Cover
List of Contributors
Foreword
Preface
Part One: Strategies & Resistance
1. Chapter 1: Comparison of Anti‐ectoparasite and Anti‐endoparasite Therapies and Control Strategies
  1. Introduction
  2. Approaches for Ectoparasite and Endoparasite Control
  3. Challenges for Ecto‐ and Endoparasite Control
  4. Perspectives on Current and Future Strategies for Ecto‐ and Endoparasite Control
  5. References
2. Chapter 2: Vaccination Against Ticks
  1. Tick Evolution and Life Cycle
  2. Commercially Available Vaccines
  3. Rational Tick Vaccine Development
  4. Future Developments
  5. References
3. Chapter 3: Blocking Transmission of Vector‐borne Diseases
  1. Introduction
  2. Vector‐borne Transmitted Pathogens
  3. Drug Profile for Blocking Pathogen Transmission
  4. Conclusion
  5. Acknowledgments
  6. References
4. Chapter 4: The Threat and Reality of Drug Resistance in the Cattle Tick Rhipicephalus (Boophilus) microplus
  1. The Cattle Tick Rhipicephalus (Boophilus) microplus
  2. History of Acaricidal Compound Classes: Their Introduction on the Market and Emergence of Resistance
  3. Mechanisms of Resistance in Cattle Ticks
  4. Resistance Management
  5. References
5. Chapter 5: Monitoring Drug Sensitivity in Cattle Ticks
  1. Introduction
  2. How to Identify Acaricide Resistance
  3. Bioassays
  4. Biochemical Tools
  5. Molecular Tools
  6. Use of Resistance Bioassays in the Discovery of New Acaricides
  7. Resistance Management
  8. References
6. Chapter 6: New Developments in the Control of Human Lice
  1. Introduction
  2. Pediculosis and Medical Importance of Head and Body Lice
  3. Pediculicides and Resistance
  4. Development of the In Vitro Rearing System: Maintenance of Insecticide‐susceptible and ‐resistant Strains and Determination of Resistance
  5. Sequencing of the Human Louse Genomes and Transcriptomes
  6. Head Louse Resistance to Pyrethrins, Pyrethroids, and Malathion
  7. Optimization of the Noninvasive Induction Assay to Identify Detoxification Genes Involved in Insecticide Tolerance as a Proactive Resistance Monitoring Approach
  8. New Pediculicides, Infestation Deterrents/Repellents, and Metabolic Synergists with Novel Modes of Action
  9. Sustainable Resistance Management
  10. References
Part Two: Screens & Models
1. Chapter 7: Molecular Targets to Impair Blood Meal Processing in Ticks
  1. Introduction
  2. Processes Associated with Blood Feeding and Digestion
  3. Conclusions
  4. Acknowledgments
  5. References
2. Chapter 8: Whole‐organism Screens for Ectoparasites
  1. Purpose, Advantages, and Limitations of Whole‐organism Screens
  2. General Workflow for Ectoparasiticide Discovery Screens
  3. Basic Principles in Ectoparasite Screen Design
  4. Considerations in Assay Implementation and Screen Execution
  5. Specific Whole‐organism Ectoparasite Assays
  6. Conclusion
  7. References
3. Chapter 9: In vitro Feeding Methods for Hematophagous Arthropods and Their Application in Drug Discovery
  1. Introduction
  2. AFS‐Components: The Membrane
  3. AFS Components: The Blood Meal
  4. AFS Components: Temperature Control Systems
  5. Artificial Feeding Methods and Applications
  6. Artificial Feeding Systems in Ectoparasiticide Drug Discovery
  7. Conclusions
  8. References
4. Chapter 10: Testing in Laboratory Animal Models for Ectoparasiticide Discovery and Development
  1. Introduction
  2. Developing Laboratory Animal Models of Ectoparasite Infestation
  3. Available Ectoparasite Models
  4. Tick Models
  5. Flea Models
  6. Other Ectoparasite Models
  7. Conclusions
  8. Acknowledgments
  9. References
5. Chapter 11: Testing in Target Hosts for Ectoparasiticide Discovery and Development
  1. Role of Animal Models in Ectoparasiticide Discovery and Development
  2. Alignment of Animal Model Assays with Desired Product Characteristics
  3. Specific Animal Model Ectoparasite Assays
  4. Specific Examples of Animal Model Uses in the Development of Ectoparasiticide Products
  5. Extension to Human Conditions
  6. Conclusion
  7. References
Part Three: Isoxazolines
1. Chapter 12: Isoxazolines: A Novel Chemotype Highly Effective on Ectoparasites
  1. Arthropod Ectoparasites: Burden to the Agricultural and Veterinary Sectors
  2. Ligand‐gated Chloride Channels as Suitable Targets for Ectoparasiticides
  3. Mode of Action
  4. Isoxazolines: Novel Ectoparasiticides Acting on GABACls and GluCls
  5. Structure and Active Sites of Chloride Channels
  6. Isoxazoline Mode of Action and Binding Site
  7. Selectivity and Safety Profile
  8. Isoxazoline Derivatives: Continuous Exploration of the Novel Chemotype
  9. Conclusions
  10. Acknowledgments
  11. References
2. Chapter 13: The Discovery of Afoxolaner: A New Ectoparasiticide for Dogs
  1. Introduction
  2. Development of Chemical–Biological Structure–Activity Relationships
  3. Mode of Action
  4. Summary
  5. References
3. Chapter 14: Development of Afoxolaner as a New Ectoparasiticide for Dogs
  1. Introduction
  2. Study Compliance with Regulatory Requirements
  3. Background
  4. Establishing Proof of Concept in Dogs
  5. Dose Level and Formulation Selection
  6. ADME Properties and Pharmacokinetics
  7. Toxicology and Safety of Afoxolaner
  8. Pivotal Dose Determination, Confirmation and Field Trials
  9. NexGard Spectra
  10. Future Direction
  11. Conclusions
  12. Acknowledgments
  13. Disclaimer
  14. References
4. Chapter 15: Discovery, Development, and Commercialization of Sarolaner (Simparica^®), A Novel Oral Isoxazoline Ectoparasiticide for Dogs
  1. Introduction
  2. Discovery of Sarolaner
  3. Laboratory Studies
  4. Flea and Tick Field Studies
  5. Mite Efficacy
  6. Prevention of Tick‐borne Disease Transmission
  7. Comparative Flea and Tick Efficacy
  8. Commercialization of Sarolaner
  9. Acknowledgments
  10. References
5. Chapter 16: Isoxazolines: Preeminent Ectoparasiticides of the Early Twenty‐first Century
  1. Introduction
  2. Patent Literature
  3. Structural Variations
  4. Conclusion
  5. Disclaimer
  6. References
Index
End User License Agreement

