compounds.4 The medium used was prepared as described by Brimer et al.2 Twenty mites were added to each of the treated and control (negative and ivermectin positive—10 mg/mL) wells using a teasing needle. Wells were covered with

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16 mm diameter porous disks to avoid mites escaping. The dishes were incubated at 21 6 2 C and 70 6 2 C for 72 h in the dark. Mite mortality based on motility with a teasing needle was assessed at 24, 48, and 72 h PT. Contact and fumigation test—paraffin gel Contact and fumigation bioassays were performed on S. scabiei mites collected from experimentally infested pigs.10 For contact bioassays, essential oils were diluted with paraffin to yield concentrations at 10%, 5%, and even 1% for the most efficient oils. The mites were inspected under a stereomicroscope 10, 20, 30, 40, 50, 60, 90, 120, 150, and 180 min after contact. For fumigation bioassay a filter paper was treated with 100 µL of the pure essential oil. The mites were inspected under a stereomicroscope for the first 5 min, and then every 5 min until 1 h.

In vivo method(s) Parasiticide efficacy studies against S. scabiei should be designed and conducted in accordance with the WAAVP guidelines for evaluating the efficacy of acaricides against (mange and itch) mites on ruminants (see Appendix A of Ref. [11]). Efficacy studies against S. scabiei on experimentally infested pigs have been successfully conducted. In one such study the efficacy of ivermectin against S. scabiei var suis was evaluated.12 In this study, 12-week-old pigs were experimentally infested using B4800 mites and an unknown number of eggs. All of the pigs were positive for sarcoptic mange at the start of the treatment. Mite counts were conducted weekly up to 28 days PT, and the efficacy of ivermectin was assessed by the reduction in mite counts (control vs treated) in combination with the improvement of clinical signs using the scratching index with a cutoff value of 0
4.13 Efficacy studies against S. scabiei on dogs usually involves the enrollment of naturally infested animals due to the occurrence of self-cure when attempting to experimentally infest study animals.14
17 Efficacy evaluation in these studies are primarily based on the reduction of Sarcoptes mites counted in skin scrapings as well as the number of mite-free dogs compared to the controls. Skin scrapings ( 6 4 cm2) are performed prior to treatment and monthly thereafter up to 2 months from five different body areas suspected of being infested based on the lesions (crusts, hair loss, erythema, and/or papules) observed. The presence of clinical signs (crusts, hair loss, erythema, and/or papules) associated with mange infestations are also recorded/sketched on a dog silhouette (left- and right-hand sides). Pruritus was assessed on each dog on the days when scrapings are made during a 5-min observation period and documented as present or absent. The difference in the number of mites between treated and control animals was calculated. A percentage reduction in mite counts was considered to be a criterion

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for treatment success as well as the proportion of the total number of mitefree dogs in each group.

Notoedres cati (Hering, 1838) Railliet, 1893—feline scabies mite Biology and life cycle Notoedres cati is a parasitic mite of cats, primarily causing face or head mange, but can also infest rats, rabbits, and occasionally humans.1 The mite is closely related to Sarcoptes mange mites of dogs, but much smaller, and has been reported in Australia, Europe, India, Indonesia, Japan, the Middle East, and North and South America.18 It is morphologically distinguished from Sarcoptes by its dorsal anus. Adult male mites are round measuring 200 3 240 µm with very short legs and sucker-like feet on the first, second, and fourth pairs of legs. Female mites are about the same size and have sucker-like feet only on the first pair of legs. Females burrow deep within the dermis (stratum germinativum) and lay eggs that hatch to six-legged larvae (87
166 µm long 3 130
183 µm wide) with sucker-like appendages only on the first two pairs of legs. The life cycle of N. cati has not been described in detail. Based on the life cycle of Notoedres notoedres from the white rat, 11
25 eggs are laid in the burrow in the epidermis. Eggs hatch in 4
5 days and the larvae exit the burrow, crawl on the skin, and dig a molting burrow where they molt to the first-stage nymphs in 5
7 days. Nymphs leave the burrow, wander on the skin, and burrow to the stratum corneum with the posterior of the nymph protruding the tunnel. After 3
5 days, nymphs molt to the second-stage nymphs. They remain 3
5 days, then exit the burrow, and dig a third molting burrow to molt to the adult mite. The life cycle takes about 12 days from hatched larvae to adults. Male mites seek out females and mate in the nymphal molting burrow. Transmission between hosts occurs with larvae and nymphs.

Rearing method(s) No specific rearing methods for N. cati could be found in the literature, but the methods described for S. scabiei using rabbits as host could be considered. In vitro method(s) No specific in vitro methods for assessing the acaricidal activity of test compounds against N. cati could be found in the literature or known to the authors. The in vitro methods described for S. scabiei could be used for the evaluation of acaricidal compounds against N. cati.

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In vivo method(s) Naturally infested cats with N. cati were recruited for clinical efficacy studies based on clinical signs and positive results of viable mites by skin scrapings.19 Infested cats were randomly allocated to treatment and control groups by the pretreatment mite counts and assessments of notoedric skin lesions was performed weekly up to 28 days PT. The severity of notoedric lesions was scored as follows: 0 5 no skin lesions, no alopecia, and no scratching; 1 5 mild skin lesions, mild alopecia, and occasionally scratching; 2 5 moderate skin lesion, moderate alopecia, intensive scratching, and scratching wounds; and 3 5 severe skin lesion, severe alopecia, thick/crusty and scabby appearance of the skin, intensive scratching, and scratching wounds. The extents of lesions were scored as follows: 0 5 no skin lesions, 1 5 , 50% of the body skin surface affected, and 2 5 $ 50% of the body skin surface affected. Both scores were added up and expressed as a Notoedres-induced skin lesions score (NISLS) with sum values between 0 and 5. A final clinical assessment of Notoedres-induced skin lesions was performed based on NISLS on the final assessment day or any other earlier day of removal. The following outcomes were possible: (1) clinical cure 5 NISLS reduced to zero, (2) clinical improvement 5 NISLS ,50% of NISLS on TD 0 (Day 30), and (3) clinical failure 5 NISLS $ 50% of NISLS on TD 0 (Day 30).

