Chemosensory Transduction in Arthropods
Abstract and Keywords
Reception of chemicals via olfaction and gustation are prerequisites to find, distinguish, and recognize food and mates and to avoid dangers. Several receptor gene superfamilies are employed in arthropod chemosensation: inverse 7-transmembrane (7-TM) gustatory and olfactory receptors (GRs, ORs), 3-TM ionotropic glutamate-related receptors (IRs), receptor-guanylyl cyclases, transient receptor potential ion channels, and epithelial sodium channels. Some of these receptor gene families have ancient origins and expanded in several taxa, producing very large, variant gene families adapted to the respectively relevant odor ligands in species-specific environments. Biochemical and electrophysiological studies in situ as well as molecular genetics found evidence for G-protein-dependent signal transduction cascades for ORs, GRs, and IRs, suggesting that signal amplification is paramount for chemical senses. In contrast, heterologous expression studies argued for primarily ionotropic transduction as a prerequisite to interstimulus intervals in the range of microseconds.
Chemoreception is an ancient sense, absolutely required for survival. It is present in all uni- and multicellular organisms on earth. Different types of chemoreceptors with various transduction mechanisms were acquired during evolution. They convert information about quality, quantity, and kinetics of the chemical stimulus into electrically encoded information in the detecting receptor cell. Although there is general agreement about chemoreceptors and their signal transduction cascades in vertebrate chemoreception (Kato & Touhara, 2009; Persuy et al., 2015), there is an ongoing, lively debate about the function of arthropod chemoreceptors that is not yet resolved. Thus, chemoreception in land- or water-living arthropods is still not fully understood. The dispute focuses on the question of whether arthropod chemoreceptors are ionotropic receptors developed for extreme speed and/or G-protein-coupled receptors, employing signal amplification for maximizing sensitivity. Most data were collected in insects such as the fruitfly Drosophila melanogaster and the hawkmoth Manduca sexta as representatives for land-dwelling species (for reviews, Nakagawa & Vosshall, 2009; Stengl, 2010). Among water-living arthropods, crustaceans such as the lobster Panulirus argus are studied best (McClintock et al., 2006; Derby & Sorensen, 2008). The focus of this review lies on olfactory transduction of olfactory receptors (ORs) in insects and ionotropic glutamate-related (IRs) in crustaceans, while details of sensory organs morphology and perireceptor events were described elsewhere (Altner & Prillinger, 1980; Stocker, 1994; Leal, 2012; Pelosi et al., 2006, 2014).
Olfaction and Taste
Chemoreception is subdivided into olfaction and taste. The “sense of smell” or “odor reception” (p. 346) implies long-distance detection of volatile chemicals at low thresholds in air and water. Odor-detecting olfactory receptor neurons (ORNs) innervate mostly hair-like multipore sensilla on head appendages such as insect antennae or antennules of crustaceans (Altner & Prillinger, 1980; Schmidt & Gnatzy, 1984). Short-distance detection of less volatile chemicals mostly at higher thresholds is termed “sense of taste,” or “gustation.” The taste-detecting gustatory receptor neurons (GRNs) innervate stiff bristles with a terminal pore, primarily on legs and on appendages around the mouth. While gustation guides primarily reflexive and simple, consumatory behaviors such as feeding, olfaction mediates more complex behaviors such as search for distant odor sources, or courtship, and other social interactions (Derby & Sorensen, 2008). Nevertheless, these definitions of olfaction and taste are two extremes of a continuum, rather than describing two completely separate senses. They are especially difficult to separate in water-living organisms.
The Nature of Chemical Stimuli
Chemical molecules mostly occur in mixtures of seemingly infinite variability and abundance, over many orders of magnitude. In nature, animals are exposed to chemicals as solids, liquids, or gases from living and nonliving matter. The chemicals are released into the surrounding medium: the air or water, dependent on temperature. Land-living invertebrates either sample chemicals (gustatory stimuli) from solids or fluids directly via their gustatory (taste) organs. With their antennae/antennules they detect volatile chemicals (odorants, odors) in the air or odors dissolved in water from odor sources over long distances of up to several kilometers. Chemicals from the outside environment, from other species (allomones), from conspecifics, and from the sensing organism itself (pheromones) are detected and evaluated. Thus, chemicals are everywhere, omnipresent, and extremely complex. The chemical bouquets are highly variable, even if emitted from the same object over time, such as a ripening or rotting fruit. They present an overwhelming source of information about the environment, as well as about internal physiology and behavioral state of the chemically communicating senders and receivers. Furthermore, very rarely odors build up smooth concentration gradients with long odor durations of seconds or minutes while spreading only via undisturbed diffusion from an odor source. This may happen underground, in the soil, but it does not occur in air or water. In air and water, odor signals are intermittent with durations in the range of milliseconds, because of turbulences. Information of distance to an odor source, thus, is reflected in the frequency of odor filaments and in the slope of the concentration gradient within odor filaments (for review, Vickers, 2006).
Animals that sense an odor first need to decide whether they should ignore, avoid, or seek an odor source. Consequently, first of all, chemosensory organs need to filter relevant from irrelevant stimuli to avoid damage via overstimulation and to concentrate on what is most relevant. Very relevant are chemical stimuli for intra- and interspecific communication such as pheromones and allomones (Kaissling, 2014). Usually, they are costly for the producer and are sent off at very low concentrations. Thus, it is advantageous to develop highly sensitive, selective chemoreceptors for pheromones. In addition, there is the neverending necessity to detect new food sources. Therefore, the challenge is to detect and distinguish as many different odor/taste qualities as possible over a broad concentration range. Newly developing plants may advertise their flowers or fruits with specific, new mixtures of odors at changing, low or high concentrations. Consequently, arthropods need to detect, distinguish, learn, and memorize new chemically defined objects, despite changes in odor concentration or composition (Stopfer et al., 2003; Shen et al., 2013; Mizunami et al., 2015). Then, they need to decide whether to ignore, seek, or avoid the odor source (Li & Liberles, 2015). Once a new attractive scent is detected, the contest for finding the odor source starts. Only who is first at a new food source can exploit it best. Thus, new mechanisms for better odor source location are advantageous such as fast-adapting sensory neurons with membrane potential oscillations that tune into frequency detection of flickering odor filaments via resonance (Kato et al., 2014; Park et al., 2016).
To summarize, during evolution the ability to focus on relevant stimuli, to first detect, locate, and remember new food sources or mates, and to avoid poisons or predators will present advantages to survival and reproduction. Chemical senses need to develop specialist-receptor neurons with maximized sensitivity, as well as broadly tuned generalists. For odor source location via odor filament frequency tracking, it is advantageous to develop fast-following receptor neurons that can keep relevant stimulus memories if odor trails are lost. Finally, to detect the frequently fluctuating odor concentrations, receptor neurons need to span a (p. 347) wide range of stimulus concentrations with sliding thresholds adjustable to currently encountered odor concentrations.
Arthropod Chemoreceptors Belong to Different Families
In contrast to vision, which relies on two to four photoreceptors, chemical senses employ a very large number of receptors encoded by about 1% to >5% of the entire genome of an organism. The first family of chemoreceptors discovered was the family of vertebrate G-protein-coupled receptors (GPCRs) from the rhodopsin-related receptor superfamily (Buck & Axel, 1991). These vertebrate-type ORs are found from fish to men and are employed also in Caenorhabditis elegans (for review, Bargman, 2006). However, they appear not to function as chemoreceptors in arthropods. Because during evolution different gene families were recruited for chemosensation, chemoreceptors are highly diverse in different animal taxa and could not be detected in arthropods through homology screens. Thus, it took several years for arthropod chemoreceptors to be identified (Clyne et al., 1999; Vosshall et al., 1999; Benton et al., 2009). The predominant invertebrate chemoreceptors (Fig. 13.1) belong to two divergent families of receptors with seven transmembrane domains (7-TM receptors): gustatory receptors (GRs) and olfactory receptors (ORs) (Fig. 13.1A, B). Furthermore, two large families of 3-TM receptors related to ionotropic glutamate receptors (IRs) are employed as arthropod chemoreceptors (Fig. 13.1C). One family of IRs is expressed primarily in the antennae and the other in gustatory organs. In addition, ligand-gated mostly orphan receptor guanylyl cyclases (rGCs), 6-TM transient receptor potential (TRP) ion channels, and 2-TM degenerin/epithelial sodium channels (Deg/ENaCs = Ppks) are employed in arthropod chemosensation (Fig. 13.1D–F) (for reviews, Stengl, 2010; Rytz et al., 2013; Joseph & Carlson, 2015; Robertson, 2015; Saina et al., 2015).
Gustatory Receptors Are Ancient, Inverse 7-TM Receptors
GRs belong to a novel superfamily of 7TM receptors with inverse (intracellular N-terminus) membrane topology. It is assumed that relatives of the basal GR family are very ancient and originated early in or before animal evolution (Robertson, 2015; Saina et al., 2015). Gustatory receptor-like genes (Grls) were identified in genomes of all protostomia examined and in deuterostomia such as ambulacraria, but not in chordata. The Grls were found also in nonbilateria such as cnidaria and in placozoa, but not in porifera and ctenophora. They were described in genomes of different species such as placozoans, an anemon, coral, polychaete, leech, the nematode C. elegans, three molluscs (the limpet Lottia gigantean, the oyster Crassostrea gigas, and the sea hare Aplysia californica), a sea urchin, and an acorn. Because Grls were completely absent in some animal lineages such as chordates and were not found in the ctenophore Mnemiopsis leidyi, the demosponge Amphimedon queenslandica, and two choanoflagellates, it is unresolved whether this superfamily evolved before or during animal evolution (Giribert, 2015; Robertson, 2015). Apparently, two Grls of the cnidarian Nmatostella vectensis were located in the blastula and gastrula early during embryonic development, but not to adult putative chemosensory organs (Saina et al., 2015). Thus, arthropod GRs possibly originated from receptors that were involved in ligand-dependent pattern formation during embryogenesis before they were recruited to chemoreception and other senses.