List of Tables

Chapter 3
1. Table 3.1 Human vector‐transmitted pathogens.
2. Table 3.2 Companion animal vector‐transmitted pathogens.
3. Table 3.3 Farm animal vector‐transmitted pathogens.
Chapter 8
1. Table 8.1 Ectoparasiticide standards in the mosquito larval assay.
2. Table 8.2 Activity of ectoparasiticide standards in the flea contact assay.
3. Table 8.3 Activity of ectoparasiticide standards in the flea ingestion assay.
4. Table 8.4 Activity of ectoparasiticide standards in the tick contact assay.
5. Table 8.5 Activity of ectoparasiticide standards in the stable fly contact assay.
Chapter 10
1. Table 10.1 Principal features of laboratory models used for drug screening and characterization.
2. Table 10.2 Efficacious doses for known ectoparasiticides in the corresponding laboratory models in comparison with dosages used in the marketed product.
Chapter 12
1. Table 12.1 Ectoparasiticides acting on ligand‐gated chloride channels.
2. Table 12.2 Isoxazoline‐derived parasiticides.
3. Table 12.3 Ectoparasiticide activity (IC50 (nM)) in binding assays on M. domestica head membrane.^a)
Chapter 13
1. Table 13.1 Membrane feeding and oral gavage results for monoamide naphthalene isoxazolines on fleas.
2. Table 13.2 Membrane feeding and oral gavage results for diamide naphthalene isoxazolines on fleas.
3. Table 13.3 Membrane feeding and oral gavage results for diamide naphthalene isoxazolines on fleas.
4. Table 13.4 Tick control by oral gavage.
Chapter 14
1. Table 14.1 Weight bands and active ingredient amounts in the four sizes of NexGard chewable tablets for dogs.
2. Table 14.2 Mean ± standard deviation of the effective concentration for 90% efficacy (EC₉₀) determined using a Sigmoid E_max model for percent efficacy against R. sanguineus, D. variabilis, and A. americanum following oral administration of afoxolaner in a chewable formulation to dogs.
3. Table 14.3 Mean percent efficacy against eight tick species on day 30 or 31 following treatment with NexGard (48‐ or 72‐h counts).
Chapter 15
1. Table 15.1 Comparative in vitro whole parasite efficacy for afoxalaner, fluralaner, and sarolaner fed in blood to C. felis and O. turicata (n = 3).
2. Table 15.2 Summary of dose confirmation laboratory flea efficacy studies [9].
3. Table 15.3 Summary of dose confirmation laboratory tick efficacy studies [10, 11].
4. Table 15.4 Flea efficacy: comparative speed of kill [29–31].