Psoroptidae Psoroptes ovis (Hering, 1838)—sheep scab mite Biology and life cycle Psoroptes ovis is a nonburrowing parasitic mite of sheep and cattle found in Europe, Asia, and South America, having been eradicated from North America and Australasia. Bovine psoroptic mange is of increasing importance in the United States and Europe. Adult mites are 50 µm long, oval shaped with all the legs projecting beyond the body margin. The adult mites have piercing
chewing mouthparts and have rounded abdominal tubercles with characteristic three-jointed pedicels-bearing trumpet-shaped, sucker-like pulvilli. Males have copulatory suckers and paired posterior lobes with pulvilli on the first three pairs of legs and setae on the fourth pair. Female mites have a life span of about 16 days and can lay about 40
50 eggs on the skin surface of the host. Hexapod larvae (330 µm long) hatch from the oval eggs (250 µm in length) within 4 days and develop through two nymphal stages (protonymph and tritonymph) and then to mature adult mites. The life cycle from egg to egg is about 10 days. Mite populations on sheep grow at B11%/day and doubles every 6.3 days. Psoroptic mange lesions occur on the neck, shoulders, back, and flank

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of sheep, but they may be found on any part of the body. Transmission from host to host is by direct contact, however, adult mites can survive for about 18 days or more off the host allowing for fomite transmission.20,21

Rearing method(s) P. ovis mite colonies can be maintained on stanchioned cattle. Mites are collected using a thermal plate apparatus and held at 28 C and 85% RH in glass jars.22 Adult female mites can be fed in vitro using a Sykes
Moore chamber (32 mm in diameter 3 7 mm in height). The viewing area is 2.83 cm2, and the vertical working area is 1.5 mm with a volume of B0.65 mL. A silicone gasket (25 cm diameter) provides an internal seal between the top and bottom of the chamber. The top of the chamber is fitted with a glass cover slip that prevents evaporation and the bottom with a 112-mesh nylon netting. Mites (30) are introduced into the chamber with a 23-gauge needle by puncturing the O-ring gasket in the side of the chamber. The Sykes
Moore chamber is placed on a Conway diffusion chamber containing the feeding solution. The inside of the netting becomes wet by capillary action allowing the mites contact with the feeding solution. After feeding, mites are washed in Sykes
Moore chambers in a mild nondetergent soap solution to remove feeding solution, then rinsed in distilled water, and blown dry with compressed air. A positive feeding response is determined by a red color change in the mites when fed on whole blood or fed on other solutions containing 4
8 µg/mL of neural red dye (pH 6.8
8.0) added to the solution.23 Feeding responses were about 85% for female mites held off host cattle for 1 day with mites preferring water, plasma or serum and low-salt diets over whole blood. Feeding occurred within 5 min of diet exposure with a maximum response within 30 min of contact. Mites ingested an average of 0.29 µL of diet solution/100 mites with no differences observed between 10 C and 42 C. Live mites were also maintained in the laboratory using in vitro chambers constructed using two Perspex slides (30 3 60 mm) with a 15 mm diameter hole cut in the center of both.24 A 25 mm sterile circular filter-paper disk (Grade 1 Whatman) is placed in between the slides and secured with elastic bands or masking tape at each end. Mites are then placed, using either a mounted needle modified with a plastic hair or a fine paintbrush onto the filter paper with a cover slip secured on top with two elastic bands. Water, lamb serum, and two previously reported preferred diets of P. ovis were tested by administering 10 µL to the underside of the filter paper, daily.25 Mite chambers are then placed onto a moist tray of cotton wool and covered loosely with cling film to maintain a high RH.24,26 The chambers are placed in an incubator at 24 6 2 C and RH was measured using a HT50 meter. The positions of the chambers on the tray are rotated daily to avoid any incubator effects. Mite chambers are checked daily under a dissecting microscope to

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ensure sufficient moisture levels in the chambers and also for mortality. Mites are pronounced dead if they fail to move in response to manipulation with a paintbrush.27

In vitro method(s) The response of P. ovis collected from sheep to pyrethroids was evaluated in vitro by enclosing mites in small “tea bags” made from heat-sealable paper prior to dipping in insecticide.7 Mites failed to die 24 h after a 1 min dip in working concentrations of insecticidal sheep dips. With flumethrin a variety of different conditions were tested, but most failed to improve the efficacy of flumethrin. In vitro evaluation of bacteriophage therapy against P. ovis has been conducted.27 A Tranjector 5246 2 in 0.5 µm Femtotips was used at a constant pressure of 80 psi for a range of injection times (0.1, 0.5, 1, and 5 s). The average injection volume was calculated based on a scalene ellipsoid equation using P. ovis measurements from SEM images with the aim of injecting no more than 1% of the total mite volume.28 Mites from an in vivo culture within 48 h of harvest were cooled on ice and then stuck upside down onto double sticky
sided tape in a line on top of four microscope slides with a needle inserted at a 150 degrees angle, moving only the microscope stage.29 A back pressure of about B100 psi was used. The needle was aimed at the center on the side of the mite gut cavity to minimize damage.24 In addition, a needle-stick-only and noninjection control was used. Mite mortality was checked every 2 h for the first 8 h. The effect of fungal pathogens on P. ovis was evaluated in vivo and in vitro.30 In this study, mites were confined in 25 mm diameter chambers that were attached to the backs of 6 scab-naive sheep. In some treatments, mites were exposed to the fungal pathogens for 48 h in vitro prior to being placed on the host, while other treatments involved mites with no prior exposure placed directly onto the skin of a host treated with a fungal pathogen. After 48 h on the host, mites were removed, incubated individually, and all fungal infections were recorded. Fungal infection was observed in all treatments, except untreated controls. Psoroptes mites can become infected by entomopathogenic fungi on the skin of sheep and provides the first demonstration of the potential of this technology for the control of sheep scab. In vivo method(s) Parasiticide efficacy studies against P. ovis should be designed and conducted in accordance with the WAAVP guidelines for evaluating the efficacy of acaricides against (mange and itch) mites on ruminants (see Appendix A of Ref. [11]).