Insect Gustatory Receptors Serve Different Behaviors
Vertebrates and insects distinguish the same five canonical tastes: sweet, bitter, amino acids (umami), salty, and sour (acidic). In addition they taste non-volatile pheromones (Ebbs & Amrein, 2007). With their sense of taste (gustation) they detect and locate nutrients, while avoiding the intake of bitter-tasting toxins. In insects gustatory stimuli are detected via primary sensory neurns with an axon (gustatory receptor neurons = GRNs) that expresses GRs in their dendritic membranes. The GRNs of head appendages project their dendrites into stiff long bristles with a terminal pore (sensilla chaetica) or into cone-shaped pegs, while sending their axons to the subesophageal ganglion (Stocker, 1994). One taste bristle is innervated by one mechanosensory neuron and two to four GRNs. Each GRN responds to one distinct taste modality, such as sweet, bitter/high-salt, low-salt, or water, as shown in extracellular recordings (Ishimoto & Tanimura, 2004). Fruitfly GRNs in different organs serve different functions (for reviews, Liman et al., 2014; Miyamoto & Amrein, 2014; Freeman & Dahanukar, 2015; Fuji et al., 2015; Joseph & Carlson, 2015; Harris et al., 2015). The taste sensilla which control feeding are located on the antenna, on the labellum, the proboscis, and the tarsi of the legs. Swallowing and processing of nutrients is mediated via GRNs in the labral sense organs of the (p. 348) pharynx and in the gut (LeDue et al., 2015). In addition, a fructose receptor (GR43a) is expressed in the central brain in the fruitfly, apparently sensing the internal nutrient status (Miyamoto et al., 2012, 2014; Miyamoto & Amrein, 2014). The GRNs at the wing margin and at distal tarsal segments of the legs appear to control grooming behavior, while other tarsal GRNs are involved in sexual behavior. Furthermore, GRNs at the ovipositor supervise egg laying. In addition, GRNs innervate the oviduct and the ejaculatory duct, apparently playing important roles in reproduction (Miyamoto & Amrein, 2014; Joseph & Carlson, 2015).
In the drosophila Brain the Processing of Aversive Versus Appetitive Tastes Is Segregated
Two parallel neuronal circuits in the brain control either avoidance or attraction. Thus, taste encoding follows the “labeled line model” according to valence. This means that apparently all aversive taste modalities activate the same specific neuronal circuit that mediates aversion. All appetitive tastes are encoded via a different, specific neuronal pathway that elicits attraction and consumatory behavior (Harris et al., 2015; Joseph & Carlson, 2015). In contrast, in Drosophila there is no confirmation of (p. 349) the “across-fiber pattern” model, which proposes that information from GRNs encoding different taste modalities is integrated in higher brain centers of insects, resulting in multimodal interneurons (Harris et al., 2015; Joseph & Carlson, 2015).
Different Chemoreceptor Families Are Involved in Insect Taste Perception
In insects the sense of taste is mediated via different receptor gene families such as GRs, IRs, members of the pickpocket (ppk) family of epithelial sodium channels, and TRPs (Cameron et al., 2010; Chen et al., 2010; Pikielny, 2012; Kang et al., 2010; Kim et al., 2010; Zhang et al., 2013; Zhang & Montell, 2013; Koh et al., 2014; for reviews, Miyamoto et al., 2014; Freeman & Dahanukar, 2015). The GR-dependent taste perception is best studied and most is known about gustation in D. melanogaster. A total of 60 extremely divergent GR genes encoding 68 different GRs via alternative splicing are expressed in GRNs of the fruitfly (Clyne et al., 2000; Dunipace et al., 2001; Scott et al., 2001; Robertson et al., 2003; for reviews, Liman et al., 2014; Joseph & Carlson, 2015). Insect GRs are highly diverse 7TM receptors with inverted membrane topology which can have as little as 8% of amino acid identity (Robertson et al., 2003; Xu et al., 2014; Zhang et al., 2011).
Gustatory Receptor Neurons Coexpress Many Gustatory Receptors of the Same Taste Quality
Expression studies showed that the sets of GRs expressed in sweet- or bitter-sensing neurons are mutually exclusive and that taste encoding occurs in segregated labeled lines in the brain (Harris et al., 2015). Furthermore, each GRN expresses many (up to 28) GRs of the same modality, such as bitter-sensing GRNs expressing different GRs for various bitter substances and sweet-sensing GRNs expressing different GRs for several sugars. These conclusions were partly derived from expression studies employing GAL4/UAS with Gr43aGAL4 in taste receptor neurons combined with different cell markers. The studies confirmed that the fructose-receptor GR43a is coexpressed with the sweet-sensing GR64f in a subset of sweet neurons in the tarsi and labellum, while it is not coexpressed with various bitter receptors tested (Miyamoto et al., 2012). Despite the fact that sweet- and bitter-GRs are not coexpressed in the same GRN, there is at least one peripheral and one central mechanism of lateral inhibition between sweet- and bitter-sensing GRNs (Jeong et al., 2013; French et al., 2015). This possibly serves for focusing on the most relevant stimulus.
Gustatory Receptors Can Only Be Exogenously Expressed in Gustatory Receptor Neurons but Not in Olfactory Receptor Neurons
Expression studies in the “empty neuron system” (sensory neuron with deleted chemoreceptors) of Drosophila as well as loss-of function experiments examined ligand/taste sensitivity of GRs. In the fruitfly the CO2-sensing ab1C GRN innervates the ab1 large basiconic sensillum together with three other sensory neurons. It expresses no ORs or IRs but expresses only CO2-sensitive GR21a and GR63a (Larsson et al., 2004; Jones et al., 2007; Kwon et al., 2007; Benton et al., 2009). Exogenous receptor expression in the ab1C GRN with the GAL4/UAS system employing GR21a or GR63a promoter sequences was employed to characterize GRs (Freeman et al., 2014). However, it was usually not possible to express GR genes in an “empty neuron system” that employed ORNs instead of GRNs (Jones et al., 2007; Kwon et al. 2007). Possibly, the GRs depended on an unknown GRN-specific chaperon or on an obligatory coreceptor to be stably expressed in dendritic cilia.
Eight Broadly Tuned Related Insect Gustatory Receptors of the Sweet Clade Encode Sweet Tastes
Sweet-sensing fruitfly GRNs express different combinations of eight GRs: GR5a, GR61a, and GR64a–f, which appear to belong to one clade of arthropod receptors (Dahanukar et al., 2007; Jiao et al., 2007; Slone et al., 2007; Kent & Robertson, 2009; Miyamoto et al., 2012). It was suggested that the GRs of the sweet clade form two subgroups of heteromeric receptor complexes which require coexpression of either GR5a or GR64a together with several other GRs (Dahanukar et al., 2007; Jiao et al., 2008; Freeman et al., 2014). While GR5a is required for the detection of trehalose and melezitose (Dahanukar et al., 2001, 2007; Ueno et al., 2001), GR64a is necessary for sucrose and maltose detection (Dahanukar et al., 2007; Jiao et al., 2007). In contrast, GR64f is required for all sugars tested except for fructose sensing, despite being coexpressed also with the fructose receptor GR43a. Possibly, GR64f is a very broadly tuned sugar receptor or an obligatory coreceptor or chaperon (Jiao et al., 2008). Thus, the broad tuning (p. 350) of the GRNs with overlapping response spectra appears to be due to expression of multiple ligand-dependent GRs in one GRN. Next to the broadly tuned heteromeric GR receptor complexes of the sugar clade there is also the narrowly tuned GR43a, which is specialized for fructose sensing (Miyamoto et al., 2012). GR43a is expressed in taste organs such as the tarsi, but it is also expressed in the central brain, apparently measuring hemolymph fructose levels (Miyamoto et al., 2012). Despite its narrow tuning, GR43a is necessary for detection of different sugars, most likely because these sugars are metabolized and change blood fructose levels (Mishra et al., 2013).
Most Insect Gustatory Receptors Encode Bitter Tastes
Usually, poisons taste bitter. Bitter tastes of alkaloids, phenols, and terpenoids are detected by bitter-sensing GRs such as GR66a, GR33a, GR93a that are partly coexpressed in the same GRN, the bitter/high-salt GRN (Moon et al., 2006, 2009; Lee et al., 2009; Weiss et al., 2011; Miyamoto et al., 2012; for review, Liman et al., 2014). The labellar taste sensilla of the fruitfly were grouped into four bitter-sensitive types (Weiss et al., 2011). Two types are narrowly tuned to specific bitter compounds, while the other two are broadly tuned with differing activity patterns. Reporter gene constructs showed that 33 of 38 GRs in labellar sensilla were expressed in bitter-sensitive GRNs (Weiss et al., 2011). As expected, the broadly tuned GRNs expressed more GRs than the narrowly tuned GRNs. Thus, most of the GRs of the fruitfly respond to bitter substances while only nine GRs detect sweet compounds. Furthermore, as deletion studies combined with heterologous expression showed, functional bitter receptors comprise heteromeric complexes of more than three subunits. Single GRNs express up to 28 ligand-binding GR subunits (Kwon et al., 2011; Weiss et al., 2011). Since GR32a, GR33a, GR66a were obligatory for most bitter chemicals tested, they appear to be broadly tuned and are coexpressed with varying numbers of additional GRs in all bitter-sensitive GRNs (Moon et al., 2009; Lee et al., 2010). In contrast, GR8a is narrowly tuned to detect L-canavanine, while GR93a responds specifically to caffeine (Lee et al., 2009, 2012). The vast number of different bitter-sensing GRs in Drosophila that are coexpressed in single GRNs appear to reflect the necessity to adapt to the increasing number of new poisons that plants develop in their attempt to fight off hungry insect larvae.
Tasting Transient Receptor Potential Channels and Gustatory Receptor That Encode Other Modalities
Coexpressed with GRs in bitter-sensitive GRNs are at least three different types of TRP ion channels: TRPA1 that is activated by aristolochic acid (Kim et al., 2010; Afroz et al., 2013), the TRPA-related channel Painless that is required for the detection of isothiocyanates (wasabi) (Al-Anzi et al., 2006), and the TRPL channel that senses camphor (Zhang et al., 2013; Liman et al., 2014). Still not identified is the low-salt and water-sensing GR (Dethier, 1976; Liu et al., 2003; Cameron et al., 2010). Furthermore, GRs of the Gr28b gene locus encode temperature sensors or a light sensor similar to a related GR which is required for phototaxis in C. elegans (Robertson et al., 2003; Thorne & Amrein, 2008; Xiang et al., 2010; Liu et al., 2010; Ni et al., 2013).