List of Illustrations

Chapter 1
1. Figure 1.1 Consolidation of animal health companies 1990–2016.
Chapter 2
1. Figure 2.1 Schematic representation of the systems that are being targeted to develop vaccines against tick infestation.
2. Figure 2.2 Simplified schematic of the blood coagulation processes that are modulated by tick proteins. Compounds in green fields and printed in italics are targets of tick serine protease inhibitors. Compounds in blue fields are substrates of tick proteases. Compounds in yellow are activated by tick proteins. ADP, adenosine diphosphate; VWF, von Willebrand factor.
Chapter 3
1. Figure 3.1 Generic sketch of transmission of diseases by ectoparasites (vectors). Blocking of transmission can, in principle, occur at every stage, but most drugs aim to interfere during the “Attachment” phase and/or “Feeding and Transmission” phase.
Chapter 4
1. Figure 4.1 Chronological order of introduction of acaricides for use against cattle ticks (green date markers) and the first documentation of resistance against the respective class (red date markers).
Chapter 5
1. Figure 5.1 The cattle tick R. (B.) microplus has attachment sites where it may not be easily detected. In the present case, a heifer was captured on pasture and laid down for determination of the tick burden. Using such a technique before and after acaricide treatment to identify resistance is very resource intensive and time consuming.
2. Figure 5.2 Schematic depiction of the most commonly used bioassays for in vitro identification of cattle tick resistance to acaricides. (a). AIT: Adult engorged female ticks are immersed in defined dilutions of acaricide. The ticks are dried, placed in Petri dishes and kept at 27 to 28 °C and 80–95% relative humidity (RH). After 2 weeks, eggs are weighed, transferred to tubes, and incubated at the before‐mentioned climatic conditions for ~3 additional weeks to evaluate the larval hatching. In a simplified protocol, oviposition is evaluated approximately 1 week after incubation, without further evaluation of the larval hatching. (b). LPT: Tick larvae are incubated at 27 to 28 °C and 85–95% RH for 24 h in filter paper pouches that were impregnated with acaricides. The pouches are opened to visually determine the viability of the larvae. (c). LIT: The tick larvae are immersed for ~10 min in defined concentrations of acaricide. This is followed by a transfer into filter paper pouches (no impregnation with acaricide) in which the larvae are incubated for 24 h at 27 to 28 °C and 80–90% RH. The viability of the larvae is determined directly in the pouch by removing the clips, analogous to the LPT. (d). LTT: Tick eggs are placed into 96‐well flat bottom plates which were pre‐coated with defined concentrations of acaricides. The plates are sealed and incubated at 27 to 29 °C and 70–80% RH. Two weeks after egg hatching the larval mortality is determined in each well.
3. Figure 5.3 Position of the so far identified mutations in the voltage‐gated sodium channel of R. (B.) microplus, which lead to resistance to synthetic pyrethroids. Three mutations lead to changes in the S4‐S5 linker of domain II: T170C (red) [20], C190A (green) [21], and G215T (blue) [22]; and one in the S6‐region of domain III: T2134A (violet) [23].
Chapter 6
1. Figure 6.1 Assembly of the in vitro rearing system for lice. ()
2. Figure 6.2 Comparative sodium current traces from the house fly VSSC variants with and without head louse mutations expressed in Xenopus oocytes before and after exposure to increasing concentrations of permethrin. ()
3. Figure 6.3 Relative transcript levels (panel a and b) and mortality responses (panel c and d) of body louse females to a lethal contact amount of ivermectin (5% IVM) following injection of dsRNA targeting either louse CYP9AG2 or ABCC4. Lice were also injected with either dsRNA of the odd‐paired gene, opa, (GeneBank accession # S78339) for P450 silencing or with dsRNA of the E. coli plasmid, pQE30, for ABC transporter silencing as sham injected controls. Asterisks (*) in panels a and b indicate that CYP9AG2 and ABCC4 dsRNA significantly suppress the levels of CYP9AG2 and ABCC4 transcripts, respectively (Student’s t‐test, P < 0.05). In panel c, the bioassay was started 48 h after CYP9AG2 dsRNA injection. In panel d, the bioassay was started 12 h after ABCC4 dsRNA injection. Asterisks (*) in panels c and d indicate that the mortality responses of lice injected with dsRNAs were significantly different from their respective controls (buffer or water only injected, maximum log‐likelihood ratio test, P < 0.05). ()
Chapter 7
1. Figure 7.1 Physiological processes associated with blood meal processing in ticks. A schematic overview of tick tissues and related processes of blood meal uptake, digestion, heme and iron metabolism, detoxification and inter‐tissue transport that may serve as rational targets for “anti‐tick” intervention. The red and blue arrows indicate blood meal uptake and reverse water secretion, respectively.
2. Figure 7.2 The current model of uptake and digestion of major host blood proteins by Ixodes ricinus digestive cells. Red arrows – expressional and secretory pathway of hydrolases involved in protein digestion. Black arrows – endocytic pathways of hemoglobin and serum albumin. RME – receptor‐mediated endocytosis of hemoglobin; FPE – fluid‐phase endocytosis of albumin (and other serum proteins). HbR – hemoglobin receptor (putative, yet not identified); CP – coated pits, E‐l – large endosomal vesicles containing hemoglobin; E‐s – small endosomal vesicles containing dissolved serum albumin [40]; Hs – hemosomes containing condensed heme [42]; Golgi – Golgi apparatus; ER – endoplasmic reticulum; PL – primary lysosomes. IrAE – I. ricinus legumain/AE; IrCD – I. ricinus cathepsin D; IrCB – I. ricinus cathepsin B; IrCC – I. ricinus cathepsin C; IrSCP – I. ricinus serine carboxypeptidase; IrLAP – I. ricinus leucine aminopeptidase. For details, see the text.
