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EN58CH06-Casida ARI 5 December 2012 8:11

Neuroactive : Targets, Selectivity, Resistance, and Secondary Effects

John E. Casida1,∗ and Kathleen A. Durkin2

1Environmental Chemistry and Toxicology Laboratory, Department of Environmental Science, Policy, and Management, 2Molecular Graphics and Computational Facility, College of Chemistry, University of California, Berkeley, California 94720; email: [email protected], [email protected]

Annu. Rev. Entomol. 2013. 58:99–117 Keywords The Annual Review of Entomology is online at , calcium channels, GABAA receptor, nicotinic ento.annualreviews.org receptor, secondary targets, sodium channel This article’s doi: 10.1146/annurev-ento-120811-153645 Abstract Copyright c 2013 by Annual Reviews. Neuroactive insecticides are the principal means of protecting crops, people, All rights reserved livestock, and pets from pest insect attack and disease transmission. Cur- ∗ Corresponding author rently, the four major nerve targets are acetylcholinesterase for organophos- phates and methylcarbamates, the nicotinic receptor for , the γ-aminobutyric acid receptor/chloride channel for by Public Health Information Access Project on 04/29/14. For personal use only.

Annu. Rev. Entomol. 2013.58:99-117. Downloaded from www.annualreviews.org polychlorocyclohexanes and fiproles, and the voltage-gated sodium channel for and dichlorodiphenyltrichloroethane. Species selectivity and acquired resistance are attributable in part to structural differences in binding subsites, receptor subunit interfaces, or transmembrane regions. Additional targets are sites in the sodium channel ( and metaflumizone), the glutamate-gated chloride channel (), the octopamine receptor ( metabolite), and the calcium-activated calcium channel (diamides). Secondary toxic effects in mammals from off-target serine hydrolase inhibi- tion include -induced delayed neuropathy and disruption of the cannabinoid system. Possible associations between pesticides and Parkinson’s and Alzheimer’s diseases are proposed but not established based on epidemiological observations and mechanistic considerations.

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WHY NEUROACTIVE INSECTICIDES? Insecticides are principal defenses against insect pests of crops, livestock, pets, and people. Most PCCA: insecticides are nerve poisons and have been since dichlorodiphenyltrichloroethane (DDT) and polychlorocycloalkane various polychlorocycloalkanes (PCCAs) were introduced in the 1940s, followed by organophos- Target site: enzyme, phates (OPs) in the 1950s, methylcarbamates (MCs) in the 1960s, pyrethroids in the 1970s, and receptor, or channel neonicotinoids in the 1990s (26, 138). Neurotoxicants are the major synthetic insecticides for site at which specific several reasons (28). They act rapidly to stop crop damage and disease transmission. There are binding initiates the many sensitive sites at which even a small disruption may ultimately prove to be lethal. A lipoidal physiological change sheath protects the insect nerve from ionized toxicants but not from lipophilic insecticides. Poor Cross-resistance: detoxification mechanisms in nerves provide prolonged toxicant effects. target site (or metabolic) resistance The nervous system has at least 11 targets or defined modes of action for neuroactive in- conferred by a secticides (7, 8, 23, 26, 32) (Figure 1, A–K). Primary target site selectivity between insects and common target (or people plays an important role in safety, but secondary targets must also be considered. detoxification Selection of pest populations for target site resistance and cross-resistance is delayed or circum- mechanism) vented by shifting modes of action. A diversity of targets is therefore critical for insect and in- Secondary effect: secticide management (95). This review considers the targets and species selectivity of neurotoxic toxicity from action at insecticides, the resistance mechanisms in insects, and the secondary effects in mammals, with a secondary target emphasis on research by the authors. Structures for the insecticides mentioned here are given in Ordeal poison: recent manuals or reviews (8, 95, 138). a poison given to the accused to determine guilt (dies) or innocence (lives) ACETYLCHOLINESTERASE

Primary Target and Inhibitor Action The toxicity of OPs and MCs to insects and mammals is attributable to inhibition of acetyl- cholinesterase (AChE), which is responsible for the hydrolysis of acetylcholine (ACh) at synaptic regions of nerve endings (19, 46, 74) (Figure 1). AChE inhibitors cause ACh to accu- mulate, resulting in excessive stimulation of cholinergic receptors. The MC from calabar beans (Physostigma venenosum) was used first as an ordeal poison in West Africa and then to establish the relationships between ACh neurotransmission and AChE activity and its inhibition (20). AChE inhibition by OP and MC insecticides involves phosphorylation and carbamoylation, respectively, of serine in the esteratic site (Figures 1, 2). AChE is inhibited by MCs as the re- versible AChE–MC complex or as the methylcarbamoylated enzyme at serine that reactivates spontaneously over a short time span (75). The corresponding reaction with OPs yields the phos- by Public Health Information Access Project on 04/29/14. For personal use only. Annu. Rev. Entomol. 2013.58:99-117. Downloaded from www.annualreviews.org phorylated AChE that undergoes aging or reactivates slowly, except with an oxime reactivator such as (19, 46).

−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−→

Figure 1 Proposed sites of neuroactive insecticide action on nerve impulses, neuromuscular transmission, and synaptic receptors adapted from Bloomquist (8), Casida & Quistad (28), and many other sources. The presynaptic nerve terminals (–, ) and the postsynaptic cell (–, ) or muscle cell () are shown. In the nicotinic and γ-aminobutyric acid (GABA) receptor/channel cross-sections (, ), four of the five subunits and one binding site are arbitrarily shown per channel. The number of commercial insecticides or prototype natural products (number of compounds) (95) that act at each site (A–K) are given in parentheses. Abbreviations: ACh, acetylcholine; DDT, dichlorodiphenyltrichloroethane; MC, methylcarbamate; nAChR, nicotinic acetylcholine receptor; OP, organophosphate; PCCA, polychlorocycloalkane.

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Neurotransmitters 11. AChE and nicotinic receptor/cation channel excitatory synapse ACh CH C(O)OCH CH N+(CH ) AChE A 3 2 2 3 3 ACh + acetate K+ ACh Excitatory ACh GABA NH2CH2CH2CH2C(O)OH ACh postsynaptic ACh ACh potential Extracellular Vesicles B Na+ Octopamine HO CHOHCH2NH2 ACh ACh ACh Action potential Intracellular OO BD– + Na Glutamate HO OH

NH2 22. GABA receptor/CI– channel inhibitory synapse

GABA Insecticides (number of compounds) G opens G GABA OP (65), MC (26), physotigmine: Vesicles Extracellular A AChE inhibitors Cl– F G G Nicotine, neonicotinoids (7): Inhibitory B nAChR agonists G = GABA postsynaptic potential E Intracellular analogs (4): Cl– EF– Block of action potential C nAChR blocker

33. Glutamate receptor excitatory synapse Spinosyns (2): nAChR allosteric D activators

Glu Fiproles (2), PCCAs (2), picrotoxin: Na+ E noncompetitive antagonists, blockers and convulsants Ca++ Excitatory postsynaptic potential Avermectins (4): activators, F modulators Glu Cl – F DDT, pyrethroids, , G veratridine (44): modulators 4.4 Voltage-dependent Na+ channel H Indoxacarb (1): blocker

GI– I (1):blocker Na+ Ca++

Ryanodine, diamides (2): activators Action J potential Glu Glu Type 1 pyrethroids by Public Health Information Access Project on 04/29/14. For personal use only. Control Type 2 pyrethroids Annu. Rev. Entomol. 2013.58:99-117. Downloaded from www.annualreviews.org action repetitive firing suppression of Vesicles Glu K Amitraz (1): mimics Glu potential action potential

Glu = glutamate

5.5 Ca2+–activated Ca2+ channel 66. Octopamine receptor coupled to second messengers

J ATP Mg2+ Ca2+ receptor OA OAOA Elevation of cyclic AMP Sarcoplasmic Ca2+ Muscle reticulum contraction OA OA K Neuroexcitation Transverse OA = octopamine

Cell membrane tubule Ca2+ Voltage channel sensor

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a AChE – oxonb nAChR –

nAChR-heteropentamer AChBP-homopentamer O O Cl N P Extracellular O O Side view Cl binding domain Cl Trans- membrane BP BP E334* domain Top view

Intracellular BP F295 BP BP Y337 H447* N131 Cl S203* F338 L141 L143 N

Y231 W174 C227 N G121 Y118 N G120 N N+ C226 H O– O Y119 Y224 W79

+ c GABAAR – d Na channel - Heteropentamer Homopentamer O α β 1 3 N Br L L V T A A T O L β3 L A β3 Br β3 T β3 T T L T L A NCA A NCA V L T L T A T L T α1 L γ S A β3 2 β3 O by Public Health Information Access Project on 04/29/14. For personal use only.