The Signal Transduction of Gustatory Receptors Is Not Resolved Yet
Signal transduction cascades of the highly diverse GRs with their little amino acid identity might not be the same for each GR. While heterologous expression studies suggested that the fructose receptor GR43a is a ligand-gated ion channel (Sato et al., 2011), several studies implied the involvement of different, partly parallel G-protein-dependent and/or cGMP-dependent signal transduction cascades for sweet and bitter tastes (Talluri et al., 1995; Koganezawa & Shimada, 2001; Ueno et al., 2006; Vermehren et al., 2006; Ueno & Kidokoro, 2008; Bredendiek et al., 2011; Vermehren-Schmaedick et al., 2011; Masek & Keene, 2013). Further experiments are necessary to resolve the issue of GR-dependent transduction cascades for each of these GRs in different species of arthropods.
To summarize, Drosophila GRs are inverse 7 TM receptors that form heteromeric complexes of unknown stoichiometry. Multiple GRs of the same modality and same valence are expressed per single sensory neuron. One bitter GRN can express not only many bitter-sensitive GRs but also other receptors such as TRPs that bind aversive compounds. Thus, tastes are encoded as valence-dependent labeled lines. Specialist receptors such as the fructose receptor GR43a co-occur next to broadly tuned generalists. Broad tuning appears to be accomplished via combination of different GR-subunits within GR-receptor complexes in a single sensory neuron. GR-dependent transduction mechanisms are not resolved yet and may differ (p. 351) between members of this highly diverse superfamily, depending on the respective needs for either amplification or reaction speed.
Insect Odorant Receptors Are Related to Gustatory Receptors and Are Primarily Expressed in Filiform Sensilla Trichoidea and Sensilla Basiconica of Insect Antennae
Insect olfactory receptors (ORs) belong to the same superfamily of 7TM chemoreceptors as GRs, unrelated to vertebrate ORs. The insect-type ORs appeared to diversify from one lineage of the highly diverse gustatory receptors (GRs) at the base of the hexapoda. The ORs expanded substantially to become the dominant class of odorant receptors in insects most likely around the time when flowering plant species and winged insects diversified (Carraher et al., 2015; Wicher, 2015). In contrast to gustatory stimuli, insects detect volatile odors with ORNs innervating multiporous hair-like sensilla trichoidea, basiconica, or sensilla coeloconica (pegs in a cuticular groove), and sensilla placodea on antennae and maxillary palps. Like other sensory neurons the ORNs are compartmentalized. Axons are isolated by a glial cell and extend from the antenna into the antennal lobes of the brain. Soma and dendrite are wrapped by an auxillary thecogen cell and are not exposed to the sensillum lymph. Only the cilium of the ORN extends into the sensillum lymph of the hair shaft, which is formed by thormogen and trichogen cells (Altner & Prillinger, 1980; Keil & Steinbrecht, 1984; Lee & Strausfeld, 1990; Stocker, 2001). Odorants pass through cuticular pores in the hair shafts or pore plate and reach the sensillum lymph, which contains high concentrations of odorant- or pheromone-binding proteins (OBPs or PBPs). The OBPs/PBPs accumulate odorants/pheromones partly with highly selective binding and prevent direct degradation via enzymes. Furthermore, the PBPs transport lipophilic pheromones through the aqueous sensillum lymph to their specific receptors in the cilia of the ORNs. Thus, insect OBPs/PBPs serve various functions and contribute to the specificity, sensitivity, and kinetics of odorant responses (Vogt & Riddiford, 1981, 1986; Steinbrecht et al., 1995; Ziegelberger, 1995; Pophof, 2004; Leal et al., 2005; Xu et al., 2005; Forstner et al., 2006; Pelosi et al., 2006; Grosse-Wilde et al., 2006, 2007; Laughlin et al., 2008; for reviews, Kaissling, 2009; Stengl, 2010; Pelosi et al., 2014). In vivo tip recordings of odor-sensitive hawkmoth sensilla showed that insect ORNs react to odor application with a specific action potential response that can be divided into three consecutive phases. The first phasic action potential response occurs during less than the first 100 ms of the odor response and is the only part of the response that encodes odor concentration changes. The second tonic odor response has a duration of several hundred milliseconds and does not necessarily encode odor pulse duration. The third delayed, long-lasting odor response occurs after about 1 second of the odor stimulus and can last over many seconds to minutes. The different kinetic components of the insect odor response hint that different processes/ion channels are involved in insect odor transduction (for review, Stengl, 2010; Stengl & Funk, 2013; Nolte et al., 2013).
Usually One Odorant Receptor Is Expressed per Olfactory Receptor Neuron Together With Orco, an Obligatory Coreceptor
Odorant receptors (ORs) of D. melanogaster were discovered first with a clever cloning strategy as well as with novel computer software predicting peptide sequences and characteristic protein domains from available genomic sequence databases (Clyne et al., 1999; Gao & Chess, 1999; Vosshall et al., 1999). So far, 60 OR genes were identified in the fruitfly genome, encoding for 62 different novel 7TM proteins with 17%–76% amino acid identity among its members (Clyne et al., 1999; Gao & Chess, 1999; Vosshall et al., 1999; for review, Joseph & Carlson, 2015). RNA in situ hybridizations combined with neuronal or nonneuronal markers confirmed that ORs are expressed in distinct subpopulations of ORNs of antennae and maxillary palps. In contrast to GRNs, ORNs express only one to three odorant-binding ORs together with a ubiquitous protein (A45 = OR83b = Orco) called olfactory receptor-coreceptor (Orco) (Vosshall & Hansson, 2011). Orco shares only 24% sequence identity with classical ORs and is larger than the ligand-binding ORs. Orco is expressed in all OR-expressing ORNs of sensilla trichoidea and basiconica. In addition, Orco shares up to 94% sequence identity with orthologs from different insect species in contrast to the highly divergent ORs (Krieger et al., 2002, 2003, 2005, 2009; Sakurai et al., 2004; Grosse-Wilde et al., 2007; Brigaud et al., 2009; Review; Nakagawa, & Vosshall, 2009). Orco and ORs appear to be specific for insect ORNs since so far no homologs were found in crustaceans (Peñalva-Arana et al., 2009; Corey et al., 2013).
(p. 352) Olfactory Receptors and Orco Are 7TM Receptors With Intracellular N-Termini, Forming Homo- and Heteromeric Complexes
Compared to vertebrate 7TM olfactory receptors insect ORs as well as Orco are inversely inserted into the membrane with an intracellular N-terminus (Benton et al., 2006; Wistrand et al., 2006; Lundin et al., 2007; Smart et al., 2008; Guo & Kim, 2010; Tsitoura et al., 2010). In heterologous expression ORs and Orco form homo- and heteromeric complexes of unknown stoichiometry (Neuhaus et al., 2005; German et al., 2013). In vivo they are assumed to associate with additional molecules such as sensory neuron membrane proteins (SNMPs) to form large complexes in signalosomes (Rogers et al., 1997, 2001a, 2001b; Benton et al., 2007; Forstner et al., 2008; Jin et al., 2008; Zhang et al., 2015; for reviews, Vogt et al., 2009; Stengl, 2010; German et al., 2013; Stengl & Funk, 2013).
Orco Does Not Bind Odors as Olfactory Receptors Do but Functions as an Obligatory Chaperon for Olfactory Receptors
While it is firmly established that odorant response specificity relies on the ligand-binding OR (Dobritsa et al., 2003; Elmore et al., 2003; Hallem et al., 2004), the role of Orco for odorant transduction is still under debate. Orco null mutants showed severely disrupted localization of ORs to olfactory cilia and rescue experiments restored stable expression of ORs in dendritic cilia. Thus, it is generally agreed upon that Orco acts as chaperon required for the location and maintenance of the ligand-binding ORs to the membranes of the dendritic cilia of ORNs (Larsson et al., 2004; Benton et al., 2006). Because expression of ORs in dendritic cilia of ORNs is a prerequisite to odor responses, Orco mutant flies and flies with RNAi-dependent knockdown of Orco did not respond to odorants (Larsson et al., 2004; Neuhaus et al., 2005). Therefore, Orco´s function as chaperon is required for OR-dependent odorant detection.
Orco Is a Pacemaker Channel Controlling Spontaneous Activity of Olfactory Receptor Neurons
Independent of coexpression with ORs in heterologous expression systems, Orco from different insect species forms a spontaneously opening Ca2+-permeable, nonspecific cation channel (Sato et al., 2008; Wicher et al., 2008; Jones et al., 2011; Sargsyan et al., 2011; Nolte et al., 2013). If Orco also forms a spontaneously opening cation channel in vivo, it would depolarize ORNs from the negative resting potential to spike threshold and would generate spontaneous action potential activity, as only pacemaker channels do. Indeed, ORNs from Orco-mutant flies had strongly decreased spontaneous activity (Larsson et al., 2004; Benton et al., 2007; Deng et al., 2011). Thus, Orco likely serves as a pacemaker channel in fruitfly ORNs that promotes spontaneous membrane potential depolarizations, controlling spontaneous action potential activity. As pacemaker channel that controls membrane potential oscillations and spike threshold, Orco will affect threshold and kinetics of odorant responses. Furthermore, endogenous subthreshold membrane potential oscillations are hallmarks of temporal encoding, turning ORNs into temporal filters. In addition, ultradian membrane potential oscillations allow for recruitment of neuronal ensembles via different means of synchronization (Nadasdy, 2010). Thus, it was proposed that Orco´s function as pacemaker channel is a prerequisite to improve resolution of odor pulses via resonance (Stengl, 2010).