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Psoroptes cuniculi (Delafond, 1859)—rabbit ear mite or ear canker mite Biology and life cycle Psoroptes cuniculi is a nonburrowing parasitic mite of rabbits. The mite has essentially the same morphology and life cycle identical to P. ovis.21

Rearing method(s) No specific rearing methods for P. cuniculi could be found, but the methods described for P. ovis could be considered.

In vitro method(s) The acaricidal activity of aqueous extracts of chamomile flowers, Matricaria chamomilla, against P. cuniculi was tested in vitro using a contact test.31 Male and female adult mites used for the tests were isolated from scabs and ear cerumen collected from the ears of an infested rabbit. The test plates used were 4 cm in diameter, and 30 mites/plate, each containing 2 mL of a single extract to be tested (nine replicates/extract), were used. Another nine plates (30 mites/plate) containing 2 mL of saline solution were used as controls. Each plate was placed in a 9 cm diameter Petri dish, and these were placed in a humidity chamber (90%) at 22 C. After 24 h, the extracts were aspirated with a pipette from three of the nine plates of each extract and replaced with 2 mL of saline solution for 24 h and subsequently examined using a stereomicroscope. The same procedure was adopted for the remaining plates after 48 h (three plates) and 72 h (three plates). Mites found motionless after stimulation with a needle were regarded dead. In another study the efficacy of an essential oil of Eugenia caryophyllata against P. cuniculi was evaluated using a similar test as previously described.32 In this test, mites were also isolated from naturally infested rabbits, but placed in 6-cm Petri dishes. The essential oil of leaves of E. caryophyllata was diluted from the concentration of 10%
0.03% (10%, 5%, 2.5%, 1.25%, 0.62%, 0.31%, 0.16%, 0.010% 0.06%, and 0.03%) in paraffin oil and 2.5 mL of each solution was added to Petri dishes, six replications were made for each concentration. For negative and positive control dishes, six Petri dishes containing only 2.5 mL of paraffin oil and six Petri dishes containing 2.5 mL of a pyrethrum extract containing 25% of pyrethrins were used. Plates were placed in a humidity chamber in saturated humidity conditions at 22 C, and after 24 h, mites were transferred to clean Petri dishes containing 2.5 mL of paraffin oil. The Petri dishes were subsequently examined after a further 24 h, and all the motionless mites were stimulated with a needle and regarded dead if remaining motionless.

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The hatchability of P. cuniculi eggs was assessed in vitro after exposure to acaricidal compounds.33 In this test, adult females and eggs were isolated randomly, using a fine needle, from scabs removed from the ears of naturally infested rabbits. All bioassays were performed on either 50 eggs or 50 adult females maintained at 27 C. RH of 100% was obtained by sealing the handling chambers with Parafilm, and all ovicidal and adulticidal assays were conducted in quadruplicates and triplicates, respectively. Three kinds of handling chambers were evaluated. In the first approach (humidity chamber), eggs or adult females were transferred to 1.5 mL Eppendorf tubes containing a piece of cotton wool saturated with a solution of TC-100 insect medium to which 2% penicillin
streptomycin had been added. For each assay, 50 or 5 tubes containing 1 egg or 10 females, respectively, were used. Tubes were examined under a dissecting microscope (30
40 3 magnification) by picking up the cotton wool and counting the larvae/nonhatched eggs to determine the hatching rate. The viability of the larvae and adult females was determined based on motility following a gentle mechanical stimulation. Assessments were done on TD 8 for the eggs and on TD 3 for the females. In the second approach (liquid medium), eggs or adult females were immersed into 500 mL of TC-100 insect medium containing 2% penicillin
streptomycin and 10% horse serum in 24-well microplates. Ten eggs or 10 adults were placed in each of five wells. Eggs, larvae, and adult mites were counted daily under an inverted microscope (25 3 magnification) without opening the plates, and the number of living mites was estimated according to the presence or absence of motility during a 2 min observation period. The third approach (agarose dishes) was adapted from Bowman et al.34 In this bioassay, 3 mL of 1% liquid agarose in distilled water was added to 3.5-cm Petri dishes. Ten eggs and 10 adult mites were laid down in the center of each of five dishes and sealed. The sealed dishes were subsequently examined daily under a dissecting microscope (30
40 3 magnification) to determine hatching and survival rates. In addition, the position of each nonmotile mite was recorded on the dish allowing for the confirmation of mite death during the next observation. Dishes were examined for up to 8 days to evaluate egg hatching. Amitraz (125 mg/mL) and phoxim (500 mg/mL) diluted in test media at their working concentrations of 0.2% and 0.1%, respectively, were evaluated using these bioassays. The test media used were: TC-100 insect medium plus penicillin
streptomycin, TC-100 plus penicillin
streptomycin with horse serum, and agarose for approaches 1, 2, and 3, respectively. The test compounds were evaluated in vitro using the three protocols. For approach 3 the insecticides were added just before polymerization (approximate temperature of 55 C). Insecticide-free test media were used as controls.