Annu. Rev. Entomol. 2013.58:99-117. Downloaded from www.annualreviews.org F1530 L L932* C933 9' CF3 L F1534 L L T929* L T T F1538 6' T Cl Cl F1537 N T N NH2 L925* M918*

2' A A S CF A A N O 3

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Selectivity and Resistance Inhibitor specificity between the AChE of insects and mammals and between susceptible and tol- erant strains of insects contributes to selective toxicity and resistance (19, 46). For example, the Agonist: substance selectivity of methyl for inhibiting house fly (Musca domestica) versus bovine erythrocyte that combines with a AChE is greatly increased on introducing a 3-methyl substituent to yield the of the fenitroth- receptor to initiate a response ion insecticide (58). Further, is a relatively poor inhibitor for honey bee (Apis mellifera) compared to house fly AChE (18). Mutations in some resistant strains confer low OP Antagonist: substance that combines with a and MC inhibitor sensitivity (101, 142) and potentially altered inhibitor specificity. As yet another + − receptor to oppose a example, ( / )- is effective on some insects resistant to other OPs, possibly because response (+)-profenofos acts directly and (−)-profenofos after biosulfoxidation (53, 146), resulting in two toxicants with different leaving groups on AChE phosphorylation. Unlike humans, some insects (e.g., aphids and mosquitoes) have a sensitive cysteine in the acyl pocket of AChE, providing an opportunity, not yet fully realized, for target site selective sulfhydryl reagent inhibitors (108, 117).

NICOTINIC RECEPTOR

Primary Target and Agonist Action ACh is the endogenous agonist and principal excitatory transmitter for rapid neurotransmission in the insect central nervous system and the mammalian central and peripheral nervous systems (90, 130, 131, 133). ACh released from the presynaptic membrane interacts with the agonist binding site at the extracellular domain of the ACh receptor (AChR) ion channel complex (92, 131, 151) (Figure 1). The receptor then undergoes a conformational change leading to channel opening, influx of extracellular Na+, and efflux of intracellular K+. Actions resembling those of nicotine are associated with the nicotinic receptor (nAChR) and those similar to muscarine are attributed to the muscarinic receptor (mAChR), i.e., ACh has both nicotinic and muscarinic actions. The search for muscarinic antagonists and agonists selective on insects relative to mammals has thus far been unsuccessful (61). Nicotine is more toxic to mammals than to most insects, and systematic structural changes have not greatly improved its potency or safety (150). The breakthrough discovery of the highly insecticidal but photolabile with a nitromethylene moiety and no cationic substituent (126) acting as an nAChR agonist (119) led ultimately to photostabilized compounds selective for insects relative to mammals, commercialized as imidacloprid (IMI) (70, 71) and six analogs designated neonicotinoids (67, 68, 133, 151). The guanidine or amidine unit is coplanar ←−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−− by Public Health Information Access Project on 04/29/14. For personal use only. Annu. Rev. Entomol. 2013.58:99-117. Downloaded from www.annualreviews.org Figure 2 Structural models of four primary targets of major neuroactive insecticides showing the proposed docking positions and some binding site residues designated as in the cited literature. (a) Chlorpyrifos oxon and AChE (mouse) showing the catalytic charge relay complex ∗ ( ) of S203, E334, and H447, with specificity conferred by interactions with additional sites and pockets (not shown) (98). (b) Imidacloprid and nAChR showing structural comparison of Torpedo nAChR and Aplysia AChBP (which has no transmembrane domain or cytoplasmic end) (135) and homology model of Myzus α2β1 nAChR based on AChBP, with proposed docking position of IMI and some binding site residues (137). The binding region includes 118, 174, 224, 226, 227, and 231 of the α subunit and 79, 131, 141, and 143 of the β subunit. (c) Fipronil and GABAAR (human) chloride ionophore shown as a native (α1)2(β3)2γ2 heteropentamer and a recombinant β3 homopentamer of enhanced insecticide or noncompetitive antagonist (NCA) sensitivity. The proposed β3 homopentamer channel gating amino acids are A2,T6,andL9 (31), with the 6T facing into the channel as shown in the upper cross-sections. (d ) Deltamethrin and voltage-gated Na+ channel of house fly (Musca domestica) in an open conformation with residues (∗) implicated in binding of M918, L925, T929, and L932 in the S4-S5 linkers, S5 helices, pore helices, and S6 helices (38, 39, 140). Abbreviations: AChBP, acetylcholine binding protein; AChE, acetylcholinesterase; BP, binding pocket; GABA, γ-aminobutyric acid; GABAAR, GABA type A receptor; nAChR, nicotinic acetylcholine receptor.

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and electronically conjugated with the nitro or cyano substituent, which facilitates partial negative (δ−) charge flow toward the tip (135, 136). They act as agonists at multiple nAChR subtypes, with differential selectivity between insects and mammals conferred by only minor structural changes nAChR: nicotinic acetylcholine receptor (132, 134). AChBP: acetylcholine binding protein Selectivity and Resistance GABA R: GABA A 3 receptor type A Electrophysiological experiments and radioligand binding studies with [ H]nicotine, [3H]epibatidine, and [3H]IMI (83, 84) establish remarkable similarities within insect nAChRs and selectivity between insect and mammalian receptors (133, 134, 135). Nicotine, epibati- dine, and desnitro-IMI (referred to collectively as nicotinoids) are protonated at physiological pH and undergo cation-π interaction with tryptophan at an nAChR subsite of the α4/β2 interface in mammals (135, 136). Precise binding site interactions were established by crystallography and photoaffinity labeling studies, with ACh binding protein (AChBP) from hemolymph of the mollusk Aplysia californica and the snail Lymnaea stagnalis as models for the insect and mammalian nAChR, respectively, followed by homology modeling (90, 128, 135, 137), illustrated for Myzus in Figure 2. Binding subsite specificity plays a major role in selective toxicity for protonated nicotinoids in mammals and electronegative tip neonicotinoids in insects. The sulfoximine insecticides (i.e., sulfoxaflor) have both structural similarities to and differences from neonicotinoids consistent with their unique behavior as nicotinic agonists (145). resistance associated with a modified nAChR binding site has been observed in planthoppers, aphids, and a few other species (3, 85, 86, 90, 92).

Channel Blockers and Allosteric Activators Cartap is a noncompetitive blocker of the nAChR (81) and spinosyns act at yet another unique site in the nAChR (73), and as such there is no cross-resistance between neonicotinoids, cartap, and spinosyns (95).