Olfactory Receptor–Orco-Dependent Odor Transduction Cascades in drosophila Are Under Debate
Based upon their unexpected inverse membrane topology, it was questioned whether insect ORs couple to trimeric G-proteins as vertebrate 7TM ORs do (for review, Nakagawa & Vosshall, 2009). Sato et al. (2008) suggested that in the fruitfly, moth, and mosquito OR-Orco form odorant-dependent receptor-ion channel complexes that underlie ionotropic odor transduction, without any contribution of metabotropic cascades. In contrast, Wicher et al. (2008) proposed a low-sensitive ionotropic and a high-sensitive metabotropic odor transduction cascade for D. melanogaster. Both hypotheses were based upon heterologous coexpression of different fruitfly ORs + Orco. Strong (100 µM) odor stimulation lasting for seconds only evoked nonselective cation currents under these conditions (Sato et al., 2008). The authors concluded that the odor-binding OR together with Orco form an odor-dependent receptor-ion channel complex where both subunits are necessary for ion channel pore formation. In other studies heterologous expression of ORs from fruitflies and moths in the absence of Orco already evoked odor-dependent responses (Wetzel et al., 2001; Sakurai et al., 2004; Grosse-Wilde et al., 2006; Smart et al., (p. 353) 2008; Deng et al., 2011). Because odor-ligand-gated ORs were not able to form ion channels if expressed alone, it was concluded that they couple to trimeric G-proteins present in the heterologous cells without any need to heteromerize with Orco. Accordingly, odor-dependent rises in cAMP levels were measured with heterologous expression of fruitfly OR22a + Orco (Wicher et al., 2008). In addition, antennal homogenates from cockroaches and moths rapidly and transiently increased IP3 but not cAMP levels after species-specific pheromone stimulation, while cGMP levels increased more slowly and more sustained (Boekhoff et al., 1990, 1993; Breer et al., 1990; Ziegelberger et al., 1990). These biochemical experiments argue for the activation of different metabotropic signal transduction cascades in different insects in addition to activation of a G-protein-independent activation of receptor-guanylyl cyclases in insect odor and pheromone transduction (Stengl et al., 2001; for review, Stengl, 2010).
In contrast, Sato et al. (2008) did not confirm the activation of a G-protein-dependent adenylyl cyclase or phospholipase C after odor application. Because no odor-dependent cAMP rises were measured after odor stimulation in heterologous expression of Drosophila OR47a + Orco, and inclusion of cAMP, cGMP, or IP3 in the patch pipette in HeLa cells that coexpressed fruitfly OR47a + Orco did not elicit currents, it was concluded that ORs and Orco are not directly second messenger dependent and that odor-dependent currents in these cells were not elicited via metabotropic cascades (Sato et al., 2008). In contrast, solely ionotropic odor transduction was claimed to occur in fruitflies and moths because neither the phospholipase C antagonist U73122 nor nonhydrolysable GDPβS strongly affected odor responses of heterologously expressed OR47a + Orco or OR43b + Orco, or pheromone responses of the silkmoth’s bombykal receptor OR1 and Orco (Sato et al., 2008; Smart et al., 2008). Furthermore, pharmacological interference with metabotropic cascades in single sensillum recordings only moderately changed responses to general odorants in D. melanogaster (Yao & Carlson, 2010). To reconcile these negative findings with published contradictory data, it remains to be examined whether changes in the experimental conditions, such as different extracellular Ca2+ concentrations, Zeitgebertime, or different doses of antagonists/second messengers might have revealed other results. Furthermore, parallel G-protein-dependent cascades in insect ORNs together with the presence of odor-dependent but G-protein-independent receptor guanylyl cyclase, as described in the hawkmoth (Stengl, 2010), might be responsible for the moderate effects of pharmacological interference with metabotropic cascades.
In support of an ionotropic cascade were outside-out patch clamp recordings from heterologously expressed Drosophila OR 47a + Orco and Anopheles GPROR2 + Orco eliciting spontaneous ion channel opening that increased odor dose-dependently without the need to coapply ATP or GTP. Thus, the authors followed that Orco from different species is a spontaneously opening ion channel that increases its open time probability after application of odors via direct activation of an OR-Orco ion channel complex (Sato et al., 2008). Furthermore, Wicher et al. (2008) measured in HEK-cells with whole-cell patch clamp recordings an odor-dependent inward current dependent on heterologous expression of fruitfly OR22a + Orco that neither depended on coapplication of GTP nor ATP. While these experiments argued for an insensitive ionotropic transduction based upon OR-Orco receptor ion channel complexes that can be activated at very high odor concentrations, it could not be disproven that excised patches contained signalosomes with metabotropic cascades. In addition, it might not have been necessary to add ATP and GTP because HEK293 cells maintained enough endogenous ATP and GTP during the first second of the whole-cell recordings.
In the fruitfly D. melanogaster, experimental evidence from different groups suggested that metabotropic odor transduction cascades are employed (Talluri et al., 1995; Kain et al., 2008; Wicher et al., 2008; Chatterjee et al., 2009; Boto et al., 2010; Bredendiek et al., 2011; French et al., 2011). At physiologically low odor doses the ligand binding OR couples to Gαs activating adenylyl cyclase. The rises in intracellular cAMP then open the cation channel Orco, if Orco is previously phosphorylated via a protein kinase C via unknown mechanisms (Wicher et al., 2008; Sargsyan et al., 2011). It remains to be studied further whether an OR-Orco-based metabotropic mechanism is activated in vivo that controls phosphorylation of Orco Ca2+-dependently or whether in vivo odor-dependent openings of OR-Orco receptor ion channel complexes are the first transduction events to occur underlying the phasic odor response.
(p. 354) Olfactory Receptor–Dependent Metabotropic Pheromone Transduction in Moths
Besides the genetic model system, Drosophila insect olfaction has been studied best in the large hawkmoth Manduca sexta (Stengl, 2010; Martin et al., 2011). Female hawkmoths release a species-specific sex pheromone blend in pulsatile fashion that the males track to locate their mates. Next to biochemical experiments that described sharp, transient pheromone-dependent rises in antennal IP3 concentrations underlying phasic pheromone responses (Boekhoff et al., 1993), electrophysiological recordings found evidence for a phospholipase Cβ-dependent odor transduction cascade (Stengl, 2010). In patch clamp experiments of primary cell cultures of antennal ORNs from M. sexta, application of sex-pheromone at physiologically low concentrations elicited a sequence of three inward currents (Stengl & Hildebrand, 1990; Stengl et al., 1992; Stengl, 1993, 1994). Pharmacological experiments and ion exchange experiments suggested that the first very transient inward current was caused by IP3-dependent opening of a plasma membrane Ca2+ channel. The resulting Ca2+ influx then opened a Ca2+-dependent nonspecific cation channel. Further Ca2+ concentration rises in the presence of pheromone then activated protein kinase C–dependent, nonspecific, sustained-opening cation channels with low Ca2+ permeability (Stengl, 1993, 1994). Finally, very strong or long pheromone stimuli activated a G-protein-independent receptor-guanylyl cyclase causing rises in intracellular cGMP concentrations (Ziegelberger et al., 1990; Stengl et al., 2001). To examine whether the first, transient inward current was preceded by an ionotropic mechanism based upon OR-Orco, in vivo experiments were performed with tip recordings of single pheromone-sensitive sensilla trichodea (Nolte et al., 2013). Pharmacological interference with Orco revealed that Orco was not active during the first 1,000 msec of the phasic pheromone response. Instead, Orco opened during a later time window of the late, long-lasting pheromone response, when pheromone elevated cGMP concentrations (Ziegelberger et al., 1990; Stengl et al., 2001; Nolte et al., 2013). Thus, it is intriguing to hypothesize that Orco is activated voltage- and second messenger-dependently during the late, long-lasting pheromone response to maintain an odor memory over seconds to minutes after the pheromone stimulus. If so, Orco’s control of the membrane potential and membrane potential oscillations should greatly affect pheromone detection thresholds and pheromone pulse tracking via resonance (Nadasdy, 2010; Stengl, 2010; Stengl & Funk, 2013). Further in vivo experiments in different species need to examine whether, when, and how Orco is opened during pheromone and odor stimulation.
To summarize, there is no general agreement of OR-dependent signal transduction cascades in the scientific community, not even for the fruitfly D. melanogaster. However, evidence based upon morphological, biochemical, and physiological data is accumulating for metabotropic signal transduction cascades employed by the odor-binding OR despite its inverse membrane topology. There is general consensus that the non-ligand-binding coreceptor Orco functions as a “chaperon” localizing and maintaining odor-binding ORs to the dendritic cilia’s membrane. In addition, it is agreed upon that Orco functions as a spontaneously opening pacemaker ion channel that controls the spontaneous activity of ORNs. Orco appears to turn ORNs into ultradian oscillators that are optimized to track odor pulses for odor source location. However, it is not agreed upon whether Orco together with ORs not only in vitro but also in vivo functions as an odor-dependent receptor ion channel complex. At least in the hawkmoth there is no evidence for OR-Orco-based ionotropic pheromone transduction. Further in vivo experiments need to examine the function of Orco in odor/pheromone transduction in different species to reconcile open disputes.
Ancient Chemoreceptors Related to Ionotropic Glutamate Receptors Termed Ionotropic Receptors
In the fruitfly D. melanogaster a novel family of chemosensory receptors was identified with a bioinformatic screen for antennal-expressed genes combined with molecular genetics (Benton et al., 2007, 2009). BLAST searches identified 61fruitfly genes that form a divergent subfamily of ionotropic glutamate receptors (iGluRs) termed ionotropic receptors (IRs). The IRs are ancient ancestral protostome chemoreceptors that are absent in deuterostomia but are expressed in olfactory organs of all protostomia examined (Croset et al., 2010). Protostomia comprise the clades ecdysozoa with the taxa nematoda (e.g., C. elegans) and arthropoda (e.g., insects, crustaceans), as well as the clade spiralia with the taxon mollusca (e.g., snails). In contrast to ORs, the IRs are expressed in both gustatory and olfactory organs, found in (p. 355) nematodes, molluscs, crustaceans, and insects. Thus, two subfamilies are distinguished that are either expressed in the antenna, the olfactory organ, or in gustatory organs such as mouth appendices and tarsi.