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In vivo method(s) Parasiticide efficacy studies against P. cuniculi on sheep and goats should be designed and conducted in accordance with the WAAVP guidelines for evaluating the efficacy of acaricides against (mange and itch) mites on ruminants (see Appendix A of Ref. [11]). The efficacy of selamectin for the treatment of P. cuniculi infestation in rabbits was investigated in vivo.35 In this study, rabbits were evaluated for the severity of mite infestation prior to treatment based on ear lesion size. Lesions were recorded as the sum of the area of the lesions in each ear, measured to the nearest 0.25 cm2. Rabbits were grouped according to lesion size and randomly allocated to treatment groups (eight rabbits/group) so that each treatment group contained rabbits with a similar range of lesion severity. After assignment to treatment groups (TD 0), rabbits were treated topically with vehicle or selamectin twice, 1 month apart. Mite viability PT was assessed by otoscopic examinations and ear lesion size weekly up to 1 month after the last treatment. At the conclusion of the study, rabbits were euthanatized, and quantitative counts of viable P. cuniculi mites (larvae, nymphs, and adults) were performed on all rabbits. In order to assess the acaricidal activity C. zeylanicum essential oils on P. cuniculi, rabbits were experimentally infested and a qualitative assessment done to assess efficacy.36 In this study, the qualitative assessments were done on TD 0, 7, 14, and 30 PT by examining the rabbit’s ears with an otoscope for the presence of scabs. If observed, the cerumen and/or scabs were collected and microscopically observed to evaluate the presence of mites and the degree of infestation evaluated on the basis of the following scoring system: 0 5 the absence of scabs and/or mites; 0.5 5 irritation in ear canal, but no mites observed; 1 5 small number of scabs in the ear canal, mites present; 2 5 external ear canal filled with scabs, mites present; 3 5 scabs in ear canal and proximal one-fourth of pinna, mites present; 4 5 1/2 pinna filled with scabs, mites present; 5 5 three-fourths of the pinna filled with scabs, mites present; and 6 5 all internal surface of the pinna full of scabs, mites present.37 The hormonal and behavioral changes induced by acute and chronic experimental infestation with P. cuniculi were investigated in experimentally infested rabbits.38 Rabbits were infested by placing 150 mites in the ear canal that was occluded with cotton held with tape. To confirm that rabbits were infested, macroscopic lesions were identified with an otoscope. In this study the changes in exploratory behavior and scent marking evoked was investigated by acute (1
9 days) and chronic (25
33 days) experimental infestations.

Chorioptes bovis (Hering, 1845)—chorioptic mange mite Biology and life cycle Chorioptes bovis is a parasitic nonburrowing surface mite infesting mainly cattle and horses, but also sheep, goats, and rabbits. The mite is distributed

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geographically worldwide. Adult mites have chewing mouthparts and are oval shaped with long legs and cup-shaped sucker on unsegmented pedicles. Male mites have two broad, flat setae on the posterior lobes. Adult female mites measure B300 µm long with a pair of long, terminal whip-like setae on tarsi 3. Female mites have a life span of 4
6 weeks and can lay about 17 eggs on the skin surface of the host. Larvae hatch from the eggs within 4 days, develop through two nymphal stages, and then to mature adult mites. The life cycle takes about 3 weeks. Predilection sites of chorioptic mange include the skin of the legs, udder, abdomen, perineum, and base of the tail. Transmission from host to host is by direct contact, however, adult mites can survive for B3 weeks off the host allowing for fomite transmission.20

Rearing method(s) A technique has been developed for rearing reproducing populations of C. bovis for periods of 3
8 months, using mites sourced from foot scrapings of a cow.39 Mites together with epidermal debris and hair in the scraping were placed in a series of vials that, in turn, were maintained in a jar with a volume of 1-gal. Paraffin was placed in the bottom of the jar (at least 1 cm deep, appropriate for excavation) securely holding about 10 vials (5 cm high). To maintain humidity within the jar at about 80%, sulfuric acid on the paraffin was used. The lip of each vial was circumvallate with lead in order to keep the vial upright in the paraffin when the solution was present. The solution also served as a moat preventing the exchange of mites between vials. Mites were retained in the vials using bolting silk, held in place with elastic bands. The jar was sealed with petroleum jelly and placed in an incubator at 35 C. This simple technique was suitable for rearing mites through sequential generations for several months. By transferring mites to vials with unused epidermal debris, it was possible to perpetuate them longer.

In vitro method(s) The acaricidal efficacy of extracts obtained from the plant Eupatorium adenophorum against the common cattle mite, Chorioptes texanus, was evaluated using the methods described by Fichi et al.32 as summarized for P. cuniculi by Nong et al.40

In vivo method(s) Parasiticide efficacy studies against C. bovis should be designed and conducted in accordance with the WAAVP guidelines for evaluating the efficacy of acaricides against (mange and itch) mites on ruminants (see Appendix A of Ref. [11]).

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The efficacy of an eprinomectin pour-on was evaluated in treating chorioptic mange in dairy cattle.41 In this study, 12 healthy female Fleckvieh (Simmentaler) cattle (6 heifers and 6 nonlactating dairy cows naturally infested with C. bovis) were included. Infestations were confirmed by the examination of skin scrapings taken the day before treatment (TD 21). To exclude effects on production resulting from the anthelmintic properties of eprinomectin, all animals were dewormed with FBZ administered PO at twice the manufacturer’s recommended dose on TD 26. Based on TD 21 mite counts, the animals were randomly allocated to untreated controls or were treated with eprinomectin 0.5% pour-on solution at 1 mL/10 kg bw (0.5 mg eprinomectin/kg bw). The animals were individually tethered in a barn in such a way as to prevent direct contact and were randomly allocated to blocks of adjacent stanchions and within blocks randomly assigned to a position between them. Live mites were counted in scrapings collected from the edges of active lesions or, if lesions regressed during the study, from the area where active lesions were at study commencement. Six scrapings were made on TD 21, TD 7, and then at weekly intervals until TD 56 on each animal using a sharp spoon from an area B3 3 3 cm. Samples were evaluated within 8 h of collection by washing the scraped material out onto black filter-paper disks that were examined thoroughly for live mites under a stereomicroscope. The live mites present from each scraping were counted up to a maximum of 100. Counts above 100 were recorded as .100. Moreover, the site of each scraping was recorded on a silhouette together with a description of the mange lesions at each examination. The lesions were scored as follows: 0 5 healthy skin; (1) 5 healing lesion, crusts lifted detached easily but hair growth not complete; 1 5 active lesion, extent of less than the palm of the hand; 11 5 active lesion, extent of more than the palm of the hand; and 11 1 5 active lesion, extent of more than the half of the body of the animal. In addition, all animals were weighed on TD 21 and 56 and individual feed consumption was measured daily from TD 0
56. To calculate the efficacy the mite counts were transformed to the natural logarithm (count 1 1) for the analysis and calculation of geometric means.