3. Figure 7.3 A model of putative tick iron and heme metabolic pathways. Iron and heme pathways in ticks are independent as ticks are not capable of heme degradation given the absence of heme oxygenase. Iron is most likely acquired from the host serum transferrin digested in the acidic environment of small endosomal vesicles (E‐s) of midgut digestive cells. Upon reduction, ferrous iron is transported from the lysosome by the divalent metal transporter 1 (Dmt1). Once in the cytoplasm, Fe²⁺ ions are scavenged by intracellular ferritin 1 (Fer1), whose translation is strictly regulated by the cytoplasmic Aconitase/Iron responsive protein1 (Aco/IRP1), which senses the iron cellular level. Iron destined to be delivered to the peripheral tissues is transported from digestive cells via DMT1 to the hemolymph and bound to the iron secreted ferritin 2 (Fer2) that functions as an iron transporter. The excessive iron in peripheral tissues is scavenged and stored in Fer1. Heme released from the digested host hemoglobin in the large endosomal vesicles (E‐l) is transferred to the cytoplasm via the heme responsive gene 1 (HRG‐1) transporter. Ticks detoxify the majority of acquired heme by an ABC transporter (ABC)‐mediated transport to hemosomes. Glutathione‐S‐transferase(s) (GST) serve as an intracellular scavenger of free heme. A small portion of acquired heme required for proteosynthesis of endogenous hemoproteins is exported from the digestive cells to the hemocoel by FLVCR transporter. In hemolymph, heme is bound by the abundant carrier protein(s), heme‐lipo‐glycoprotein (HeLp), which serves in all developmental stages both as a scavenger of excessive heme and transporter into peripheral tissues. In the post‐repletion period of fully engorged females, most of the heme is bound to vitellogenins (Vg) and transported to the ovaries to supply heme metabolic demands of developing embryos and larvae. After entry into the developing oocytes via vitellogenin receptor (VgR), Vg is proteolytically processed to vitellins (Vns) by aspartic proteases BYC and THAP, and cathepsin L‐like activity of VTDCE. For details, see the text.
Chapter 8
1. Figure 8.1 Ectoparasiticide discovery flow diagrams: (a) Part 1; (b) Part 2.
2. Figure 8.2 Typical 96‐well plate setup for mosquito larval assay.
3. Figure 8.3 (a–c) Apparatus for flea contact assay.
4. Figure 8.4 (a) Flea eggs, larvae, fras in Petri dish. (b) Flea larvae in Petri dish.
5. Figure 8.5 (a) Ticks in shipping container. (b) Ticks on white paper.
6. Figure 8.6 (a) Dv larvae nymph, adult. (b) Vial on hotdog roller. (c) Amitraz effect.
7. Figure 8.7 A Cheyletiella spp. mite.
8. Figure 8.8 (a) Adult flies in Bugdorm^® cage. (b) Flies in the Petri dish assay.
9. Figure 8.9 (a) Fly capture system. (b) Setup to anesthetize the flies.
Chapter 9
1. Figure 9.1 The “Greyhound” artificial feeding system. ()
2. Figure 9.2 Rutledge‐type feeder. The Rutledge feeding system consists of a jacketed hollow cone, the base of which can be covered by an artificial membrane or animal skin (a). Blood is introduced into the cone through a tube (b) that extends from its vertex. Using a circulating water bath, heated water is pumped through the cylindrical water jacket that surrounds the cone and tube (c), thereby warming the blood meal. ()
3. Figure 9.3 Example of an in vitro flea feeding system, showing three Rutledge‐type feeders (a) attached to laboratory support stands (b) and connected in series to tubing attached to a circulating water bath. The continuous flow of warm water heats the blood meal inside the Rutledge feeders to approx. 37°C. Flea cages (c) are brought into contact with the Parafilm membrane covering the blood meal using a laboratory jack (d). Each flea cage has a feeding screen which directly touches the Parafilm membrane and a bottom screen which allows flea eggs and feces to fall through in a dish containing culture media (e), from where the eggs can be harvested. ()
4. Figure 9.4 An in vitro feeding unit modified after Ref. [53]. It consists of glass tubing and mosquito netting (N) glued to a silicone membrane (M) which closes the unit on one side. Additional attachment stimuli such as hair or hair extracts can be placed on the membrane. A movable rubber ring (R) around the unit keeps the blood meal below the membrane when the feeding unit is placed in a blood container such as a glass beaker or the well of tissue culture plate. A perforated plastic stopper wrapped with muslin (S) is inserted in the unit to confine the ticks during feeding. ()
Chapter 10
1. Picture 10.1 Non‐murine rodents like gerbils can be used successfully for evaluating ectoparasiticides. To our experience, they are easy to handle and are very good hosts for ticks and cat fleas.
2. Figure 10.1 Tick rat model, diagrammed based on description by Gutierrez et al., 2006 [18].
Chapter 11
1. Figure 11.1 Prototypical workflow and decision tree for ectoparasiticide development.
Chapter 12
1. Figure 12.1 Schematic representation of the transmembrane domain of a ligand‐gated chloride channel. (a) Four transmembrane helices (M1–M4) that make up one subunit are depicted with their connecting loops. (b) Arrangement of the transmembrane subunits to form a pentameric ion channel with M2 Cl⁻ helices (orange) lining the pore, M1 and M3 linking the adjacent subunits, and M4 facing the outside of the pore. Helices M3^′ and M4^′ of the foremost subunit have been omitted for clarity. The composition of helices is illustrated for only three of the five subunits.
2. Figure 12.2 Homology model of GABACl in a proposed closed form. Sequences of Erwinia chrysanthemi GABACl and dlr‐GABACl of Ctenocephalides felis were aligned using the BLAST algorithm, and final modeling was performed with the MOE software package [41]. The five individual subunits are shaded in different colors. The Cys loop of one subunit is highlighted in light green (light‐green arrows), the resistance‐inducing mutation in dark green (dark‐green arrow). (a) Protein shown parallel to membrane; horizontal gray bars indicate membrane boundaries. (b) View along the channel pore axis from the extracellular side, with M2 helices visible as the inner pore lining. (c) View along the channel pore axis from the intracellular side, with resistance‐inducing residue (red dot) highlighted with a red arrow.