A comparison of the structural domains of iGluRs and IRs revealed interesting differences (Croset et al., 2010; Abuin et al., 2011). The iGluRs occur in eucaryotes and prokaryotes, in plants and animals alike. Apparently, they originated when distinct genes were combined which encoded different characteristic protein domains. One such iGluR-domain is the extracellular amino-terminal domain that is involved in assembly of iGluR subunits into four-unit heteromers. Another is the ligand-binding domain, which forms a S1-S2- “Venus flytrap” clasping glutamate (Fig. 13.1C). Then, there is the ion channel pore between S1 and S2, consisting of two transmembrane regions and a pore loop. Finally, the channels have a third transmembrane domain and a cytosolic carboxy-terminal region. The ligand-binding domains of odor-sensitive IRs are highly divergent from each other and from ligand-binding domains of iGluRs, as expected for the recognition of different odor ligands (Benton et al., 2009). Furthermore, the pore domain of odor-binding IRs expresses little conservation compared to the pore domain of iGluRs (Croset et al., 2010; Abuin et al., 2011), suggesting loss of function as ion channels. In addition, loss of the amino-terminal and reduction of the C-terminal domains in the divergent odor-binding IRs suggested an increasing specialization of these subunits for odor recognition. Apparently, all odor-binding IRs need either of two very conserved, broadly expressed coreceptors, the IR25a or IR8a, for localization and maintenance into the membranes of dendritic cilia of ORNs, comparably to ORs’ need of Orco.
Orthologs of IR25a are expressed in chemosensory neurons of nematods, molluscs, and insects and thus appear to be the ancestral IR, which diverged first from the dGluRs (Croset et al., 2010). With tissue-specific real time-PCR, RNA in situ hybridizations, and immunocytochemistry, it was confirmed that IRs are expressed in chemosensory organs of the fruitfly (Benton et al., 2009). Double RNA in situ hybridization with probes for Orco and probes for different IR genes determined that IRs and ORs are usually not coexpressed in the same fruitfly sensilla. While ORs are expressed in long trichoid sensilla and shorter sensilla basiconica, which are not present in paleoptera such as the mayfly Rhithrogena semicolorata, IRs are expressed in sensilla coeloconica, which occur in both paleoptera and neoptera (Benton et al., 2009). Exception to the rule is one coeloconic sensillum, the funicular ac3 sensillum of D. melanogaster. It is innervated by two sensory neurons. One expresses IR75a, IR75b, IR75c, and IR8a and responds to propionic acid. The other ORN responds to γ-hexalactone and expresses OR35a and OR83b, in addition to IR76b. While the γ-hexalactone-sensitivity neither depends on IR8a, nor on IR25a, it is not known whether and how OR35a/OR83b and IR76b together contribute to odor sensitivity in this ORN (Abuin et al., 2011).
The IR25a is conserved and widely expressed in the ORNs of sensilla coeloconica of insects, similar to the more recently evolved insect-specific dublicate IR8a, as confirmed with RNA in situ hybridizations. Both IR25a and IR8a are most closely related to iGluRs, as compared to other IRs (Benton et al., 2009; Abuin et al., 2011). They appear to bind no odor ligands but appear to function as coreceptors which locate and maintain odor-binding IRs to the ciliary membranes. In contrast to other IRs that have many substitutions in the pore domain, IR25a and IR8a have pore domains most similar to the iGluR. Thus, IR25a and IR8a might still function as ion channels, possibly similar to Orco in the OR-expressing ORNs. The “ligand-binding” domain and the large intracellular C-terminus of IR8a and IR25a are both necessary for IR complex formation and localization to the cilia. Also the coreceptors depend on the odor-binding IRs for correct cilia targeting. With fluorescent protein-tagged IRs and internal reflection fluorescence microscopy in Xenopus oocyte expression studies, the stoichiometry of IR complex formation was examined (Abuin et al., 2011). It was suggested that two IR8a or two IR25a coassemble with two ligand-binding IRs in a proposed 2 + 2 stoichiometry for correct localization to the dendritic cilia (Abuin et al., 2011). It remains to be examined whether this stoichiometry is employed in situ in the dendritic cilia of the sensory neurons.
In situ hybridizations mapped the expression patterns of D. melanogaster IRs to ORNs of sensilla coeloconica of the funiculus, the sacculus, and the arista of the antenna. The patterns were conserved between both sexes and across individual flies (Benton et al., 2009). Each sensillum coeloconicum is innervated by two to three ORNs. In contrast to OR-expressing ORNs, there are up to four different ligand-binding IRs coexpressed together with either IR8a or IR25a pro ORN (Benton et al., 2009).
(p. 356) Signal Transduction of Ionotropic Glutamate-Related Receptors Is G-Protein Coupled in Crustaceans
IR-dependent signal transduction has been studied most intensely in crustaceans (for review, Ache & Young, 2005). In crustaceans, molecular genetics neither identified mammalian-type ORs nor insect-type inverse ORs. In contrast, IRs were the only type of olfactory receptor detected in lobster olfactory tissue so far (McClintock et al., 2006; Stephanyan et al., 2006; Corey et al., 2013). Electrophysiological, biochemical, and immunocytochemical experiments provided evidence for two different metabotropic signal transduction cascades that either activated or inhibited lobster ORNS (for review, Ache & Young, 2005). Different G-proteins, phospholipase Cβ, and adenylyl cyclase, as well as IRs, were located in lobster ORNs with genomic techniques and with immunocytochemistry (Fadool & Ache, 1992; McClintock et al., 2006; Corey et al., 2010, 2013). Also, biochemical studies showed that different odors either increased phospholipid- and/or cAMP-concentrations in lobster olfactory organs matching their physiological profile as predominantly excitatory or inhibitory odors (Boekhoff et al., 1994; Corey et al., 2010). Furthermore, electrophysiological studies characterized phospholipid- and cAMP-gated ion channels, which resembled odor-dependent ion channels in lobster aesthetasc sensilla. Thus, it was concluded that excitatory odors activate Gαq-dependently phospholipase Cβ. The resulting rises in IP3 then activate ion channels in the plasma membrane of ORNs that cause depolarizations (Fadool & Ache, 1992; Boekhoff et al., 1994; Fadool et al., 1995; Bobkov et al., 2010). In contrast, it was followed that inhibitory odors activate Gαs that activates adenylyl cyclase. The rises in cAMP concentrations activate potassium or chloride channels that hyperpolarize lobster ORNs (Doolin et al., 2001; Doolin & Ache, 2005). As further evidence for IR-dependent metabotropic signal transduction cascades, phosphoinositide 3-kinase is activated in lobster ORNs odor and G-protein dependently, and different phosphoinositides rise after odor stimulation in the antenna, which are able to modulate odor-dependent ion channel activity (Bobkov et al., 2010; Corey et al., 2010). Thus, there is strong evidence based on different experimental techniques in situ that IRs are G-protein coupled in crustaceans.
Functional studies of IR-dependent signal transduction in insects are very rare and are mostly based upon genetic studies, on considerations of structural relationship of IRs and iGluRs, and on heterologous expression (Benton et al., 2009; Abuin et al., 2011). In heterologous expression studies employing Xenopus oocytes fruitfly IR84a + IR8a or IR75a + IR8a were coexpressed to test for either phenylacetaldehyde or propionic acid-dependent currents. Very high odor concentrations of 1 mM were necessary to obtain responses. Respective odor stimulation evoked nonspecific cation currents which did not significantly differ between both receptor combinations (Abuin et al., 2011). No experiments were performed to exclude coupling of expressed IRs to G-proteins present in the heterologous expression system. In addition, the pharmacology of these odor-dependent currents did not resemble properties of iGluRs, and no convincing blocker of odor-dependent currents was found. Furthermore, considerable sequence divergence between predicted pore regions of IRs and iGluRs pore domains suggests that there was no need to conserve an ion channel pore for IR function. Single amino acid mutation of a residue in the predicted pore region in IR84a which controls Ca2+ permeability in iGluRs reduced inward currents in Ca2+-, but not in Na+-solutions, while it almost deleted all currents after mutation of the respective residue in IR8a. Although the authors argue that their experiments support the hypothesis that IRs function as ion channels, it could also be concluded that this residue appears to be important either for plasma membrane localization, for general protein structure, or for G-protein coupling, and cannot be specifically accounted for pore filter properties. Furthermore, it cannot be excluded that all odor-dependent currents observed are due to coupling of IR-complexes to G-proteins activating second messenger–dependent ion channels in the heterologous expression system.
In summary, expression of up to five different IRs per chemosensory neuron of sensilla coeloconica makes combinatorial heteromultimerization possible that might allow for an expansion of chemosensitivities and/or receptors for different stimulus concentration ranges. In vivo stoichiometry still awaits confirmation. Assembly with conserved coreceptors IR25a or IR8a is necessary for ciliary localization/maintenance. It is not known whether and how the conserved coreceptors participate in chemosensory transduction mechanisms or in the control of spontaneous activity. It is not known whether the coreceptors are regulated via cyclic nucleotides as suggested for the OR-specific coreceptor Orco.
(p. 357) The term “ionotropic receptor” (IR) could be misleading because at least some IRs are coupled to different G-proteins in crustaceans. Furthermore, there is still no convincing evidence that insect IRs function as ionotropic odor receptors. Because the pore region of the different members of this superfamily of receptors diverged either little or to quite some extent from the iGluRs, it remains to be studied whether specific members in different species retained ion channel function or not.
More Need for Speed or More Need for Sensitivity?
Given the controversy of whether GRs, ORs, and IRs employ ionotropic signal transduction via ligand-gated receptor-ion channel complexes or whether they are solely or additionally metabotropic receptors, different evolutionary pressures for fast or sensitive chemotransduction are recapitulated. Focus is the need for sensitive detection of sex pheromones by crepuscular moths that rely on maximized signal amplification for successful mating partner location (Kaissling, 2014). Furthermore, we discuss how fast the senses of smell and taste need to be to provide advantages for survival and reproduction. Maximal stimulus-triggered reaction times and maximized resolution of odor pulse frequencies are considered to estimate required speeds of chemodetection.