Otodectes cynotis (Hering, 1838) Canestrini, 1894—dog ear mite Biology and life cycle This parasitic mite infests the ears of the dog, cat, fox, and ferrets. The geographical distribution of the mite is worldwide. This mite does not burrow the skin but has a predilection for the ears. The mites are visible macroscopically appearing as small, white, and mobile organisms with males measuring 274
362 µm long with two ventrally situated suckers and caruncles on all four pairs of legs. Female mites measure 345
451 µm long with long hairs or setae on the third and fourth pairs of legs. The anterior pairs of legs at the

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pretarsi have white-glass-shaped caruncles on a short pedicel. Female mites glue eggs to the ear canal that hatch in 4 days. Larval mites measure 138
224 µm long and develop to protonymphs in 3
5 days, molting in B24 h. Protonymphs develop to deutonymphs in about the same time and molt to adults. Ovulation is believed to occur when adult females shed its deutonymph exoskeleton. The life cycle takes B18
28 days from egg to egg. Hosts probably become infested by direct contact.18

Rearing method(s) The most widely used methods for obtaining the required number of mites needed for in vitro or in vivo studies are to either harvest the mites from naturally infested animals or to establish infestations experimentally on donor animals. To harvest mites from the ears of infested animals the following ear flushing technique can be used. Prior to ear flushing, the ear canals have to be examined with an otoscope to determine with reasonable certainty whether or not mites are present, without removing mites. If established that an infestation is present, the animal is sedated using a suitable sedative. Once sedated, the ear duct is filled with 5% aqueous solution of docusate sodium and slightly massaged to soften the ear duct content. The content of the ear duct is subsequently removed from the ears and filtered through a 38-µm sieve followed by a flushing with lukewarm saline solution that is poured through the same sieve. Once completed, the ear duct must be examined otoscopically and, if needed, the flushing process is repeated until the ear ducts are determined clean (no visible cerumen or mites). The sieve contents can then be rinsed with water and transferred to a Petri dish, and all live mites (adults, larvae, and nymphs) counted under a stereomicroscope. The required number of mites needed for an in vitro test can then be pipetted from the solution, or if required to infest a recipient animal, each ear flushing can be filtered through filtration paper filled at the bottom by a small tuff of hair taken from the donor animal. The extra tuff of hair serves as an additional filtration medium and also assists in collecting the mites for infestation after drying. Each filtration packet should contain at least 50
200 mites to ensure a successful experimental infestation. Allow the filtration packets to dry sufficiently before experimental infestation. When infesting an animal, the filtration packet is opened and examined visually for mites. Using a pair of forceps, the tuff of hair can be taken and mites from the filtration paper collected by lightly brushing over them with the tuff of hair. The tuff of hair is subsequently placed into the ear canal of the sedated recipient animal. After collecting the mites from the filtration paper, the filtration paper is rubbed gently against the inside of the animal’s ear to ensure that any mites that stayed behind are transferred to the animal. The ear of the animal infested is wrapped for up to 1 day to prevent the animal from removing the infested

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material. If mites are present in the ears of recipient animals (as confirmed by an otoscopic examination) after at least 2
3 weeks, the artificial infestation can be considered to be successful.

In vitro method(s) The acaricidal activity of four monoterpenes and solvents against Otodectes cynotis were assessed in vitro.42 In this study the mites were collected from six naturally infested cats. Mites were collected from all the animals by swabbing both ears and placing the swabs in 15 mm Petri dishes for identification and classification within 12 h of collection. The mites were classified as adult male (M), adult female (F), nymph (N), or larva (L). The four monoterpenes to be tested were first diluted to 10% in paraffin oil that was found to be the most effective solvent and further diluted to 5% and 1%. The test was repeated three times for each monoterpene and dilution. A total of 1080 O. cynotis mites were used (438 L, 298 N, and 344 adults—233 M and 111 F) to test monoterpenes. After identification under the stereomicroscope, the various life cycle stages were placed in different Petri dishes, and weak or dead mites were immediately removed from the dishes. The remaining mites were evenly divided into groups (30 mites/group) and placed in the Petri dishes (4 cm diameter). The test Petri dishes were prepared by fitting the lids with blotting paper disks (5 cm diameter) and adding the test solution to the blotting paper. After several attempts with different amounts of liquids, 100 µL was considered most suitable to add to the blotting paper. After 5 min the fluid spread completely and uniformly over the paper, and the mites were transferred for the various tests. To guarantee saturated humidity and to reduce evaporation of the test products, the Petri dishes were placed in a 25 3 15 3 5 cm plastic box with waterlogged absorbent tissue and covered with aluminum foil. The plastic box containing the Petri dishes was placed in a constant temperature cabinet at 35 C. All the dishes were examined hourly by stereomicroscopy up to the fourth hour, every 12 h until the 60th hour, and every 24 h until the death of the last mite (the end of the trial). Lack of natural movements, even when stimulated with a needle, was considered indicative of death. In another study a rapid quantitative assay was used to assess the acaricidal effects on O. cynotis.43 As for the previous study, mites were obtained from naturally infested cats and transferred to empty blood test tubes. Mites were isolated by incubating the tubes at 35 C for 30 min before the ear wax was transferred to empty polystyrene Petri dishes. Mobile mites leaving the ear wax were separated under a stereomicroscope and gently transferred to test plates. The ear wax with mites could be stored in the blood test tubes for 4
5 days at 5 C without a reduction in the ability of mites to migrate. Test plates (internal diameter 8.5 cm) were prepared as described by Brimer et al.,2 with diazinon added as an ethanolic solution. The ethanol concentration