3. Figure 12.3 GluCl crystal structure of C. elegans represented in an activated, open‐channel state. These images are based on an original X‐ray crystallographic structure (PDB ID: 3RHW) [49] and were produced with the MOE software package [41]. Fab molecules are omitted for clarity. The five individual subunits are shaded in different colors. The surface of one ivermectin‐binding pocket is highlighted in yellow. The Cys loop of one subunit is highlighted in light green (light‐green arrows). (a) Protein parallel to membrane, horizontal gray bars indicate membrane boundaries. (b) View along the channel pore axis from the extracellular side, with M2 helices visible as the inner pore lining. (c) View along channel pore axis from the intracellular side.
4. Figure 12.4 Schematic representation of CysLGCCs in a sectional view with only three units of the pentameric transmembrane region depicted for clarity. Positioning of isoxazolines is putative.
Chapter 13
1. Figure 13.1 Isoxazoline insecticide afoxolaner.
2. Scheme 13.1 Lead compounds in the discovery of afoxolaner.
3. Scheme 13.2 Synthesis of 4. (a) nBuLi, DMF, THF; (b) NH₂OH, EtOH; (c) NCS, Et₃N, DMF; (d) CO, PdCl₂dppf, NH₂CH₂CF₃, toluene.
4. Scheme 13.3 Synthesis of 14. (a) NCS, Et₃N, DMF; (b) PdCl₂dppf, CO, NH₂CHR₃CO₂Me, Et₃N, toluene; (c) LiOH, THF/H₂O (1 : 1) then HCl; (d) (COCl)₂, DMF, CH₂Cl₂, Et₃N, NH₂R_2.
5. Scheme 13.4 Synthesis of DD‐16 and DD‐17. (a) CO, PdCl₂dppf, MeOH, Et₃N, toluene; (b) LiOH, THF/H₂O then HCl; (c) oxalyl chloride; (d) (R)‐1‐(methylthio)propan‐2‐amine, Et₃N, CH₂Cl_2; (e) HPLC separation of diastereomers by chiral OJ‐RH column with MeOH/CH₃CN as eluent
6. Scheme 13.5 Synthesis of 22. (a) Pd(PPh₃)₄, 4 N KOH, THF/DME, 80 °C; (b) CuCN, NMP 150 °C 15 h; (c) NH₂OH, EtOH, rt, 3 h; (d) NaOCl, THF, 0 °C to rt, 30 min; (e) triazole, K₂CO3, CH₃CN, 80 °C, 18 h.
7. Figure 13.2 CPD I (22) inhibits GABA‐gated currents in American cockroach, P. americana, neurons. Dissociated neurons were clamped at a holding potential of −60 mV and repeatedly stimulated with pulses of 100 μM GABA (inset, solid trace). Perfusion of CPD I (22) inhibited the GABA response (inset, dashed trace) in a dose‐dependent manner with an IC₅₀ = 10.8 nM. Following prolonged saline rinse, a partial recovery of the GABA response (inset, dotted trace) was observed. ()
8. Figure 13.3 Inhibitory effect of afoxolaner, on GABA‐gated Cl⁻ currents recorded from Xenopus oocytes expressing either ^wtRDL or ^A302SRDL(resistant) receptors. Oocytes were recorded using TEVC (two electrode voltage clamp method) with a holding potential of ‐60 mV. ()
9. Figure 13.4 Contact toxicity of afoxolaner (square), fipronil (triangle), and dieldrin (circle) against wild‐type (Canton‐S, closed symbol) and cyclodiene‐resistant (Rdl, open symbol) strains of Drosophila. Mortality measurements were taken 72 h after flies were transferred to treated glass vials. The resistance ratio (RR) was calculated as Rdl LD₅₀/Canton‐S LD₅₀ for each compound.
Chapter 14
1. Figure 14.1 Percent efficacy of afoxolaner against Rhipicephalus sanguineus following a single oral treatment to dogs at 3 dose levels: 1.5, 2.5, and 3.5 mg/kg.
2. Figure 14.2 Percent efficacy against Dermacentor variabilis versus afoxolaner plasma concentration following oral administration of afoxolaner in a chewable formulation to Beagle dogs.
3. Figure 14.3 Proposed pathway for the formation of the hydroxylated metabolite of afoxolaner.
4. Figure 14.4 Percent efficacy of afoxolaner against C. felis following a single oral treatment of NexGard to dogs.
5. Figure 14.5 Efficacy of NexGard against existing flea (C. felis) infestations on dogs.
6. Figure 14.6 Efficacy results from NexGard field trials.
Chapter 15
1. Figure 15.1 Structure of sarolaner (1‐(5′‐((5S)‐5‐(3,5‐dichloro‐4‐fluorophenyl)‐5‐(trifluoromethyl)‐4,5‐dihydroisoxazol‐3‐yl)‐3′‐H‐spiro(azetidine‐3,1′‐(2) benzofuran)‐1‐yl)‐2‐(methylsulfonyl) ethanone): (1) phenyl head group, (2) isoxazoline core, (3) spiroazetidinebenzofuran moiety, and (4) methylsulfonylethanone tail.
Chapter 16
1. Scheme 16.1 Diamide starting point resulting in divergent compounds.
2. Figure 16.1 Other commercial isoxazolines for use against fleas and ticks.
3. Figure 16.2 Potential first commercial isoxazoline for crop protection.
4. Scheme 16.2 Two general pathways for the synthesis of parasiticidal isoxazolines.
5. Scheme 16.3 Fractional crystallization and recycling processes in the synthesis of Lotilaner.
6. Scheme 16.4 An alternate route to access enantiomerically enriched antiparasitic isoxazolines.
7. Figure 16.3 Lipophilicity and molecular weight of representative compounds.
8. Figure 16.4 Hydrogen bond acceptors and donors for representative compounds.
9. Figure 16.5 Calculated solubility and polar surface area for representative compounds.
10. Figure 16.6 General structures for literature analyses.
11. Figure 16.7 Patent applications 2005–present.
12. Figure 16.8 Common substitution patterns on the isoxazoline core.
13. Figure 16.9 Phenyl group replacements.
14. Figure 16.10 Phenyl group replacements.
15. Figure 16.11 Common amide substitutions.
16. Figure 16.12 Amide‐related functional groups in isoxazoline analogs.
17. Figure 16.13 Alternative substitutions para to the isoxazoline.
18. Figure 16.14 Thio substitutions in lieu of an amide.
19. Figure 16.15 Ketone substitutions on the phenyl linker.
20. Figure 16.16 Substitutions on the isoxazoline linker.
21. Figure 16.17 Single‐atom variants of isoxazolines.
22. Figure 16.18 Additional isoxazoline replacement rings.
23. Figure 16.19 Fused ring isoxazoline replacements.
24. Figure 16.20 Acrylamide replacements for the isoxazoline ring.
25. Figure 16.21 “Open ring” isoxazoline replacements from Dow Agrosciences.
26. Figure 16.22 Vinyl‐extended phenyl isoxazolines from Syngenta.