Sensitive Detection of Allelochemicals Such as Sex Pheromones Is Maximized in Crepuscular Moths
Different species employ allelochemicals such as pheromones, which can occur at very low concentrations in the pM range (Karlson & Lüscher, 1956; Whittaker & Feeny, 1971). The most sensitive chemoreception known in invertebrates is the detection of sex pheromones by crepuscular moths which employ GR-related inverse ORs. Female moths release a species-specific blend of sex pheromones in pulsatile fashion to attract their mates (for reviews, Stengl, 2010; Kaissling, 2014). Pairs of olfactory receptor neurons innervating thousands of long, hair-like trichoid sensilla on the moth’s antennae are specialized to detect the main pheromone components. Absolute sensitivity of ORNs was determined employing radiolabeled sex pheromones (bombykol) of the silkmoth Bombyx mori (Kasang, 1968) for extracellular electrophysiological recordings (tip recordings) of single, intact antennal pheromone-sensitive trichoid sensilla. It was calculated that one bombykol molecule was sufficient to elicit one action potential (for review, Kaissling, 2014). A similar sensitivity is documented for invertebrate photoreceptors that respond to one photon with one action potential (Scholes, 1965; Kirschfeld, 1966; Yau & Hardie, 2009). In 50% of the silkmoth males, weak pheromone stimuli (10−5 µg/fp) elicited a behavioral response within about 700 ms, while strong stimuli (10−2 µg/fp) aroused males within about 400 ms. With an estimated signal-to-noise ratio of ~6 (theoretical minimum = 3), the moth brain detects sex pheromones guiding the male mating behavior. Whereas the moth’s threshold for bombykol detection is ~3 × 103 molecules/ml, the most sensitive odorant detection for humans is 1.3 × 108 molecules/ml (sec-butyl mercaptan) and for dogs 9 × 103 molecules/ml (butyric acid) (Kaissling, 2014). Frogs smell with almost as much sensitivity as moths with a calculated detection threshold of 0.1 pM employing G-protein coupled ORs (Bhandawat et al., 2010). In frog ORNs it takes about 35 odorant binding events to trigger an action potential during one brief odor pulse. Only ~1.2 pA receptor current is sufficient to depolarize the resting potential-to-spike threshold. In comparison, the marine ragworm (polychaete) Nereis succinea detects and follows a trail of the tetrapeptide pheromone cysteinyl-glutathione at a threshold concentration of ~10−8 mol/l to search for the releasing female worms (Ram et al., 2008). At several magnitudes higher threshold concentration of 0.4 µl of a 10−5 M solution (1.7 ng), sperm release is induced. Also in C. elegans, chemoreceptors with different thresholds can trigger different behavioral responses (Taniguchi et al., 2014). In decapods crustaceans single-cell analysis with extracellular electrophysiological recordings from chemosensory neurons employing IRs recorded response thresholds as low as 10−12 M for odor detection and 10−4 M for gustatory stimuli (Schmidt & Gnatzy, 1984; for review, Atema, 1985). Sea urchin sperm which employs a receptor-guanylyl cyclase as chemoreceptor also displays extreme sensitivity with single-molecule detection (Pichlo et al., 2014).
The Speed of Invertebrate Chemoreception Lies in the Range of Milliseconds but Not Microseconds
How fast are chemical senses and how fast do odor- or taste-dependent behaviors need to be for different arthropods? Spitting out rotten, sour food should be elicited fast within a few milliseconds via reflexes to prevent poisoning. Because sour receptors are TRP-ion channel heteromers, it is likely that they (p. 358) employ fast, ionotropic mechanisms of signal transduction. Because ingested sour food is present at high stimulus concentrations, these taste receptors do not need to be very sensitive and can work at higher stimulus thresholds. Control of ion homeostasis, which strongly affects receptor potentials and osmotic balance, also needs to be regulated as quickly as possible. Accordingly, detection of salts employs ion channel receptors but of a different gene family: degenerin/epithelial sodium channels (Deg/ENaCs = Ppks) detect sodium salts, while different potassium channels are selective for potassium salts. Also the physiological concentrations of salts are rather high in the µM–mM range, making signal amplification superfluous.
Other decisions based on taste or olfactory stimuli do not need to be that fast. Bees in a hive tapping their antennae on a recruiting dancer with pollen from a new food source integrate information over seconds to minutes before they initiate a behavioral response (Gilley, 2014). In a natural airborn odor plume, odor contact with an insect antenna is brief (few ms) and frequent (up to 5 Hz) as measured with electroantennograms in flying (~3.5 m/s) or fixed insects (Vickers et al., 2001). A critical prerequisite for starting and maintaining the male moth’s characteristic zigzagging anemotaxis in search for their mates is the intermittency of the pheromone signal (Kennedy et al., 1981; Murlis & Jones, 1981; Willis & Baker, 1984; Baker et al., 1985; Baker & Haynes, 1989; Tumlinson et al., 1989; Vickers & Baker, 1992, 1994; Vickers, 2000; Koehl, 2006; Lei et al., 2009). Pulse frequencies of about 10 Hz allowed for almost straight upwind flight as compared to zigzagging flight patterns at lower pulse frequencies (for review, Vickers, 2006). Constant or very strong pheromone stimulation results in cessation of the male’s search, possibly caused by adaptation of the sensory cells (Baker et al., 1988). Also, odor stimulations with frequencies exceeding ~30 Hz are perceived as a constant odor stimulus and terminate upwind flight (Vickers, 2006). In wind tunnel experiments a moth zigzagging upwind to search for its sex-pheromone-releasing mate will switch within 300 ms to a casting search mode if it lost the pheromone trail (Vickers & Baker, 1994). Electrophysiological recordings of single pheromone-sensitive antennal trichoid sensilla confirmed that, depending on the species, moths can resolve the temporal structure of pulsed pheromone plumes between ~5 and 25 Hz (Bau et al., 2005). Antennogram measurements probing the very limits of temporal resolution of insect antennae determined odor response delays of only 2 ms and resolution of odor pulse frequencies of more than 100 Hz (Szyszka et al., 2014). Although these fast reaction times were not measured for ORNs in vivo (Cao et al., 2016) and these high frequencies may not be encountered in the natural surround, it is nevertheless very interesting that the population response of antennal sensory neurons can be that fast. Whereas vertebrates actively and regularly sample odors with their rhythmic breathing, invertebrates also sample actively and regularly. Lobsters in the water flick their antennules to maximize detection of odor concentration changes with a frequency of about 0.1–3.0 Hz (Schmitt & Ache, 1979; Koehl, 2006; Reidenbach & Koehl, 2011; Pravin & Reidenbach, 2013). Flying hawkmoths beat their wings at a frequency of up to 30 Hz, thereby producing turbulences which exchange the layer of air around the antennal sensilla (Tripathy et al., 2010; Daly et al., 2013). Thus, behaviorally measured reaction times match sampling frequencies for these species. Finally, because insect olfactory receptor neurons are not only ultradian but also circadian pacemakers, we need to consider that reaction times and odor detection thresholds differ daytime dependently (for review, Stengl, 2010). Perforated patch-clamp recordings of Drosophila IRs in vivo also confirmed reaction times of ORNs in the behaviorally relevant time range. In addition, they revealed distinct properties in adaptation and response kinetics between OR- and IR-expressing ORNs (Cao et al., 2016). While IR-expressing ORNs had odor response latencies of 35± 16 ms, OR-expressing ORNs responded slower with latencies of 66 ± 25 ms. Both response latencies suggest metabotropic rather than ionotropic odor transduction in the fruitfly.
In summary, odor detection of arthropods employing inverse ORs, IRs, or rGCs can be as sensitive as odor detection in vertebrates which employ G-protein-coupled ORs for signal amplification. Employing chemoreceptors with different response thresholds allows for the control of different behaviors dependent on stimulus concentration. The speed of odor detection is largely limited by stimulus-sampling behaviors such as antennal flicks of ~5 Hz or wing beating of ~30 Hz and by adaptation/disadaptation characteristics of phasic-tonic olfactory receptor neurons. Thus, reaction times of chemoreceptors and chemoreceptor neurons need to be in the range of milliseconds, but not microseconds for species with low stimulus sampling. Is there more need for high speed or high sensitivity? Because for different species the preferred sampling frequency as (p. 359) well as the odor concentrations for various odors or tastes differs over several orders of magnitude, the answer may not be the same for each species or even for each chemical signal detected via the same class of chemoreceptors. Therefore, I would expect that only chemosensory stimuli that require a fast response and are always present at high concentrations could select for ionotropic mechanisms, whereas costly, rare stimuli, like pheromones, require detection mechanisms of signal amplification such as receptor–G-protein coupling.
In the future, behavioral constraints and environmental conditions need to be considered for each species, and each chemical signal needs to be examined to obtain meaningful answers. In vitro analysis of receptor molecules is still important to reveal the molecular mechanics of chemoreceptors, but they cannot be taken as the sole basis for conclusions about their in vivo workings. In addition, it is not possible to generalize from a few examples of specialized species to the manifold of different taxa in their multitude of ecological niches. As odor-concentration-dependent and daytime-dependent behaviors indicate, it becomes increasingly evident that it is very important to pay attention to respective odor concentrations tested at different times of the day in different behavioral contexts since there may be different transduction cascades employed. Also, future experiments need to examine more closely amine/neuropeptide actions at the periphery since mechanisms of learning and memory in sensory neurons might not be very different from learning at central synapses. Mechanisms of gain control and expansion of response ranges tuned to physiological needs in ORNs and GRNs possibly mediated via neuropeptides and hormones are still far from understood. Thus, many interesting tasks still need to be tackled in arthropod olfaction next to the search for the transduction cascades of excitatory or inhibitory tastes and odors.
I thank Dr. Uwe Homberg, University of Marburg, for significant improvement of my writing. I thank Petra Gawaleck for supplying the figure and Christina Wollenhaupt for editing the References section.