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was ,1% v/v of prepared agar and concentrations of diazinon were expressed in mg/g of solidified gel. For the preparation of test plates with amitraz, a commercial formulation was used and diluted with sterile water to a set of standard solutions that were added, as microemulsions, to sterile agar solution (Columbia agar base 40 g/L) at 50 C. No more than 5 mL of the standard solutions were used per liter of prepared agar. The stability of the two acaricides was determined by analysis of plates containing diazinon or amitraz after various periods of storage at different temperatures. To evaluate the acaricidal effect, 10 O. cynotis mites were deposited on each test dish equally distributed along a line 2 cm from the edge of the 12 cm Petri dish. Plates were placed upside down, and each Petri dish was covered with an extra lid that was painted black to protect the mites from the light. The plates were subsequently incubated at RT for 6 h arranged in a circle below a tungsten bulb (60 W), with a vertical and horizontal distance to the bulb of 10 and 25 cm, respectively. The effect on the mites was calculated (on the basis of five plates/concentration) as follows: 1. The number of mites, out of the 10, having demonstrated the ability to migrate on the surface of a plate, was established (n). 2. The final position of each of these mites was marked with a pen in the bottom of the Petri dish. 3. The dish was subsequently placed on a scale with the starting line for the mites covering the starting line on the scale. The scales had three parallel lines, that is, the starting line and two additional lines giving rise to three zones, of which the outermost was open toward the edge of the dish. The scales had the following graduation: 1 point (0
1.5 cm), 2 points (1.5 , x # 4.5 cm), and 3 points ( . 4.5 cm). 4. The number of mites in each section of the scale was counted, and each mite was given a score according to the scale, thus giving rise to the total score per plate(s). 5. The mean migration score for each plate was calculated on basis of the migrating mites only, as s 5 n. 6. For each concentration a migration index was calculated as (Psm 5 number of plates).

In vivo method(s) The efficacy of a new spot-on formulation of selamectin plus sarolaner in the treatment of O. cynotis in cats was evaluated in vivo.44 Mite infestations were induced by the interaural transfer of mites from naturally infested donor animals as described previously in the rearing method(s) section above for O. cynotis. Cats were included in the studies if they harbored at least five live mites in at least one ear. Cats were blocked based upon whether they had bi-or unilateral infestations at study start and upon body weight and then

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were randomly assigned to placebo-treated or selamectin/sarolaner-treated groups. Cats were examined otoscopically for the presence of live mites 14 days PT, and total ear mite counts done 30 days PT. For total mite counts, cats were sedated and each ear was flushed, and the ear canal contents were processed separately as described previously in the in vitro rearing section. All live mites (adults, larvae, and nymphs) were counted, and the counts for the two ears were summed for each animal’s total ear mite count. Efficacy (% reduction) was calculated using the Abbott formula.45 In another study the efficacy of afoxolaner against O. cynotis infestations in dogs was evaluated in vivo.46 In this study, experimental infestations were also performed as previously described, and B50
100 mites, depending on the intensity of infestation in donor animals, were transferred into each ear of the recipient animal. For each dog a total of .10 live mites in at least one ear, as well as the presence of $ 1 live mite in the other ear, was necessary for inclusion in the study. Once qualified, the dogs were randomly allocated to the study groups. The otoscopic assessments were performed on all dogs prior to treatment and biweekly after treatment to confirm the initial presence, and subsequent presence or absence, of live mites. At 28 days PT a qualitative assessment of the ear canals was performed prior to the flushing procedure to determine the presence of viable mites. This was followed by the quantitative assessment of ear mites by ear duct flushing (as previously described) with complete mite collection and count. The efficacy was calculated as described for the previous study. Similar studies to assess the efficacy of fluralaner against O. cynotis infestations in dogs and cats in vivo has also been done by Taenzler et al.47

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8. Gu X, Fang C, Yang G, Xie Y, Nong X, Zhu J, et al. Acaricidal properties of an Ailanthus altissima bark extract against Psoroptes cuniculi and Sarcoptes scabiei var. cuniculi in vitro. Exp Appl Acarol 2014;62(2):225
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27. Hill S. Novel control of the sheep scab mite, Psoroptes ovis, through the application of bacteriophage therapy [Ph.D. thesis]. The University of Edinburgh; 2011, 250. 28. Jasinskiene N, Juhn J, James AA. Microinjection of Aedes aegypti embryos to obtain transgenic mosquitoes. J Vis Exp 2007;5:219. 29. Presnail JK, Hoy MA. Stable genetic transformation of a beneficial arthropod, Metaseiulus occidentalis (Acari: Phytoseiidae), by microinjection technique. Proc Nat Acad Sci USA 1992;89:7732
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45. Abbott WS. A method for computing the effectiveness of an insecticide. J Econ Entomol 1925;18(2):265
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Chapter 4f

Arachnida, Cryptostigmata Josephus J. Fourie, PhD1, Maxime Madder, PhD, Professor2,3 and Theo P.M. Schetters, PhD, Professor2,3 1 3

Clinvet International, Bloemfontein, South Africa, 2Clinglobal, La Mivoie, Mauritius, Department of Veterinary Tropical Diseases, University of Pretoria, Pretoria, South Africa

Arachnida Cryptostigmata Demodicidae

Demodex spp. (Demodex canis Leydig, 1859, Demodex cati Hirst, 1919; Demodex equi, Demodex ovis)—Demodectic Mange Mite or Red Mange Mite Biology and life cycle Mites of the Demodex genus are burrowing ectoparasites with recognized species named after their mammalian host, that is, Demodex canis in the dog, Demodex cati in the cat, Demodex equi in equids, Demodex ovis in ovines.1,2 The mites live in a characteristic head-downward posture as commensals within the hair follicle of the host and have a cigar shape (up to 0.2 mm long) with four pairs of stumpy legs anteriorly. The entire life cycle of most species takes place in the hair follicle. Eggs are laid within the epithelial cavity of the hair follicle and hexapod larvae hatch. The larvae develop to protonymphs and these to octopod nymphs before developing to the adults. It is probably impossible to transmit Demodex by contact, unless there is prolonged contact like during nursing of young animals. Rearing method(s) Up to date Demodex spp. have not been cultivated successfully without the use of their natural hosts. Attempts have been made to establish and maintain an in vitro culture of human Demodex spp.3 Mites, collected by the cellophane tape method from volunteer students, were cultivated in RPMI 1640 with L-glutamine and HEPES buffer. The RPMI medium was used in three combinations: RPMI enriched with human serum, RPMI enriched with FBS, and plain RPMI. The viability of the mites was observed at intervals of 8 h, recording the time of death according to the criteria of mobility of the body and its appendices for 1 min. RPMI enriched with human serum showed to be the best to grow and maintain viable mites: mortality after 8 days of culture was 40% at 15 C and 70% at 25 C.