CHLORIDE CHANNEL

Primary GABA Receptor Target and Antagonist Action γ-Aminobutyric acid (GABA) is the principal inhibitory neurotransmitter of insects and mammals and serves as the agonist for opening the pentameric transmembrane Cl− channel (6, 14, 22, 102, by Public Health Information Access Project on 04/29/14. For personal use only. Annu. Rev. Entomol. 2013.58:99-117. Downloaded from www.annualreviews.org 105) (Figure 1). The human brain GABA type A receptor (GABAAR) consists of various combi- nations of sixteen α subunits, seven β subunits, and four γ subunits, typically two α,twoβ,and one γ subunit. Insect GABARs exist as several different subtypes and form a class distinct from

any vertebrate GABAARs (14). The resistance-to- (RDL) GABAR subtype is expressed in many insects, and RDL homomers closely mimic the pharmacology of in situ insect GABARs (47, 48). Picrotoxinin (PTX), the insecticide and fish poison of the fishberry plant (Anamirta cocculus), was used to probe Cl− channel functions, showing that GABA opens the channel and PTX blocks Cl− flux (102). Several billion pounds of PCCAs, including , , and the cyclodienes such as α-, were used in crop protection before their mode of action was established (11, 22). They were joined in 1993 by the 1-phenylpyrazole fipronil, which is of lower acute toxicity to mammals. Radioligand binding and electrophysiological studies led to the recognition that the action of PCCAs and fipronil was similar to or the same as that of PTX and a series of insecticidal trioxabicyclooctanes (22, 34, 41). [3H]dihydroPTX was developed first as a

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radioligand for the PTX site (91), followed by the improved bicyclophosphorothionate [35S]TBPS and two bicycloorthobenzoates, [3H]TBOB for studies with mammals and particularly [3H]EBOB for investigations with insects and mammals (33, 42, 79, 80, 107, 127). They all block GABA- − EBOB: ethynylbicy- induced signals and Cl flux (62) on binding to a noncompetitive antagonist (NCA) or insecticide cloorthobenzoate binding site at the receptor subunit interface (Figures 1, 2). Importantly, the human GABA R A Common target: recombinant β3-homopentamer resembles the insect receptor in sensitivity and specificity for compounds acting at NCAs (114, 115), prompting exhaustive site-directed mutagenesis (cysteine scanning), which pin- thesamebindingsite, pointed the critical active site as pore-facing residues A2,T6,andL9 and led to a binding site leading to similar model that fits several widely diverse classes of insecticidal NCAs (30, 31). phenotype and cross-resistance Defined mode of Selectivity and Resistance action: target site and biochemical and The major GABAergic insecticides have higher potency on the native house fly or Drosophila recep- electrophysical effects tor and the human β3-homopentamer than on the less sensitive mammalian native (α1)2(β3)2γ2 consistent with heteropentamer brain receptor. In house fly brain, lindane, α-endosulfan, and fipronil block the organismal response

channel by binding to the critical pore residues with IC50 values of 0.5–3 nM (115). The insect versus mammalian target site specificity (105) is much greater for lindane and fipronil than for α-endosulfan (115) and is uniquely large for a highly selective isoxazoline (104). Cross-resistance between some of the PCCAs was the first indication of a common target and defined mode of action (22). Almost all major insect pests have been selected for resistance to insecticides acting at the GABA-gated Cl− channel. In most cases, the resistance is due to a single point mutation; e.g., in cyclodiene-resistant RDL Drosophila, alanine is replaced by serine in the ion channel lining of the transmembrane 2 region, which confers insensitivity and establishes the binding site as being in the channel pore (35, 47, 48). Cross-resistance usually extends from the PCCAs to fipronil, which acts at the same or closely related sites.

Glutamate and GABA Receptor Activators − and analogs open the Cl channel on binding to the GABAAR at a site coupled to but distinct from the antagonist site (40, 63). However, the nematicidal and insecticidal activity of avermectin analogs is most likely due to binding to glutamate-gated Cl− channels (37, 149), and the normally low toxicity to mammals is due to the P glycoprotein drug pump acting as a part of the blood-brain barrier that restricts avermectin entry into the mammalian brain (76). by Public Health Information Access Project on 04/29/14. For personal use only. Annu. Rev. Entomol. 2013.58:99-117. Downloaded from www.annualreviews.org SODIUM CHANNEL

Primary Voltage-Dependent Target and Modulator Action Voltage-gated Na+ channels open and close in response to changes in membrane potential (5). In the mammalian brain there are one alpha and two β subunits. The pore-forming α subunit consists of a single polypeptide chain with four internally homologous domains (I–IV), each having six transmembrane helices (S1–S6). The domain II S4-S5 linker, S5 and S6 helices, and domain III S6 helix interface the lipid bilayer and are therefore accessible to lipid-soluble ligands (38, 39, 123) The insect Na+ channel proteins also consist of four homologous domains, each with six transmembrane segments (122). The pyrethrins (from flowers), synthetic pyrethroids, and DDT, despite different origins or structures, are considered together because of similar insecticidal mechanisms. They act on axonal neurotransmission at insect voltage-gated Na+ channel recognition sites to block Na+

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transport, enhance channel inactivation, prolong the course of the Na+ current during depolar- ization, and induce a residual slow-acting current (“tail current”) (6, 94, 122). action is considered to be of two types (2, 24, 50). Pyrethrins and synthetic pyrethroids lacking an α-cyano group, e.g., allethrin and , and the α-cyano compound induce excitation (Type I action) on binding to resting or inactivated channels, shifting the voltage dependence of activation to more negative potentials and causing a slowly activating Na+ current responsible for repetitive activity. Deltamethrin and related α-cyano pyrethroids that exhibit writhing and con- vulsive signs in mammals (Type II action) induce profound use-dependent modification of Na+ currents, implying preferential binding to activated Na+ channel states, and recruit increasing numbers of Na+ channels into permanent open states, which results in use-dependent depolariza- tion, inactivation of unmodified channels, and blockage of conduction. Based on these differences, consideration has been given to whether Types I and II pyrethroids should fall into the same category or be considered separately in risk analyses and residue tolerances (125).

Selectivity and Resistance The higher potency of DDT and pyrethroids on insects than on mammals is proposed to be due to several features. First, the insect target is intrinsically more sensitive than that of mammals. Second, the target has a negative temperature coefficient; i.e., the insecticides are more effective at the ambient temperature of insects (e.g., 15◦C–20◦C) than that of people (37.5◦C). Other selectivity factors are lipophilicity for selective pickup by the insect epicutide and enzymatic detoxification. The 1R- cis and trans isomers are almost equal in insecticidal activity but greatly different in mammalian toxicity, which may be due to both target site specificity and relative detoxification rates (24, 78, 124). Fish are extremely sensitive to most pyrethroid esters but apparently less so to the pyrethroid ethers and silafluofen, possibly due to their low aqueous (138). When DDT first lost its effectiveness for controlling house flies, it was surprising to observe cross-resistance to the pyrethrins (16) and all synthetic pyrethroids, which was confirmed by electrophysiological studies on nerve sensitivity, suggesting a common mode of action (5, 122). Resistance to pyrethroids and DDT in Musca is conferred by the L1014F (kdr) mutant alone or with M918T (superkdr) in domain II of the α subunit (15, 38, 140). The current binding site model rationalizes the structure-activity relationships of pyrethroids relative to DDT, Types I and II action, and the open- and closed-channel conformations (39). Some of the residues on the helices that form the putative binding contacts are not conserved between insects and vertebrates, consistent with their contribution to species specificity (103, 122). by Public Health Information Access Project on 04/29/14. For personal use only. Annu. Rev. Entomol. 2013.58:99-117. Downloaded from www.annualreviews.org

Other Sites Four other sites in insect Na+ channels are targets for insecticidal botanicals (veratridine and isobutylamides) or synthetic insecticides (oxadiazines and metaflumizone) without cross- resistance, in any case, to pyrethroids and DDT. Veratridine, an active ingredient of sabadilla (Schoenocaulon officinale), was an early probe to study the Na+ channel, but attempts to simplify the structure and improve the insecticidal activity and safety (139) have thus far been unsuccessful. The isobutylamides or lipid amides are based on natural product prototypes (e.g., pellitorine from Piper nigrum) and can be quite potent but have not achieved the required balance of potency, stability, and safety to make them outstanding insecticide candidates (45). The oxadiazine indox- acarb is an effective proinsecticide that is enzymatically hydrolyzed to the active agent (77, 147). Metaflumizone is a triketone with excellent systemic aphicidal activity (118).

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OTHER NERVE AND MUSCLE TARGETS

Octopamine Receptor Agonists OPIDN: Octopamine and the octopamine receptor in insects, analogous to dopamine and the dopaminergic organophosphate- system in mammals, are coupled to second messengers to elevate cyclic AMP, leading to neuroexci- induced delayed tation (57) (Figure 1). Formamidines acting as octopamine mimics block the receptor and provide neuropathy broad-spectrum insecticidal activity (59, 60). , bioactivated by N-demethylation, NTE: neuropathy was the most important insecticide of this type until its carcinogenic activity was discovered and target esterase it was replaced by amitraz (59, 64).