List of Contributors

Eric A. Benner
Stine‐Haskell Research Center
DuPont Crop Protection
1090 Elkton Rd.
Newark, DE 19714
USA

John M. Clark^*
University of Massachusetts
Department of Veterinary & Animal Science
Amherst, MA 01003
USA
jclark@vasci.umass.edu

Jeffrey N. Clark^*
JNC Consulting Services
38 Wild Rose Lane
Pittsboro, NC 27312
USA
depotrd33@yahoo.com

Pat N. Confalone
Stine‐Haskell Research Center
DuPont Crop Protection
1090 Elkton Rd.
Newark, DE 19714
USA

Daniel Cordova
Stine‐Haskell Research Center
DuPont Crop Protection
1090 Elkton Rd.
Newark, DE 19714
USA

Andrew A. DeRosa
Zoetis, Veterinary Medicine Research and Development
333 Portage St. Kalamazoo
MI 49007
USA

Robert F. Dietrich
Stine‐Haskell Research Center
DuPont Crop Protection
1090 Elkton Rd.
Newark, DE 19714
USA

Christian Epe
Boehringer Ingelheim Animal Health
3239 Satellite Blvd
Duluth, GA 30096
USA

Brandon R. Gould
Stine‐Haskell Research Center
DuPont Crop Protection
1090 Elkton Rd.
Newark, DE 19714
USA

Ondřej Hajdušek
Czech Academy of Sciences
Institute of Parasitology Biology Centre
České Budějovice
Czech Republic

Eric J. Hartline
Stine‐Haskell Research Center
DuPont Crop Protection
1090 Elkton Rd.
Newark, DE 19714
USA

Gail S. Jones
DuPont Crop Protection
Stine‐Haskell Research Center
1090 Elkton Rd.
Newark, DE 19714
USA