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In vitro method(s) Demodex mites are collected by adhesive cellophane tape technique. The lack of standardized in vitro culture models to maintain mite viability and reproduce mites through all their life stages limits the options available for in vitro parasiticide screening. Apart from that, the determination of viability of Demodex mites is far from easy and most often based on the mite’s movement or motion. There is however a significant variability between investigators when it comes to motion scoring.4 Genetically immunodeficient mice (SCID mice) have been used as a xenograft model to study the biology of D. canis.5 Skin grafts from “clean” dogs were transplanted to SCID mice and subsequently, after healing, infested with D. canis mites collected from clinical cases. The collection of mites was done by skin scrapings, using a scapular blade coated in mineral oil and, second, by taking 6 mm diameter skin biopsies from clinical dogs. The scrapings were examined microscopically and applied directly to the canine skin grafts on the mice and bandaged, whereas the skin biopsies were applied to the skin grafts with the graft epidermis contacting the epidermis of the biopsy. The biopsies were helped into place with tissue adhesive and bandaged for 1 week.6 These artificially infested SCID mice with canine skin grafts could serve as a model for parasiticide testing against D. canis. It could be hypothesized that the same xenograft model could be developed for other Demodex species using skin grafts from their respective hosts. In vivo method(s) A more common parasiticide screening model for Demodex spp. is the use of naturally infested hosts. Fourie et al.7 used dogs with generalized demodicosis mange to test chewable fluralaner tablets. The evaluation of the infestation, both for randomization, before and after treatment was based on microscopic screening of deep skin scrapings of 4 cm2. Similar methodology was used by Six et al.8 to test sarolaner and by Snyder et al.9 to test lotilaner against naturally occurring Demodex spp. in dogs.

Cheyletiellidae Cheyletiella spp. (Cheyletiella blakei Smiley, 1970; Cheyletiella yasguri Smiley, 1965; Cheyletiella parasitivorax Me´gnin, 1878)— walking dandruff mite Biology and life cycle Three Cheyletiella species are surface dwelling, nonburrowing parasitic mites found on distinct hosts, that is, Cheyletiella blakei on dogs, Cheyletiella yasguri on cats, and Cheyletiella parasitivorax on rabbits. All three species are morphologically similar. Adult mites are ovoid, measure

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B400 µm long, with blade-like chelicerae and enormous hook-like accessory mouthparts (palps). The palps have curved palpal claws, and the palpal femur possesses a long, serrated seta. The body is slightly elongate with a waist per say and has been described as a shield, bell pepper, or western horse’s saddle when viewed from above. Adult mites have short legs with no tarsal claws and the empodium is a narrow pad with comb-like pulvilli at the ends of the legs. The peritreme is M shaped. The three species can be separated morphologically by the shape of the solenidion projecting on the genu of the first pair of legs, that is, conical in C. blakei, heart-shaped in C. yasguri, and globus in C. parasitivorax.1,10,11 However, identification based on this feature is difficult due to individual variability in shape and distortion during preparation for microscopic examination. The egg is encased within two layers, an outer chorion and an inner vitelline membrane. An egg buster is located inside the egg and consists of a dorsal pair of lancet-shaped blades, side by side and directed cephalad to aid in the exit of the larval stage.11 Eggs measure 235
245 3 115
135 µm and are attached to the hair shaft at the pole. The life cycle is spent entirely on the host with a prepatent period of 21
35 days. The mite developed through the egg, larva, nymph I, nymph II to the adult. Adults are highly mobile and often resemble large, mobile flakes of dandruff, hence, the phrase “walking dandruff.”

Rearing method(s) No specific rearing methods for Cheyletiella spp. have been described. All models described made use of naturally infested animals. In vitro method(s) No in vitro models have been described in literature either, most likely because longevity of mites off the host is very limited and does not allow development of models. In vivo method(s) Although no specific models or products are available for the treatment of cheyletiellosis, several articles describe the treatment of naturally acquired Cheyletiella infestations of cats and dogs by topical application of acaricides, such as fipronil, topical application of the parasiticide/anthelmintic selamectin on cats, and by oral, injectable, or pour-on ivermectin.12
16 Diagnosis of Cheyletiella mites is diagnosed by the direct examination of the dorsal midline of cats by using a magnifying glass, by microscopic examination of a piece of clear adhesive tape that was applied to the animal’s hair and skin at selected infestation sites, or by microscopic examination of hair and epidermal debris collected with a flea comb. A floatation technique

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using 10% KOH to digest hairs, and a saturated sucrose solution was also described as a diagnostic technique.14

Psorergatidae Psorobia ovis—sheep itch mite Biology and life cycle This is a nonburrowing parasitic mite of sheep geographically distributed in Australia, New Zealand, southern Africa, and North and South America. It is a small mite, roughly circular, and ,0.2 mm in diameter with very stout legs directed radially, giving a crude star-shaped appearance. The life cycle is similar to Psoroptes. All stages are parasitic and take place on the host. The stages comprise the egg, larval, and three nymphal stages (proto-, deuto, and tritonymph) and the adult mite. The entire cycle takes about 2
3 weeks. The mites live in the superficial skin layers causing chronic irritation and skin thickening. Rearing method(s) There are no specific rearing methods apart from normally acquired infestations on sheep. In vitro method(s) No in vitro methods have been developed, most likely because these mites are parasitic during their entire life cycle. In vivo method(s) Parasiticide efficacy studies against Psorobia ovis should be designed and conducted in accordance with the WAAVP guidelines for evaluating the efficacy of acaricides against (mange and itch) mites on ruminants (see Appendix A of Ref. [17]).