Homopteran Feeding Blockers Two insecticides act as selective homopteran feeding blockers. Pymetrozine is thought to bind at an ACh site of some nAChRs that is different from that of the neonicotinoids or (95). Flonicamid blocks the A-type K+ rectifying current at the presynaptic nerve terminal neurons, leading to prolonged depolarization and uncontrolled neurotransmitter release (55).

Ryanodine Receptor Modulators Ryanodine and 9,21-dehydroryanodine, the active ingredients of the botanical insecticide Ryania (no longer used), bind at the Ca2+-activated Ca2+ channel (109). [3H]ryanodine binding provides a convenient assay for the ryanodine receptors (RyRs) of mammalian nerve and muscle and insect muscle (82, 109, 144). The ryanodine site is similarly sensitive in insects and mammals and a large series of analogs was prepared, but high insecticidal potency combined with low mammalian toxicity was not adequately achieved (65, 66). However, the hydrolysis products ryanodol and 9,21-dehydroryanodol are selective for insects compared with mammals (144) and appear to act at a K+ channel site (141). The newest major class of insecticides is the diamides (flubendiamide, , and ), which selectively activate insect but not mammalian muscle on binding to a new and yet uncharacterized site on RyRs, leading to uncontrolled Ca2+ release (Figure 1), thereby conferring strong selectivity for insects (36, 44, 120).

Genetic Engineering Pest insect control with baculoviruses takes several days, allowing unacceptable crop damage (10). Baculoviruses expressing selected spider and scorpion toxins act much faster but have not yet found practical applications (72). by Public Health Information Access Project on 04/29/14. For personal use only. Annu. Rev. Entomol. 2013.58:99-117. Downloaded from www.annualreviews.org

SECONDARY EFFECTS

Serine Hydrolase Secondary Targets OP and MC pesticides inhibit not only AChE but also many other serine hydrolases in nerve tissues, as quantified by enzymatic activity assays on specific substrates (27, 29) or activity-based protein profiling (87). The major focus in safety evaluations is on AChE inhibition, but there is also concern about OP-induced delayed neuropathy (OPIDN) and interest in behavioral effects associated with disruption of the cannabinoid system (Figure 3). OPIDN involves axonopathy and peripheral paralysis and has affected approximately 50,000 people globally in the past eight decades from compounds other than insecticides; there were also incidents with the insecticides and leptofos. The target protein was designated neuropathy target esterase (NTE) (69)

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a OP-induced delayed neuropathy b Cannabinoid system

O Cannabinoid CB1 receptor Activation OH OC(CH2)nCH3 Delayed HO HO neurotoxicity OO OO axonopathy OP/MC P Choline P Choline 2-AG Anandamide Endo- –OO –OO inhibitors cannabinoids

NTE Arachidonic Prostaglandin acid precursor

RO RO Aging RO P P P R'O X R'O NTE –O NTE MAGL FAAH Hydrolases

c Parkinson’s disease d Alzheimer’s disease

Substantia nigra Transporter Cholinergic neurons Lewy body Basal forebrain MC (Aricept®) ubiquitin protea- Neurofibrillary Choline some system Rotenone tangles Choline α-synuclein HO acetyl AChE OP () Amyloid-bearing transferase HO CH2CH2NH2 plaques ACh Dopamine Other Permethrin neurotransmitters Other GABA Receptor neurotransmitters nAChR Nicotine glutamate

Figure 3 Secondary targets of neurotoxic insecticides. (a) OP-induced delayed neuropathy from inhibition and aging of lysophosphatidyl choline hydrolase (NTE). (b) OP- or MC-induced inhibition of MAGL and FAAH elevates endocannabinoid and lowers arachidonic acid levels. (c) Parkinson’s disease of the basal ganglia involves primarily dopaminergic neurons. (d ) Alzheimer’s disease of the basal forebrain involves disrupting cholinergic and other neurons, leading to tangles and plaques. Abbreviations: ACh, acetylcholine; AChE, acetylcholinesterase; 2-AG, 2-arachidonyl glycerol; FAAH, fatty acid amide hydrolase; GABA, γ-aminobutyric acid; MAGL, monoacyl glycerol lipase; MC, methylcarbamate; nAChR, nicotinic acetylcholine receptor; NTE, neuropathy target esterase; OP, organophosphate.

and identified as a lysophosphatidyl choline hydrolase (112, 113, 143). Diminished NTE activity in mice links OP exposure to hyperactivity (148) but not necessarily to attention deficit disorder in humans. NTE deficiency in mice is embryo lethal (93, 116, 148) but when localized to the hippocampus gives vacuolation of neuronal bodies and dendrites (1). by Public Health Information Access Project on 04/29/14. For personal use only. Annu. Rev. Entomol. 2013.58:99-117. Downloaded from www.annualreviews.org The cannabinoid system consists of the CB1 and CB2 G protein–coupled cannabinoid recep- tors in the central and peripheral nervous systems, respectively, and two endocannabinoid ligands, anandamide and 2-arachidonyl glycerol, biosynthesized as needed and degraded by the serine hy- drolases fatty acid amide hydrolase (FAAH) and monoacyl glycerol lipase (MAGL), respectively (Figure 3). This system is involved in appetite, pain, synaptic plasticity, mood, and the psychoac- tive effects of cannabis. Behavioral effects of some OPs suggested possible involvement of the cannabinoid system, and the serine hydrolases FAAH and MAGL were found to be sensitive to chlorpyrifos and profenofos in vivo in mice (25, 97). Potent OP irreversible inhibitors of MAGL and FAAH induce the full cannabinoid syndrome in mice associated with greatly elevated brain endocannabinoid levels and depleted free arachidonic acid (96). The current OP and MC insecti- MAGL: monoacyl cides as normally used do not appear to pose any cannabinoid-related toxicity problems. Selective glycerol lipase and reversible FAAH and MAGL inhibitors have been designed with the goal of pain relief (88).

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OP- and MC-induced teratogenesis by , , and other insecticides in hen eggs involves inhibition of N-formylkynurenine hydrolysis by arylformamidase (121) and depletion of NAD levels (110, 111). The importance of this secondary OP target in mice was later evaluated by identifying the catalytic site (106) and defining the effect of gene inactivation on the kynurenine pathway metabolites (43). Another OP-sensitive serine hydrolase is KIAA1363 with an acetyl monoalkylglycerol ether substrate that also serves as a chlorpyrifos oxon hydrolase (98, 99, 100).

Parkinson’s and Alzheimer’s Diseases The increasing incidence of Parkinson’s and Alzheimer’s diseases worldwide associated with aging is attributed to genetic and environmental factors, including exposure to neuroactive insecticides (49, 54) The Parkinson’s signs of tremor, rigidity, slowness of movement, and postural instability result from insufficient levels of dopamine in the substantia nigra of the midbrain leading to α- synuclein protein accumulation and formation of Lewy body inclusions. Some of the insecticides proposed to contribute to these changes and the incidence of Parkinson’s disease are the NADH oxidase inhibitors rotenone and deguelin (4, 17, 129), the PCCAs heptachlor and dieldrin (9, 12), and the pyrethroid permethrin (9, 52). Alzheimer’s disease is progressive and irreversible, lead- ing to memory loss, language problems, unpredictable behavioral changes, and development of amyloid plaques, neurofibrillary tangles, and loss of connections between neurons in brain tissue (51, 56). Choline acetyl transferase activity is low in Alzheimer’s, resulting in low ACh levels, and other neurotransmitters (GABA and glutamate) are also possibly involved. Mild to moder- ate Alzheimer’s signs can be relieved by nicotine and AChE inhibitors, including the important Alzheimer’s drug AriceptR and the candidate (but withdrawn) drug metrifonate (i.e., the super- seded OP insecticide trichlorfon) (13, 89), which delay ACh breakdown in the synaptic cleft, thereby enhancing cholinergic neurotransmission.