Ronald Kaminsky^*
paraC Consulting for Parasitology and Drug Discovery
79685 Haeg‐Ehrsberg
Germany
para.C@gmx.de

John B. Kinney
DuPont Crop Protection
Stine‐Haskell Research Center
1090 Elkton Rd.
Newark, DE 19714
USA

Petr Kopáček^*
Czech Academy of Sciences
Institute of Parasitology Biology Centre
Czech Academy of Sciences
České Budějovice
Czech Republic
kopajz@paru.cas.cz

George P. Lahm^*
Stine‐Haskell Research Center
DuPont Crop Protection
1090 Elkton Rd.
Newark, DE 19714
USA
george.lahm@fmc.com

Diane Larsen
Boehringer Ingelheim Animal Health
3239 Satellite Blvd
Duluth, GA 30096
USA

Laura Letendre^*
Boehringer Ingelheim Animal Health
631 Route 1 South
North Brunswick, NJ 08902
USA
laura.letendre@boehringer‐ingelheim.com

Alan Long^*
Boehringer Ingelheim Animal Health
3239 Satellite Blvd
Duluth, GA 30096
USA
Al.Long@merial.com

Jeffrey K. Long
Stine‐Haskell Research Center
DuPont Crop Protection
1090 Elkton Rd.
Newark, DE 19714
USA

Léonore Lovis
Elanco Animal Health
Mattenstrasse 24A
4058 Basel
Switzerland

Michael Mahaffey
Stine‐Haskell Research Center
DuPont Crop Protection
1090 Elkton Rd.
Newark, DE 19714
USA

Richard G. McDowell
Stine‐Haskell Research Center
DuPont Crop Protection
1090 Elkton Rd.
Newark, DE 19714
USA

Tom L. McTier
Zoetis, Veterinary Medicine Research and Development
333 Portage St. Kalamazoo
MI 49007
USA

Ard M. Nijhof^*
Freie Universität Berlin
Institute for Parasitology and Tropical Veterinary Medicine
Robert‐von‐Ostertag‐Str. 7–13
14163 Berlin
Germany
ArdMenzo.Nijhof@fu‐berlin.de

Sandra Noack
Boehringer Ingelheim Animal Health
Binger Str. 173
55216 Ingelheim
Germany

Thomas F. Pahutski
Stine‐Haskell Research Center
DuPont Crop Protection
1090 Elkton Rd.
Newark, DE 19714
USA

Cedric J. Pearce
Mycosynthetix, Inc.
505 Meadowland Drive
Suite 103
Hillsborough, NC 27278
USA

Jan Perner
Czech Academy of Sciences
Institute of Parasitology Biology Centre
Czech Academy of Sciences
České Budějovice
Czech Republic

Daniel F. Rhoades
Stine‐Haskell Research Center
DuPont Crop Protection
1090 Elkton Rd.
Newark, DE 19714
USA

Heinz Sager^*
Elanco Animal Health
Mattenstrasse 24A
4058 Basel
Switzerland
sager_heinz@elanco.com

Theo P. M. Schetters^*
ProtActivity R&D
Cuijk
The Netherlands
and
Université de Montpellier
Laboratoire de Biologie Cellulaire et Moléculaire
Montpellier
France
th.schetters@protactivity.com

Sandra Schorderet‐Weber^*
Sablons 30
2000 Neuchâtel
Switzerland
sandra.schorderet@hotmail.com

Mark E. Schroeder
Stine‐Haskell Research Center
DuPont Crop Protection
1090 Elkton Rd.
Newark, DE 19714
USA

Paul M. Selzer^*
Boehringer Ingelheim Animal Health
Binger Str. 173
55216 Ingelheim
Germany
paul.selzer@boehringer‐ingelheim.com

Rafael Shapiro
Stine‐Haskell Research Center
DuPont Crop Protection
1090 Elkton Rd.
Newark, DE 19714
USA

Wesley L. Shoop
Stine‐Haskell Research Center
DuPont Crop Protection
1090 Elkton Rd.
Newark, DE 19714
USA

Radek Šíma
Czech Academy of Sciences
Institute of Parasitology Biology Centre
Czech Academy of Sciences
České Budějovice
Czech Republic

Ben K. Smith
Stine‐Haskell Research Center
DuPont Crop Protection
1090 Elkton Rd.
Newark, DE 19714
USA

Daniel Sojka
Czech Academy of Sciences
Institute of Parasitology Biology Centre
Czech Academy of Sciences
České Budějovice
Czech Republic

Mark Soll
Boehringer Ingelheim Animal Health
3239 Satellite Blvd
Duluth, GA 30096
USA

Katharine R. Tyson
AgBiome
Research Triangle Park, NC 27709
USA

Molly E. Waddell
Stine‐Haskell Research Center
DuPont Crop Protection
1090 Elkton Rd.
Newark, DE 19714
USA

Ty Wagerle
Stine‐Haskell Research Center
DuPont Crop Protection
1090 Elkton Rd.
Newark, DE 19714
USA