Myobiidae Myobia musculi Schranck, 1781—murine fur mite Biology and life cycle Rodent fur mites are a persistent problem in laboratory mouse colonies and can be found in as many as 40% of the mouse facilities.18 Their high prevalence might be due to the relatively small size of the mites making them difficult to diagnose. The life cycle of Myobia musculi is completed in 23 days. The larval period takes 10 days followed by a 5-day nymphal period. Adults are

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observed on the 16th day, produce fertile eggs within 24 h of their appearance, and eggs hatch in 7 days.19

Rearing method(s) No rearing methods have been described for M. musculi in the literature. In vitro method(s) No in vitro parasiticide models have been described for M. musculi. In vivo method(s) As for the other mites described earlier, most in vivo models make use of naturally acquired infested mice. The efficacy of parasiticides has been described in a variety of articles. G¨onenc¸ et al.20 described the efficacy of selamectin against murine fur mites used as a spot-on at 10 mg/kg bw obtaining 100% of efficacy. Apart from topical applications, treatment and eradication of murine fur mites has been evaluated using ivermectin feed at the approximate ingested dose of 1.3 mg/kg for 1, 4, or 8 consecutive weeks. All mice remained free of fur mites for up to 21 weeks PT.18

Trombiculidae Neotrombicula (5Trombicula) autumnalis (Shaw, 1790)—harvest mite Biology and life cycle The geographical range of Neotrombicula autumnalis, the harvest mite, spreads from Western Europe to Southwestern Asia. It is the most abundant trombiculid species in Central Europe and the British Isles. The harvest mite is responsible for producing cutaneous pruriginous lesions in infested hosts. Neotrombicula autumnalis larvae appear red or orange in color and range from 0.2 to 0.4 mm in length. Adults of N. autumnalis are among the largest of the mites and can be up to 2 mm in length. They have a wide back plate and two double eyes and are not sexually dimorphic. The various stages of N. autumnalis include egg, larva, protonymph, deutonymph, tritonymph, and adult. In the spring the adult mites lay their eggs in decomposing organic material, and in only 1 week the eggs hatch into orange-colored larvae. Only the larvae are ectoparasitic on wild and domestic animals and humans, becoming active in autumn, hence, the name “harvest mite.” Larvae quest on the vegetation for a passing host and once picked up, they feed for 5
7 days on enzymatically liquefied tissue, epithelial secretions, or blood. After feeding, they drop off the host and continue their life cycle as free-living stages on the ground. Development to the nymphal stage

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takes 5
6 weeks. The protonymph, as a transitional stage, shows no active movement or feeding activity, whereas the deutonymphs and adults of these mites are soil dwellers that feed upon various arthropods and, in particular, on their eggs, and plant fluids. The tritonymph resumes a resting stage, and adults feed in a similar way as the deutonymphs.21,22

Rearing method(s) The collection of the different stages of N. autumnalis has been described by Cockings23 and Sch¨oler et al.24 The sampling of unengorged larvae from soil can be done using the tile catch method (TCM), bait animals in combination with a light source, and by flagging.25 The TCM uses bathroom tiles, 15 3 15 cm, which are firmly placed on the ground guaranteeing good contact with the soil. Larvae that climb onto the tiles are collected using a moist brush with soft hair. The bait-animal trap method in combination with a light source exposes hosts (hamster or guinea pigs), put in a wooden box with metal gauze on two sides and a light source on one side connected to wooden box by a dark paper funnel, to free host-seeking larvae at ground level.26 The bait animals can be maintained in a cage with wired floor for easy collection of the engorged larvae. Since larvae can be trapped using bait animals, such as hamsters and guinea pigs, it is hypothesized that field-captured larvae could also be fed on these hosts in the laboratory. From field-collected larvae with the bait animals, predilection sites of the mites were the genital area, the anus, perineum, and to a lesser extent the ears. The duration of feeding was estimated to be 30
60 h. Post
larval stages can be sampled from soil samples taken at different depths (between 0 and 45 cm) and collected by flotation from the water surface using warm water ( 6 30 C) after 1 min stirring. The procedure can be repeated several times until no mites are found any more. Although hardly any information is available on N. autumnalis, some attempts have been made to routinely rear Neotrombicula deliensis mites. Fully engorged larval mites can be incubated in tubes filled with moist soil and kept wet for the first 24 h at 30 C
32 C. Thereafter, larvae become nymphophanes in 7 days. In the publication of Cockings,23 several researchers had observed that nymphs and adult mites are predacious. In vitro method(s) No specific in vitro methods for testing parasiticides against N. autumnalis were found in the literature. In vivo method(s) Two Yorkshire Terriers artificially infested with 2000 larvae of N. autumnalis developed paresis in the hind legs, then the forelegs within 24 h PI.1

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Nervous signs disappeared within 3 days after repeated acaricidal (propoxur) and symptomatic therapy.

Straelensia cynotis (Le Net et al., 1999)—chigger mite Biology and life cycle Straelensia cynotis or chigger mite is another mite that caused trombiculosis in dogs and cats, or more specifically, straelensiosis, an emerging disease reported during the last decade from Southern Europe.27
29 The mite’s activity is seasonal, infestations appearing in autumn and often diagnosed in hunting dogs as autumn is the hunting season. The life cycle remains largely unknown but might be similar to that of N. autumnalis although the periods of feeding on the host can take up to 3 months.

Rearing method(s) As the life cycle of S. cynotis is not known, no rearing methods have been described as yet.

In vitro method(s) No in vitro models have been described yet for S. cynotis.

In vivo method(s) As this parasite potentially shows a reduced metabolism, looking at the duration of feeding of the parasitic larval stage, systemic antiparasiticides at recommended or slightly increased dose, mites might not obtain a sufficiently high dose to be controlled.30 Ivermectin at a SC dose of 0.2 mg/kg, twice at monthly intervals, in combination with a selamectin spot-on, three times every fortnight, did not alter the duration of clinical signs in five dogs examined. However, a single application of a combination of amitraz and metaflumizone, at a dose of 25.1 mg/kg of amitraz, did show a regression of the lesions within 2 weeks and complete healing after 3 months.

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3. Santana HJM, Meza DLM, Jaraba LJE, Escaf LC, Lizarazo M, De Egea GG, et al. A novel technique for improving an in vitro culture of Demodex spp (Acari: Demodicidae). A pilot trial. In: Front Immunol Conf Abstract: IMMUNOCOLOMBIA2015
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