FUTURE PROSPECTS

Less Emphasis on Neurotoxicants Almost all insecticides used in the 1940s to 1980s were neurotoxicants, but this had dropped to 79% of the total world market value by 1990. Between 1997 and 2010, there was a marked shift from OPs and MCs to neonicotinoids accompanied by a distinct trend toward a variety of non-neuroactive insecticides (Figure 4). After seven decades of continuing discoveries, there are indications that we may be coming to the end of the golden age (26) of neuroactive insecticide by Public Health Information Access Project on 04/29/14. For personal use only.

Annu. Rev. Entomol. 2013.58:99-117. Downloaded from www.annualreviews.org research. Exploring nature and sifting through chemical libraries have repeatedly ended up with insecticides dependent on the same major targets (21). This is disappointing when new targets are needed to circumvent cross-resistance mechanisms. Although insect growth regulators, includ- ing insect hormone analogs or agonists and inhibitors of chitin synthesis, can be highly potent, effective, and selective, they act slowly in stopping crop damage, thereby limiting their role in . Inhibitors of Complex I and other sites in the respiratory chain, including selected thioureas, organotins, and uncouplers of oxidative phosphorylation, are limited by low selectivity between insects and mammals (23, 95).

Insecticide Activation and Detoxification Neurotoxicant selectivity and resistance are attributable not only to differences in the biochemical targets considered here but also to the relative rates of P450 oxidation, hydrolysis, and conjugation

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Cholinergic Na+ AChE nAChRchannel Other

40

30 OP MC neonic pyr

20 Worldwide market (%) 10

0 Years (1997 – 2010) Figure 4 Changes in use of insecticide classes between 1997 and 2010 showing decreases for (OPs), methylcarbamates (MCs), and pyrethroids (pyr) and increases for neonicotinoids (neonic) and other compounds. Abbreviations: AChE, acetylcholinesterase; nAChR, nicotinic acetylcholine receptor. Data shown for the years 1997, 2000, 2002, 2005, 2008, and 2010 from T.C. Sparks (personal communication) are similar to those from his coauthored paper (95).

Resistance leading to activation and detoxification in insects and mammals. Many neurotoxic insecticides are management: in fact proinsecticides, such as phosphorothionates and derivatized MCs, that undergo preferential the attempt to bioactivation in pests to the active metabolites. The use of proinsecticides makes it possible to minimize the buildup alter the physicochemical properties, distribution, and persistence for improved efficacy, resistance of resistance to insecticides and other management, and safety. pesticides through various means by Public Health Information Access Project on 04/29/14. For personal use only.

Annu. Rev. Entomol. 2013.58:99-117. Downloaded from www.annualreviews.org SUMMARY POINTS 1. The four major neuroactive insecticide targets (AChE, nAChR, GABAR, and Na+ chan- nels) have the following features in common: original recognition by a botanical insecti- cide or prototype; target site cross-resistance limiting their effectiveness; available models for insecticide binding site interactions; alternative subsites for insecticides less involved in cross-resistance phenomena. 2. Downsizing of research on insecticide chemistry and toxicology has limited the discovery of new chemotypes and biochemical targets, but this is partially compensated for by new in vitro screens and testing of massive chemical libraries. 3. Neuroactive insecticides have helped clarify the role of secondary targets in OPIDN, cannabinoid-like effects, PD, and AD.

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FUTURE ISSUES 1. There is a continuing need for new chemotypes of neuroactive insecticides as the next generation of “resistance busters.” 2. An increasing diversity of non-nerve targets will help to maintain control capabilities. 3. New centers of excellence in insecticide research will meet continuing and increasing demands for safe and effective chemicals.

DISCLOSURE STATEMENT The authors are not aware of any affiliations, memberships, funding, or financial holdings that might be perceived as affecting the objectivity of this review.

ACKNOWLEDGMENTS Financial support past and present from the U.S. National Institutes of Health and the William Muriece Hoskins Chair in Chemical and Molecular Entomology (University of California, Berkeley) ( J.E.C.) and from the U.S. National Science Foundation (CHE-0840505) (K.A.D.) is gratefully acknowledged. We thank undergraduate researchers Alex Laihsu, Elodie Tong-Lin, and particularly Sean Kodani for excellent assistance and advice.

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61. Honda H, Tomizawa M, Casida JE. 2007. Insect muscarinic acetylcholine receptor: pharmacological and toxicological profiles of antagonists and agonists. J. Agric. Food Chem. 55:2276–81 62. Huang J, Casida JE. 1996. Characterization of [3H]ethynylbicycloorthobenzoate ([3H]EBOB) binding and the action of insecticides on the γ-aminobutyric acid–gated chloride channel of cultured cerebellar granule neurons. J. Pharmacol. Exp. Ther. 279:1191–96 63. Huang J, Casida JE. 1997. Avermectin B1a binds to high- and low-affinity sites with dual effects on the γ-aminobutyric acid–gated chloride channel of cultured cerebellar granule neurons. J. Pharmacol. Exp. Ther. 281:261–66 64. IRAC: Summaries and Evaluations. 1983. Chlordimeform. IRAC Monogr. 30:61–72 65. Jefferies PR, Casida JE. 1994. chemistry and action. In Natural and Engineered Pest Management Agents, ed. PA Hedin, JJ Menn, RM Hollingworth, 55:130–44. Washington, DC: Am. Chem. Soc. 66. Jefferies PR, Yu P, Casida JE. 1997. Structural modifications increase the insecticidal activity of ryan- odine. Pestic. Sci. 51:33–38 67. Jeschke P, Nauen R. 2008. Neonicotinoids—from zero to hero in insecticide chemistry. Pest Manag. Sci. 64:1084–98 68. Jeschke P, Nauen R, Schindler M, Elbert A. 2011. Overview of the status and global strategy for neoni- cotinoids. J. Agric. Food Chem. 59:2897–908 69. Johnson MK, Glynn P. 2001. Neuropathy target esterase. In Handbook of Pesticide Toxicology,ed.RI Krieger, 2:953–65. San Diego: Academic 70. Kagabu S. 1997. Chloronicotinyl insecticides: discovery, application and future perspective. Rev. Toxicol. 1:75–129 71. Kagabu S. 2003. Molecular design of neonicotinoids: past, present and future. In Chemistry of Crop Protection: Progress and Prospects in Science and Regulation, ed. G Voss, G Ramos, pp. 193–212. New York: Wiley-VCH 72. Kamita SG, Kang K-D, Hammock BD, Inceoglu AB. 2005. Genetically modified baculoviruses for pest control. In Comprehensive Molecular Insect Science, ed. LI Gilbert, K Iatrov, SS Gill, pp. 271–322. Oxford, UK: Elsevier 73. Kirst HA. 2010. The spinosyn family of insecticides: realizing the potential of natural products research. J. Antibiot. 63:101–11 74. Koelle GB. 1963. Cholinesterases and Anticholinesterase Agents. Berlin: Springer. 1220 pp. 75. Kuhr RJ, Dorough HW. 1976. Carbamate Insecticides: Chemistry, Biochemistry and Toxicology. Cleveland: CRC Press. 301 pp. 76. Lankas GR, Cartwright ME, Umbenhauer D. 1997. P-glycoprotein deficiency in a subpopulation of CF-1 mice enhances avermectin-induced neurotoxicity. Toxicol. Appl. Pharmacol. 143:357–65 77. Lapied B, Grolleau F, Sattelle DB. 2001. Indoxacarb, an oxadiazine insecticide, blocks insect neuronal sodium channels. Br.J.Pharmacol.132:587–95 78. Lawrence LJ, Casida JE. 1982. Pyrethroid toxicology: mouse intracerebral structure–activity relation- ships. Pestic. Biochem. Physiol. 18:9–14 by Public Health Information Access Project on 04/29/14. For personal use only. Annu. Rev. Entomol. 2013.58:99-117. Downloaded from www.annualreviews.org 79. Lawrence LJ, Casida JE. 1984. Interactions of lindane, toxaphene and cyclodienes with brain-specific t-butylbicyclophosphorothionate receptor. Life Sci. 35:171–78 80. Lawrence LJ, Palmer CJ, Gee KW, Wang X, Yamamura HI, Casida JE. 1985. t- [3H]butylbicycloorthobenzoate: new radioligand probe for the γ-aminobutyric acid-regulated chloride ionophore. J. Neurochem. 45:798–804 81. Lee S-J, Tomizawa M, Casida JE. 2003. Nereistoxin and cartap neurotoxicity attributable to direct block of the insect nicotinic receptor/channel. J. Agric. Food Chem. 51:2646–52 82. Lehmberg E, Casida JE. 1994. Similarity of insect and mammalian ryanodine binding sites. Pestic. Biochem. Physiol. 48:145–52 3 83. Describes high 83. Liu M-Y, Casida JE. 1993. High affinity binding of [ H]imidacloprid in the insect acetylcholine affinity and selectivity of receptor. Pestic. Biochem. Physiol. 46:40–46 the insect nAChR 84. Liu M-Y, Lanford J, Casida JE. 1993. Relevance of [3H]imidacloprid binding site in house fly head binding site. acetylcholine receptor to insecticidal activity of 2-nitromethylene- and 2-nitroimino-imidazolidines. Pestic. Biochem. Physiol. 46:200–6