Tina Weber
Merck KGaA
Frankfurter Str. 250
64293 Darmstadt
Germany

Debra J. Woods^*
Zoetis, Veterinary Medicine Research and Development
333 Portage St. Kalamazoo
MI 49007
USA
debra.j.woods@zoetis.com

Ming Xu
Stine‐Haskell Research Center
DuPont Crop Protection
1090 Elkton Rd.
Newark, DE 19714
USA

Preface

Infestation by ectoparasites has plagued humans, figuratively and literally, since ancient times; for example, lice are listed among the Biblical plagues visited upon Egypt (Exodus 8:17, KJV) and fleas transmitting bubonic plague have had devastating impacts on numerous civilizations over the centuries. Strategies for battling ectoparasites have an equally deep history, as evidenced by mummified lice found in ancient Egyptian combs and by perforated necklace beads that doubled as personal flea traps in medieval Europe. Although human ectoparasite infestations are less prevalent in modern developed countries due to dramatically improved living and hygiene conditions, infestation on domesticated animals remains a major challenge, causing nuisance in companion animals and livestock as well as lowering livestock productivity. Ectoparasites can move between animals and from animals to humans, potentially transmitting various diseases in the process. Ectoparasite control strategies must therefore contend with the ability of the target to move, often quite quickly, as anyone who has ever wanted to kill a flea can attest. This eighth volume in the Drug Discovery for Infectious Diseases series reviews strategies and models for discovery and development of ectoparasiticidal treatments for use in both human and animal health. The challenges presented by moving targets are a common theme throughout, ranging from the market requirement for a rapid speed of kill to the design of effective containment strategies in whole‐organism drug screening assays.

The first section of the volume, Strategies & Resistance, presents various perspectives on what is needed to achieve effective therapeutic control of ectoparasite infestations. The section begins with a comparison by Woods et al. of therapeutic strategies against moving target ectoparasites with those against the less‐mobile endoparasites. Weber et al. review strategies for preventing disease transmission by ectoparasite vectors, for which speed of kill is an important consideration. Schetters reviews promising progress toward development of vaccines against ticks. The emergence of drug resistance threatens the utility of ectoparasiticides, especially for cattle tick and human head lice. Sager et al. and Lovis et al. discuss the threat, reality, and monitoring of drug resistance in cattle tick, particularly relevant for Southern Hemisphere markets such as Brazil and Australia. Clark reviews new developments in the control of human lice.

The second section focuses on laboratory screens and in vivo models for discovery of new treatments against ectoparasites. Compared to human diseases, the molecular targets of parasites, especially ectoparasites, are much less clear, and few can be utilized for screening. The chapter by Kopáček considers the challenges in identifying candidate small‐molecule drug targets in ticks. Currently, discovery of new treatments against ectoparasites relies heavily on phenotypic‐based screening against whole organisms such as fleas and ticks. Chapters by Clark and Pearce and by Nijhof and Tyson discuss the design and implementation of various whole‐organism assays to detect different aspects of the desired treatments, for example, the flea ingestion assay to detect the ability of a compound to work through ingestion rather than through contact. Compared to drug discovery for humans, a major advantage of drug discovery for animal health is that a new investigative drug can be tested in the target host much sooner in the latter. This might seem to make testing in rodent models less critical. However, testing in rodent models remains an important step in drug discovery for animal health, because these models require much less quantity of a compound and save valuable animals of the target species, as discussed in depth by Weber et al. Of course, testing in the target host species is an essential aspect of late‐stage development of a new drug for animal health, and in the concluding chapter of this section Clark reviews protocols for controlled laboratory testing in host species and provides numerous examples of how these testing strategies have been applied in successful ectoparasiticide development programs.

Drugs effective against ectoparasites comprise only a few chemical classes, the pyrethroids, the phenylpyrazoles, and the macrocyclic lactones being the major ones. On average a new class appears about every 20 years. The isoxazolines are the most recent addition to the roster. The last section of this volume is devoted exclusively to this fascinating new class of ectoparasiticides, which has attracted tremendous interest in the animal health and crop protection industries. Weber and Selzer first discuss the new mode of action that underlies the rapid speed of kill by the isoxazolines. Chapters by Lahm et al. and by Letendre et al. detail the complete drug discovery and development process for afoxolaner, the first commercial product launched from this class. The development of sarolaner, reviewed by Woods and McTier, gives another story from a different setting. The final chapter by Long presents a comprehensive overview of the entire isoxazoline chemical class to date.

We thank Dr. Paul M. Selzer, the series editor, and the various representatives of Wiley for the opportunity to shepherd this volume, and for their guidance and support. We also thank the authors who have generously contributed their time and expertise. The combined result of their efforts is a volume designed to be of both interest and utility to those scientists in academia and industry willing to undertake the discovery of drugs aimed at moving targets.

April 2018

Charles Q. Meng

Duluth, Georgia

Ann E. Sluder

Boston, MA

Ectoparasites

Drug Discovery Against Moving Targets

Copyright

List of Contributors

Note

Foreword

Preface

Part One
Strategies & Resistance