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85. Liu Z, Williamson MS, Lansdell SJ, Denholm I, Han Z, Millar NS. 2005. A nicotinic acetylcholine recep- tor mutation conferring target-site resistance to imidacloprid in Nilaparvata lugens (brown planthopper). Proc. Natl. Acad. Sci. USA 102:8420–25 86. Liu Z, Yao X, Zhang Y. 2008. Insect nicotinic acetylcholine receptors (nAChRs): important amino acid residues contributing to neonicotinoid insecticides selectivity and resistance. Afr. J. Biotechnol. 7:4935–39 87. Long JZ, Cravatt BF. 2011. The metabolic serine hydrolases and their functions in mammalian physiology and disease. Chem. Rev. 111:6022–63 88. Long JZ, Li W, Booker L, Burston JJ, Kinsey SG, et al. 2008. Selective blockade of 2- arachidonoylglycerol hydrolysis produces cannabinoid behavioral effects. Nat. Chem. Biol. 5:37–44 89. Lopez-Arrieta´ J, Schneider L. 2006. Metrifonate for Alzheimer’s disease. Cochrane Database Syst. Rev. 2:CD003155 90. Matsuda K, Kanaoka S, Akamatsu M, Sattelle DB. 2009. Diverse actions and target-site selectivity of neonicotinoids: structural insights. Mol. Pharmacol. 76:1–10 91. Matsumura F, Ghiasuddin SM. 1983. Evidence for similarities between cyclodiene type insecticides and picrotoxin in their toxic mechanisms. J. Environ. Sci. Health B 18:1–14 92. Millar NS, Denholm I. 2007. Nicotinic acetylcholine receptors: targets for commercially important insecticides. Invertebr. Neurosci. 7:53–66 93. Moser M, Li Y, Vaupel K, Kretzschmar D, Kluge R, et al. 2004. Placental failure and impaired vas- culogenesis result in embryonic lethality for neuropathy target esterase-deficient mice. Mol. Cell Biol. 24:1667–79 94. Narahashi T. 2000. Neuroreceptors and ion channels as the basis for drug action: past, present and future. The 2000 ASPET Otto Krayer Award Lecture. J. Pharmacol. Exp. Ther. 294:1–26 95. Nauen R, Elbert A, McCaffery A, Slater R, Sparks TC. 2012. IRAC: insecticide resistance, and 95. IRAC relates mode mode of action classification of insecticides. In Modern Crop Protection Compounds, ed. W Kramer, of action to resistance U Schirmer, P Jeschke, M Mitschel, pp. 935–55. Hoboken, NJ: Wiley. 2nd ed. management. 96. Nomura DK, Blankman JL, Simon GM, Fujioka K, Issa RS, et al. 2008. Activation of endocannabinoid system by organophosphorus nerve agents. Nat. Chem. Biol. 4:373–78 97. Nomura DK, Casida JE. 2011. Activity-based protein profiling of organophosphorus and thiocarbamate pesticides reveals multiple serine hydrolase targets in mouse brain. J. Agric. Food Chem. 59:2808–15 98. Nomura DK, Durkin KA, Chiang KP, Quistad GB, Cravatt BF, Casida JE. 2006. Serine hydrolase KIAA1363: toxicological and structural features with emphasis on organophosphate interactions. Chem. Res. Toxicol. 19:1142–50 99. Nomura DK, Fujioka K, Issa RS, Ward AM, Cravatt BF, Casida JE. 2008. Dual roles of brain ser- ine hydrolase KIAA1363 in either lipid metabolism and organophosphate detoxification. Toxicol. Appl. Pharmacol. 228:42–48 100. Nomura DK, Leung D, Chiang TP, Quistad GB, Cravatt BF, Casida JC. 2005. A brain detoxifying enzyme for organophosphorus nerve poisons. Proc. Natl. Acad. Sci. USA 102:6195–200 101. Oakeshott JG, Claudionos C, Campbell PM, Newcomb RD, Russell RJ. 2010. Biochemical genetics and genomics of insect esterases. In Insect Pharmacology, ed. LI Gilbert, SS Gill, pp. 229–301. London: by Public Health Information Access Project on 04/29/14. For personal use only. Annu. Rev. Entomol. 2013.58:99-117. Downloaded from www.annualreviews.org Academic 102. Olsen RW. 2002. GABA. In Neuropsychopharmacology—Fifth Generation of Progress, ed. KL David, D Charney, JT Coyle, C Nemeroff, pp. 159–88. Philadelphia: Lippincott Williams & Wilkins 103. O’Reilly AQ, Khambay BPS, Williamson MS, Field LM, Wallace BA, Davies TGE. 2006. Modeling insecticide-binding sites in the voltage-gated sodium channel. Biochem. J. 396:255–63 104. Ozoe Y, Asahi M, Ozoe F, Nakahira K, Mita T. 2010. The antiparasitic isoxazoline A1443 is a potent blocker of insect ligand-gated chloride channels. Biochem. Biophys. Res. Commun. 391:744–49 105. Ozoe Y, Takeda M, Matsuda K. 2009. γ-Aminobutyric acid receptors: a rationale for developing selective insect pest control chemicals. In Biorational Control of Arthropod Pests, ed. I Ishaaya, AR Horowitz, pp. 131– 162. Dordrecht, The Neth.: Springer 106. Pabarcus MK, Casida JE. 2005. Cloning, expression, and catalytic triad of recombinant arylformamidase. Protein Expr. Purif. 44:39–44 107. Palmer CJ, Casida JE. 1989. 1-(4-Ethynylphenyl)-2,6,7-trioxabicyclo[2.2.2]octanes: a new order of po- tency for insecticides acting at the GABA-gated chloride channel. J. Agric. Food Chem. 37:213–16

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108. Pang Y-P, Ekstrom¨ F, Polsinelli GA, Gao Y, Rana S, et al. 2009. Selective and irreversible inhibitors of mosquito for controlling malaria and other mosquito-borne diseases. PLoS ONE 4:e6851 109. Pessah IN, Waterhouse AL, Casida JE. 1985. The calcium ryanodine receptor complex of skeletal and cardiac muscle. Biochem. Biophys. Res. Commun. 128:449–56 110. Proctor NH, Casida JE. 1975. Organophosphorus and methylcarbamate insecticide teratogenesis: di- minished NAD in chicken embryos. Science 190:580–83 111. Proctor NH, Moscioni AD, Casida JE. 1976. Chicken embryo NAD levels lowered by teratogenic organophosphorus and methylcarbamate insecticides. Biochem. Pharmacol. 25:757–62 112. First in vivo 112. Quistad GB, Barlow C, Winrow CJ, Sparks SE, Casida JE. 2003. Evidence that mouse brain demonstration that neuropathy target esterase is a lysophospholipase. Proc. Natl. Acad. Sci. USA 100:7983–87 NTE is a 113. Quistad GB, Casida JE. 2004. Lysophospholipase inhibition by organophosphorus toxicants. Toxicol. lysophosphatidylcholine Appl. Pharmacol. 196:319–26 hydrolase. 114. Ratra GS, Casida JE. 2001. GABA receptor subunit composition relative to insecticide potency and selectivity. Toxicol. Lett. 122:215–22 115. Ratra GS, Kamita SG, Casida JE. 2001. Role of human GABAA receptor β3 subunit in insecticide toxicity. Toxicol. Appl. Pharmacol. 172:233–40 116. Rosenbluth J, Schiff R, Lam P, Nuriel T, Chao MV. 2009. Spongiform pathology in mouse CNS lacking ‘neuropathy target esterase’ and cellular prion protein. Neurobiol. Dis. 35:433–37 117. Rowland M, Tsigelny I, Wolfe M, Pezzementi L. 2008. Inactivation of an invertebrate acetyl- cholinesterase by sulfhydryl reagents: a reconsideration of the implications for insecticide design. Chem. Biol. Interact. 175:73–75 118. Salgado VL, Hayashi JH. 2007. Metaflumizone is a novel sodium channel blocker insecticide. Vet. Parasitol. 150:182–89 119. Sattelle DB, Buckingham SD, Wafford KA, Sherby SM, Bakry NM, et al. 1989. Actions of the insecticide 2-(nitromethylene)-tetrahydro-1,3-thiazine on insect and vertebrate nicotinic acetylcholine receptors. Proc. R. Soc. Lond. Sci. Ser. B 237:501–14 120. Sattelle DB, Cordova D, Cheek TR. 2008. Insect ryanodine receptors: molecular targets for novel pest control chemicals. Invertebr. Neurosci. 8:107–19 121. Seifert J, Casida JE. 1981. Mechanisms of teratogenesis induced by organophosphorus and methylcar- bamate insecticides. Prog. Pestic. Biochem. 1:219–46 122. Soderlund DM. 2010. Sodium channels. In Insect Pharmacology, ed. LI Gilbert, SS Gill, pp. 1–24. London: Academic 123. Soderlund DM, Bloomquist JR. 1989. Neurotoxic actions of pyrethroid insecticides. Annu. Rev. Entomol. 34:77–96 124. Soderlund DM, Casida JE. 1977. Effects of pyrethroid structure on rates of hydrolysis and oxidation by mouse liver microsomal enzymes. Pestic. Biochem. Physiol. 7:391–401 125. Soderlund DM, Clark JM, Sheets LP, Mullin LS, Piccirillo VJ, et al. 2002. Mechanisms of pyrethroid neurotoxicity: implications for cumulative risk assessment. Toxicology 171:3–59 by Public Health Information Access Project on 04/29/14. For personal use only. Annu. Rev. Entomol. 2013.58:99-117. Downloaded from www.annualreviews.org 126. Soloway SB, Henry AC, Kollmeyer WD, Padgett WM, Powell JE, et al. 1978. Nitromethylene hetero- cycles as insecticides. In Pesticide Venom Neurotoxicology, ed. DL Shankland, RM Hollingworth, T Smyth, pp. 153–58. New York: Plenum 127. Squires RF, Casida JE, Richardson M, Saederup E. 1983. [35S]t-butylbicyclophosphorothionate binds with high affinity to brain specific sites coupled to γ-aminobutyric acid-A and ion recognition sites. Mol. Pharmacol. 23:326–36 128. Talley TT, Harel M, Hibbs RH, Radic´ Z, Tomizawa M, et al. 2008. Atomic interactions of neonicotinoid agonists with AChBP: molecular recognition of the distinctive electronegative pharmacophore. Proc. Natl. Acad. Sci. USA 105:7606–11 129. Tanner CM, Kamel F, Ross GW, Hoppin JA, Goldman SM, et al. 2011. Rotenone, paraquat and Parkinson’s disease. Environ. Health Perspect. 119:866–72 130. Thany SH. 2010. Insect nicotinic acetylcholine receptors. Adv. Exp. Med. Biol. 683:118 131. Thany SH. 2010. Neonicotinoid insecticides: historical evolution and resistance mechanisms. Adv. Exp. Med. Biol. 683:75–83

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132. Tomizawa M, Casida JE. 1999. Minor structural changes in nicotinoid insecticides confer differential subtype selectivity for mammalian nicotinic acetylcholine receptors. Br. J. Pharmacol. 127:115–22 133. Tomizawa M, Casida JE. 2003. Selective toxicity of neonicotinoids attributable to specificity of insect and mammalian nicotinic receptors. Annu. Rev. Entomol. 48:339–64 134. Tomizawa M, Casida JE. 2005. Neonicotinoid insecticide toxicology: mechanisms of selective action. Annu. Rev. Pharmacol. Toxicol. 45:247–68 135. Tomizawa M, Casida JE. 2009. Molecular recognition of neonicotinoid insecticides: the deter- 135. Establishes minants of life or death. Acc. Chem. Res. 42:260–69 molecular aspects of the 136. Tomizawa M, Lee DL, Casida JE. 2000. Neonicotinoid insecticides: molecular features conferring neonicotinoid binding selectivity for insect versus mammalian nicotinic receptors. J. Agric. Food Chem. 48:6016–24 site. 137. Tomizawa M, Maltby D, Talley TT, Durkin KA, Medzihradszky KF, et al. 2008. Atypical nicotinic agonist bound conformations conferring subtype selectivity. Proc. Natl. Acad. Sci. USA 105:1728–32 138. Tomlin CDS. 2009. The Pesticide Manual: A World Compendium. Hampshire, UK: Br. Crop Prot. Counc. 1457 pp. 15th ed. 139. Ujvary´ I, Eya BK, Grendell RL, Toia RF, Casida JE. 1991. Insecticidal activity of various 3-acyl and other derivatives of veracevine relative to the Veratrum alkaloids, veratridine and cevidine. J. Agric. Food Chem. 39:1875–81 140. Usherwood PNR, Davies TGE, Mellor IR, O’Reilly AO, Peng F, et al. 2007. Mutations in DIIS5 and the DIIS4-S5 linker of Drosophila melanogaster sodium channel define binding domains for pyrethroids and DDT. FEBS Lett. 581:5485–92 141. Usherwood PNR, Vais H. 1995. Towards the development of ryanoid insecticides with low mammalian toxicity. Toxicol. Lett. 82 /83:247–54 142. Villatte F, Ziliani P, Marcel V, Menozzi P, Fournier D. 2000. A high number of mutations in insect acetylcholinesterase may provide insecticide resistance. Pestic. Biochem. Physiol. 67:95–102 143. Vose S, Holland NT, Eskenazi B, Casida JE. 2007. Lysophosphatidylcholine hydrolases of human ery- throcytes, lymphocytes and brain: sensitive targets of conserved specificity for organophosphorus delayed neurotoxicants. Toxicol. Appl. Pharmacol. 224:98–104 144. Waterhouse AL, Pessah IN, Francini AO, Casida JE. 1987. Structural aspects of ryanodine action and selectivity. J. Med. Chem. 30:710–16 145. Watson GB, Loso MR, Babcock JM, Hasler JM, Letherer TJ, et al. 2011. Novel nicotinic action of the sulfoximine insecticide sulfoxaflor. Insect Biochem. Mol. Biol. 41:432–39 146. Wing KD, Glickman AH, Casida JE. 1983. Oxidative bioactivation of S-alkyl phosphorothiolate pesti- cides: stereospecificity of profenofos insecticide activation. Science 219:63–65 147. Wing KD, Sacher M, Kagaya Y, Tsurubuchi Y, Mulderig L, et al. 2000. Bioactivation and mode of action of the oxadiazine indoxacarb in insects. Crop Prot. 19:537–45 148. Winrow CJ, Hemming ML, Allen DM, Quistad GB, Casida JE, Barlow C. 2003. Loss of neuropathy target esterase in mice links organophosphate exposure to hyperactivity. Nat. Genet. 33:477–85 149. Wolstenholme AJ, Rogers AT. 2005. Glutamate-gated chloride channels and the mode of action of the by Public Health Information Access Project on 04/29/14. For personal use only.

Annu. Rev. Entomol. 2013.58:99-117. Downloaded from www.annualreviews.org avermectin/milbemycin . Parasitology 131:S85–95 150. Yamamoto I. 1999. Nicotine to neonicotinoids: 1962 to 1997. In Nicotinoid Insecticides and the Nicotinic Acetylcholine Receptor, ed. I Yamamoto, JE Casida, pp. 3–27. Tokyo: Springer 151. Yamamoto I, Casida JE. 1999. Nicotinoid Insecticides and the Nicotinic Acetylcholine Receptor.Tokyo: Springer. 300 pp.

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Annual Review of Entomology Volume 58, 2013 Contents

Life as a Cataglyphologist—and Beyond R¨udiger Wehner ppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppp1 Ecological Mechanisms Underlying Arthropod Species Diversity in Grasslands Anthony Joern and Angela N. Laws ppppppppppppppppppppppppppppppppppppppppppppppppppppppppp19 Recurrent Evolution of Dependent Colony Foundation Across Eusocial Insects Adam L. Cronin, Mathieu Molet, Claudie Doums, Thibaud Monnin, and Christian Peeters ppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppp37 The Impact of Molecular Data on Our Understanding of Bee Phylogeny and Evolution Bryan N. Danforth, Sophie Cardinal, Christophe Praz, Eduardo A.B. Almeida, and Denis Michez pppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppp57 An Emerging Understanding of Mechanisms Governing Insect Herbivory Under Elevated CO2 Jorge A. Zavala, Paul D. Nabity, and Evan H. DeLucia pppppppppppppppppppppppppppppppppp79 Neuroactive Insecticides: Targets, Selectivity, Resistance, and Secondary Effects John E. Casida and Kathleen A. Durkin pppppppppppppppppppppppppppppppppppppppppppppppppppp99 Biological Pest Control in Mexico by Public Health Information Access Project on 04/29/14. For personal use only. Annu. Rev. Entomol. 2013.58:99-117. Downloaded from www.annualreviews.org Trevor Williams, Hugo C. Arredondo-Bernal, and Luis A. Rodr´ıguez-del-Bosque pppp119 Nutritional Ecology of Entomophagy in Humans and Other Primates David Raubenheimer and Jessica M. Rothman ppppppppppppppppppppppppppppppppppppppppppp141 Conservation and Variation in Hox Genes: How Insect Models Pioneered the Evo-Devo Field Alison Heffer and Leslie Pick ppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppp161 The Juvenile Hormone Signaling Pathway in Insect Development Marek Jindra, Subba R. Palli, and Lynn M. Riddiford pppppppppppppppppppppppppppppppppp181

viii EN58-Frontmatter ARI 5 December 2012 7:43

The Adult Dipteran Crop: A Unique and Overlooked Organ John G. Stoffolano Jr. and Aaron T. Haselton pppppppppppppppppppppppppppppppppppppppppppp205 Biology of Phlebotomine Sand Flies as Vectors of Disease Agents Paul D. Ready ppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppp227 Ecdysone Receptors: From the Ashburner Model to Structural Biology Ronald J. Hill, Isabelle M.L. Billas, Fran¸cois Bonneton, Lloyd D. Graham, and Michael C. Lawrence pppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppp251 Thelytokous Parthenogenesis in Eusocial Hymenoptera Christian Rabeling and Daniel J.C. Kronauer pppppppppppppppppppppppppppppppppppppppppppp273 Red Turpentine Beetle: Innocuous Native Becomes Invasive Tree Killer in China Jianghua Sun, Min Lu, Nancy E. Gillette, and Michael J. Wingfield pppppppppppppppppp293 Vision in Drosophila: Seeing the World Through a Model’s Eyes Angelique Paulk, S. Sean Millard, and Bruno van Swinderen pppppppppppppppppppppppppp313 Intrinsic Inter- and Intraspecific Competition in Parasitoid Wasps Jeffrey A. Harvey, Erik H. Poelman, and Toshiharu Tanaka ppppppppppppppppppppppppppp333 Biology and Management of Palm Dynastid Beetles: Recent Advances Geoffrey O. Bedford ppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppp353 Odorant Reception in Insects: Roles of Receptors, Binding Proteins, and Degrading Enzymes Walter S. Leal pppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppp373 Molecular Systematics and Insecticide Resistance in the Major African Malaria Vector Anopheles funestus Maureen Coetzee and Lizette L. Koekemoer pppppppppppppppppppppppppppppppppppppppppppppp393 Biology and Management of Asian Citrus Psyllid, Vector of the Huanglongbing Pathogens Elizabeth E. Grafton-Cardwell, Lukasz L. Stelinski, and Philip A. Stansly pppppppppppp413 by Public Health Information Access Project on 04/29/14. For personal use only. Annu. Rev. Entomol. 2013.58:99-117. Downloaded from www.annualreviews.org Host Preferences of Blood-Feeding Mosquitoes Willem Takken and Niels O. Verhulst pppppppppppppppppppppppppppppppppppppppppppppppppppp433 Biology of Invasive Termites: A Worldwide Review Theodore A. Evans, Brian T. Forschler, and J. Kenneth Grace pppppppppppppppppppppppppp455 Spider-Venom Peptides: Structure, Pharmacology, and Potential for Control of Insect Pests Glenn F. King and Margaret C. Hardy ppppppppppppppppppppppppppppppppppppppppppppppppppp475 Ecdysone Control of Developmental Transitions: Lessons from Drosophila Research Naoki Yamanaka, Kim F. Rewitz, and Michael B. O’Connor ppppppppppppppppppppppppppp497

Contents ix EN58-Frontmatter ARI 5 December 2012 7:43

Diamondback Moth Ecology and Management: Problems, Progress, and Prospects Michael J. Furlong, Denis J. Wright, and Lloyd M. Dosdall pppppppppppppppppppppppppppp517 Neural Mechanisms of Reward in Insects Clint J. Perry and Andrew B. Barron ppppppppppppppppppppppppppppppppppppppppppppppppppppp543 Potential of Insects as Food and Feed in Assuring Food Security Arnold van Huis pppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppp563 A History of Entomological Classification Michael S. Engel and Niels P. Kristensen ppppppppppppppppppppppppppppppppppppppppppppppppp585 Ants and the Fossil Record John S. LaPolla, Gennady M. Dlussky, and Vincent Perrichot pppppppppppppppppppppppppp609

Indexes

Cumulative Index of Contributing Authors, Volumes 49–58 ppppppppppppppppppppppppppp631 Cumulative Index of Article Titles, Volumes 49–58 ppppppppppppppppppppppppppppppppppppp636

Errata

An online log of corrections to Annual Review of Entomology articles may be found at http://ento.annualreviews.org/errata.shtml by Public Health Information Access Project on 04/29/14. For personal use only. Annu. Rev. Entomol. 2013.58:99-117. Downloaded from www.annualreviews.org

xContents Annual Reviews It’s about time. Your time. It’s time well spent.

New From Annual Reviews: Annual Review of Statistics and Its Application Volume 1 • Online January 2014 • http://statistics.annualreviews.org Editor: Stephen E. Fienberg, Carnegie Mellon University Associate Editors: Nancy Reid, University of Toronto Stephen M. Stigler, University of Chicago The Annual Review of Statistics and Its Application aims to inform statisticians and quantitative methodologists, as well as all scientists and users of statistics about major methodological advances and the computational tools that allow for their implementation. It will include developments in the field of statistics, including theoretical statistical underpinnings of new methodology, as well as developments in specific application domains such as biostatistics and bioinformatics, economics, machine learning, psychology, sociology, and aspects of the physical sciences. Complimentary online access to the first volume will be available until January 2015.

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