Abuin, L., Bargeton, B., Ulbrich, M. H., Isacoff, Y., Kellenberger, S., & Benton, S. (2011). Functional architecture of olfactory ionotropic glutamate receptors. Neuron, 69, 44–60.Find this resource:
Ache, B. W., & Young, J. M. (2005). Olfaction: Diverse species, conserved principles. Neuron, 48, 417–430.Find this resource:
Afroz, A., Howlett, N., Shuzkla, A., Ahmad, F., Batista, E., Bedard, K., . . . Glendinning, J. I. (2013). Gustatory receptor neurons in Manduca sexta contain a TrpA1-dependent signaling pathway that integrates taste and temperature. Chemical Senses, 38(7), 605–617.Find this resource:
Al-Anzi, B., Tracey, W. D. Jr., & Benzer, S. (2006). “Response of Drosophila to wasabi is mediated by painless, the fly homolog of mammalian TRPA1/ANKTM1. Current Biology, 16(10), 1032–1040.Find this resource:
Altner, H., & Prillinger, L. (1980). “Ultrastructure of invertebrate chemo-, thermo,- and hygroreceptors and its functional significance. International Review of Cytology, 67, 69–139.Find this resource:
Atema, J. (1985). Chemoreception in the sea: adaptations of chemoreceptors and behaviour to aquatic stimulus conditions. Symposia of the Society for Experimental Biology, 39, 387–423.Find this resource:
Baker, T. C., Hansson, B. S., Löfstedt, C., & Löfqvist, J. (1988). Adaptation of antennal neurons in moths is associated with cessation of pheromone-mediated upwind flight. Proceedings of the National Academy of Sciences USA, 85, 9826–9830.Find this resource:
Baker, T. C., & Haynes, K. F. (1989). Field and laboratory electroantennographic measurements of pheromone plume structure correlated with oriental fruit moth behavior. Physiology and Entomology, 14, 1–12.Find this resource:
Baker, T. C., Willis, M., Haynes, K. F., & Phelan, P. L. (1985). A pulsed cloud of sex pheromone elicits upwind flight in male moths. Physiol. Enthomol. 10, 257–265.Find this resource:
Bargman, C. I. (2006). Comparative chemosensation from receptors to ecology. Nature, 444(16), 295–301.Find this resource:
Bau, J., Justus, K. A., Loudon, C., & Cardé, R. T. (2005). Electroantennographic resolution of pulsed pheromone plumes in two species of moths with bipectinate antennae. Chemical Senses, 30, 771–780.Find this resource:
Benton, R., Sachse, S., Michnick, S. W., & Vosshall, L. B. (2006). Atypical membrane topology and heteromeric function of Drosophila odorant receptors in vivo. PLoS Biology, 4, e20.Find this resource:
Benton, R., Vannice, K. S., Goez-Diaz, C., & Vosshall, L. B. (2009). Variant ionotropic glutamate receptors as chemosensory receptors in Drosophila. Cell, 136, 149–162.Find this resource:
Benton, R., Vannice, K. S., & Vosshall, L. B. (2007). An essential role for a CD36-related receptor in pheromone detection in Drosophila. Nature, 450, 289–293.Find this resource:
Bhandawat, V., Reisert, J., & Yau, K. W. (2010). Signaling by olfactory receptor neurons near threshold. Proceedings of the National Academy of Sciences USA, 107, 18682–18687.Find this resource:
Bobkov, Y. V., Pezier, A., Corey, E. Y., & Ache, B. W. (2010). Phosphatidylinositol 4,5. ExperimentalBiology, 213, 1417–1424.Find this resource:
Boekhoff, I., Michel, W. C., Breer, H., & Ache, B. W. (1994). Single odors differentially stimulate dual second messenger pathways in lobster olfactory receptor cells. Journal of Neuroscience, 14(5), 3304–3309.Find this resource:
Boekhoff, I., Seifert, E., Göggerle, S., Lindemann, M., Krüger, B.-W., & Breer, H. (1993). Pheromone-induced second messenger signaling in insect antennae. Insect Biochemistry and Molecular Biology, 23, 757–762.Find this resource:
Boekhoff, I., Strotmann, J., Raming, K., Tareilus, E., & Breer, H. (1990). Odorant-sensitive phospholipase C in insect antennae. Cell Signal, 2(1), 49–56.Find this resource:
(p. 360) Boto, T., Gomez-Diaz, C., & Alcorta, E. (2010). Expression analysis of the 3 G-protein subunits, Gα, Gβ, Gγ, in the olfactory receptor organs of adult Drosophila melanogaster. Chemical Senses, 35, 183–193.Find this resource:
Bredendiek, N., Hütte, J., Steingräber, A., Hatt, H., Gisselmann, G., & Neuhaus, E. M. (2011). Go α is involved in sugar perception in Drosophila. Chemical Senses, 26(1), 69–81.Find this resource:
Breer, H., Boekhoff, I., & Tareilus, E. (1990). Rapid kinetics of second messenger formation in olfactory transduction. Nature, 345, 65–68.Find this resource:
Brigaud, I., Montagne, N., Monsempes, C., Francois, M.-C., & Macquin-Joly, E. (2009). Identification of an atypical insect olfactory receptor subtype highly conserved within noctuids. FEBS, 276, 6537–6547.Find this resource:
Buck, L., & Axel, R. (1991). A novel multigene family may encode odorant receptors. A molecular basis for odor recognition. Cell, 65, 175–187.Find this resource:
Cameron, P., Hiroi, M., Ngai, J., & Scott, K. (2010). The molecular basis for water taste in Drosophila. Nature, 465, 91–95.Find this resource:
Cao, L.-H., Jing, B.-Y., Yang, D., Zeng, X., Shen, Y., & Tu, Y. (2016). Distinct signaling of Drosophila chemoreceptors in olfactory sensory neurons. Proceedings of the National Academy of Sciences USA, 113, E902–E911.Find this resource:
Carraher, C., Dalziel, J., Jordan, M. D., Christie, D. L., Newcomb, R. D., & Kralicek, A. V. (2015). Towards an understanding of the structural basis for insect olfaction by odorant receptors. Insect Biochemistry and Molecular Biology, 66, 31–41.Find this resource:
Chatterjee, A., Roman, G., & Hardin, P. E. (2009). Go contributes to olfactory reception in Drosophila melanogaster. BMC Physiology, 9(22), 1–7.Find this resource:
Chen, Z., Wang, Y., & Wang, Z. (2010). The amiloride-sensitive epithelial Na+ channel PPK28 is essential for Drosophila gustatory water reception. Journal of Neurosciences, 30, 6247–6252.Find this resource:
Clyne, P. J., Warr, C. G., & Carlson, J. R. (2000). Candidate taste receptors in Drosophila. Science, 287(5459), 1830–1834.Find this resource:
Clyne, P. J., Warr, C. G., Freeman, M. R., Lessing, D., Kim, J., & Carlson, J. R. (1999). A novel family of divergent seven-transmembrane proteins: candidate odorant receptors in Drosophila. Neuron, 22(2), 327–338.Find this resource:
Corey, E. A., Bobkov, Y., Pezier, A., & Ache, B. W. (2010). Phosphoinositide 3-kinase mediated signaling in lobster olfactory receptor neurons. Journal of Neuorchemistry, 113, 341–350.Find this resource:
Corey, E. A., Bobkov, Y., Ukhanov, K., & Ache, B. W. (2013). Ionotropic crustacean olfactory receptors. PLoS One, 8(4), 1–10 e60551.Find this resource:
Croset, V., Rytz, R., Cummins, S. F., Budd, A., Brawand, D., Kaessmann, H., Gibson, T. J., & Benton, R. (2010). Ancient protostome origin of chemosensory ionotropic glutamate receptors and the evolution of insect taste and olfaction. PLoS Genetics, 6(8): e1001064.Find this resource:
Dahanukar, A., Foster, K., van der Goes van Naters, W. M., & Carlson, J. R. (2001). A Gr receptor is required for response to the sugar trehalose in taste neurons of DrosophilaNature Neuroscience, 4, 1182–1186.Find this resource:
Dahanukar, A., Lei, Y. T., Kwon, J. Y., & Carlson, J. R. (2007). Two Gr genes underlie sugar reception in Drosophila. Neuron, 56(3), 503–516.Find this resource:
Daly, K. C., Kalwar, F., Hatfield, M., Staudacher, E., & Bradley, S. P. (2013). Odor detection in Manduca sexta is optimized when odor stimuli are pulsed at a frequency matching the wing beat during flight. PLoS One, 8, e8163.Find this resource:
Deng, Y., Zhang, W., Farhat, K., Oberland, S., Gisselmann, G., & Neuhaus, E. M. (2011). The stimulatory Gαs protein is involved in olfactory signal transduction in Drosophila. PLoS One, 6(4), e18605.Find this resource:
Derby, C. D., & Sorensen, P. W. (2008). Neural processing, perception, and behavioral responses to natural chemical stimuli by fish and crustaceans. Journal of Chemical Ecology, 34, 898–914.Find this resource:
Dethier, V. G. (1976). The hungry fly. Cambridge, MA: Harvard University Press.Find this resource:
Dobritsa, A. A., van der Goes van Naters, W., Warr, C. G., Steinbrecht, R. A., & Carlson, J. R. (2003). Integrating the molecular and cellular basis of odor coding in the Drosophila antenna. Neuron, 37(5), 827–841.Find this resource:
Doolin, R. E., & Ache, B. W. (2005). Cyclic nucleotide signaling mediates an odorant-suppressible chloride conductance in lobster olfactory receptor neurons. Chemical Senses, 30(2), 127–135.Find this resource:
Doolin, R. W., Zhainazarov, A. B., & Ache, B. W. (2001). An odorant-suppressed Cl- conductance in lobster olfactory receptor cells. Journal of Comparative Physiology A, 187(6), 477–487.Find this resource:
Dunipace, L., Meister, S., McNealy, C., & Amrein, H. (2001). Spatially restricted expression of candidate taste receptors in the Drosophila gustatory system. Current Biology, 11(11), 822–835.Find this resource:
Ebbs, M. L. & Amrein, H. (2007). Taste and pheromone perception in the fruit fly Drosophila melanogaster. European Journal of Physiology, 454, 735–747.Find this resource:
Elmore, T, Ignell, R., Carlson, J. R., & Smith, D. P. (2003). Targeted mutation of a Drosophila odor receptor defines receptor requirement in a novel class of sensillum. Journal of Neuroscience, 23(30), 9906–9912.Find this resource:
Fadool, D. A., & Ache, B. W. (1992). Plasma membrane inositol 1,4,5-trisphosphate–activated channels mediate signal transduction in lobster olfactory receptor neurons. Neuron, 9(5), 907–918.Find this resource:
Fadool, D. A., Estey, S. J., & Ache, B. W. (1995). Evidence that a Gq-protein mediates excitatory odor transduction in lobster olfactory receptor neurons. Chemical Senses, 20(5), 489–498.Find this resource:
Forstner, M., Gohl, T., Breer, H., & Krieger, J. (2006). Candidate pheromone binding proteins of the silkmoth Bombyx mori. Invertebrate Neuroscience, 6, 177–187.Find this resource:
Forstner, M., Gohl, T., Gondesen, I., Raming, K., Breer, H., & Krieger, J. (2008). Differential expression of SNMP-1 and SNMP-2 proteins in pheromone-sensitive hairs of moths. Chemical Senses, 33, 291–299.Find this resource:
Freeman, E. G., & Dahanukar, A. (2015). Molecular neurobiology of Drosophila taste. Current Trends in Neurobiology, 34, 140–148.Find this resource:
Freeman, E. G., Wisotsky, Z., & Dahanukar, A. (2014). Detection of sweet tastants by a conserved group of insect gustatory receptors. Proceedings of the National Academy of Sciences USA, 111(4), 1598–1603.Find this resource:
French, A. S., Sellier, M. J., Moutaz, A. A., Guigue, A., Chabaud, M. A., Reeb, P. D., . . . Marion-Poll, F. (2015). Dual mechanism for bitter avoidance in Drosophila. Journal of Neuroscience, 35, 3990–4004.Find this resource:
French, A. S., Torkkeli, T. H., & Schuckel, J. (2011). Dynamic characterization of Drosophila antennal olfactory neurons (p. 361) indicates multiple opponent signalling pathways in odor discrimination. Journal of Neuroscience, 19, 851–869.Find this resource:
Fuji, S., Yavuz, A., Slone, J., Jagge, C., Song, X., & Amrein, H. (2015). Drosophila sugar receptors in sweet taste perception, olfaction, and internal nutrient sensing. Current Biology, 25, 621–627.Find this resource:
Gao, Q., & Chess, A. (1999). Identification of candidate Drosophila olfactory receptors from genomic DNA sequence. Genomics, 60(1), 31–39.Find this resource:
German, P. F., van der Poel, S., Carraher, C., Kralicek, A. V., & Newcomb, R. D. (2013). Insights into subunit interactions within the insect olfactory receptor complex using FRET. Insect Biochememistry and Molecular Biology, 43(2), 138–145.Find this resource:
Gilley, D. C. (2014). Hydrocarbons emitted by waggle-dancing honey bees increase forager recruitment by stimulating dancing. PLoS One, 9(8), e105671. doi:10.1371/journal.pone.0105671.Find this resource:
Giribert, G. (2015). New animal phylogeny: Future challenges for animal phylogeny. Organisms Diversity and Evolution. doi:10.1007/s13127–015-0236–4.Find this resource:
Grosse-Wilde, E., Gohl, T., Bouche, E., Breer, H., & Krieger, J. (2007). Candidate pheromone receptors provide the basis for the response of distinct antennal neurons to pheromonal compounds. European Journal of Neuroscience, 25, 2364–2373.Find this resource:
Grosse-Wilde, E., Svatos, A., & Krieger, J. (2006). A pheromone-binding protein mediates the bombykol-induced activation of a pheromone receptor in vitro. Chemical Senses, 31, 547–555.Find this resource:
Guo, S., & Kim, J. (2010). Dissecting the molecular mechanism of Drosophila odorant receptors through activity modeling and comparative analysis. Proteins, 78(2), 381–399.Find this resource:
Hallem, E. A., Ho, M. G., & Carlson, J. R. (2004). The molecular basis of odor coding in the Drosophila antenna. Cell, 117, 965–979.Find this resource:
Harris, D. T., Kallman, B. R., Mullaney, B. C., & Scott, K. (2015). Representation of taste modality in the Drosohila brain. Neuron, 86, 1449–1460.Find this resource:
Ishimoto, H., & Tanimura, T. (2004). Molecular neurophysiology of taste in Drosophila. Cellular and Molecular Life Sciences, 61, 10–18.Find this resource:
Jeong, Y. T., Shim, J., Oh, S. R., Yoon, H., Kim, C. H., Moon, S. J., & Montell, C. (2013). An odorant-binding protein required for suppression of sweet taste by bitter chemicals. Neuron, 79, 725–737.Find this resource:
Jiao, Y., Moon, S. J., & Montell, C. (2007). A Drosophila gustatory receptor required for the responses to sucrose, glucose, and maltose identified by mRNA tagging. Proceedings of the National Academy of Sciences USA, 104, 14110–14115.Find this resource:
Jiao, Y., Moon, S. J., Wang, X., Ren, Q., & Montell, C. (2008). Gr64f is required in combination with other gustatory receptors for sugar detection in Drosophila. Current Biology, 18(22), 1797–1801.Find this resource:
Jin, X., Ha, T. S., & Smith, D. P. (2008). SNMP is a signaling component required for pheromone sensitivity in Drosophila. Proceedings of the National Academy of Sciences USA, 105, 10996.Find this resource:
Jones, P. L., Pask, G. M., Rinker, D. C., & Zwiebel, L. J. (2011). Functional agonism of insect odorant receptor ion channels. Proceedings of the National Academy of Sciences USA, 108(21), 8821–8825.Find this resource:
Jones, W. D., Cayirlioglu, P., Grunwald Kadow, I., & Vosshall, L. B. (2007). Two chemosensory receptors together mediate carbon dioxide detection in Drosophila. Nature, 445, 86–90.Find this resource:
Joseph, R. M., & Carlson, J. R. (2015). Drosophila chemoreceptors: a molecular interface between the chemical world and the brain. Trends in Genetics, 31(12), 683–695.Find this resource:
Kain, P., Chandrashekaran, S., Rodrigues, V., & Hasan, G. (2008). Drosophila mutants in phospholipid signaling have reduced olfactory responses as adults and larvae. Journal of Neurogenetics, 23(3), 303–312.Find this resource:
Kaissling, K.-E. (2009). Olfactory perireceptor and receptor events in moths: A kinetic model revised. Journal of Comparative Physiology A. Neuroethology Sensory Neural Behavioral Physiology, 195(10), 895–922.Find this resource:
Kaissling, K.-E. (2014). Pheromone reception in insects: The example of silk moths. InC. Mucignat-Caretta (Ed.), Neurobiology of chemical communication (pp. 99–145). Boca Raton, FL: CRC Press/Taylor & Francis.Find this resource:
Kang, K., Pulver, S. R., Panzano, V. C., Chang, E. C., Griffith, L. C., Theobald, D. L., & Garrity, P. A. (2010). Analysis of Drosophila TRPA1 reveals an ancient origin for human chemical nociception. Nature, 464, 597–600.Find this resource:
Karlson, P., & Lüscher, M. (1956). “Pheromones”: A new term for a class of biologically active substances. Nature (Lond.), 183, 55–56.Find this resource:
Kasang, G. (1968). Tritium labeling of the sex attractant Bobykol. Z. Naturforschung, 23b, 1331–1335.Find this resource:
Kato, A., & Touhara, K. (2009). Mammalian olfactory receptors: pharmacology, G protein coupling and desensitization. Cellular and Molecular Life Sciences, 66(23), 3743–3753.Find this resource:
Kato, S., Xu, Y., Cho, C. E., Abbott, L. F., & Bargmann, C. I. (2014). Temporal responses of C. elegans chemosensory neurons are preserved in behavioral dynamics. Neuron, 81, 616–628.Find this resource:
Keil, T. A., & Steinbrecht, R. A. (1984). Mechanosensitive and olfactory sensilla of insects. In R. C. King & H. Akai (Eds.), Insect ultrastructure, vol. 2. New York, NY: Plenum Press.Find this resource:
Kennedy, J. S., Ludlov, A. R., & Sanders, D. J. (1981). Guidance of flying male moths by wind-born sex pheromone. Physiology and Enthomology, 6, 395–412.Find this resource:
Kent, L. B., & Robertson, H. M. (2009). Evolution of the sugar receptors in insects. BMC Evolutionary Biology, 9, 41.Find this resource:
Kim, S. H., Lee, Y., Akitake, B., Woodward, O. M., Guggino, W. B., & Montell, C. (2010). Drosophila TRPA1 channel mediates chemical avoidance in gustatory receptor neurons. Proceedings of the National Academy of Sciences USA, 107, 8440–8445.Find this resource:
Kirschfeld, K. (1966). Discrete and graded receptor potentials in the coompound eye of the fly (Musca). In Proceedings of the international symposium on the functional organization of the compound eye (pp. 291–307). Oxford, England: Pergamon Press.Find this resource:
Koehl, M. A. R. (2006). The fluid mechanics of arthropod sniffing in turbulent odor plumes. Chemical Senses, 31, 93–105.Find this resource:
Koganezawa, M., & Shimada, I. (2001). Inositol 1,4,5-trisphosphate transduction cascade in taste reception of the fleshfly, Boettcherisca peregrina. Journal of Neurobiology, 51, 66–83.Find this resource:
Koh, T. W., He, Z., Gorur-Shandilya, S., Menuz, K., Larter, N. K., Steward, S., & Carlson, J. R. (2014). TheDrosophila IR20a clade of ionotropic receptors are candidate taste and pheromone receptors. Neuron, 83, 850–865.Find this resource:
Krieger, J., Gondesen, I., Forstner, M., Gohl, T., Dewer, Y. M., & Breer, H. (2009). HR11 and HR13 receptor-expressing (p. 362) neurons are housed together in pheromone-responsive sensilla trichodea of male Heliothis virescens. Chemical Senses, 34, 469–477.Find this resource:
Krieger, J., Grosse-Wilde, E., Gohl, T., & Breer, H. (2005). Candidate pheromone receptors of the silkmoth Bombyx mori. European Journal of Neuroscience, 21, 2167–2176.Find this resource:
Krieger, J., Klink, O., Mohl, C., Raming, K., & Breer, H. (2003). A candidate olfactory receptor subtype highly conserved across different insect orders. Journal of Comparative Physiology A, 189, 519–526.Find this resource:
Krieger, J., Raming, K., Dewer, Y. M. E., Bette, S., Conzelmann, S., & Breer, H. (2002). A divergent gene family encoding candidate olfactory receptors of the moth Heliothis virescens. European Journal of Neuroscience, 16, 619–628.Find this resource:
Kwon, J. Y., Dahanukar, A., Weiss, L. A., & Carlson, J. R. (2007). The molecular basis of CO2 reception in Drosophila. Proceedings of the National Academy of Sciences USA, 104, 3574–3578.Find this resource:
Kwon, J. Y., Dahanukar, A., Weiss, L. A., & Carlson, J. R. (2011). Molecular and cellular organization of the taste system in the Drosophila larva. Journal of Neuroscience, 31, 15300–15309.Find this resource:
Larsson, M. C., Domingos, A. L., Jones, W. D., Chiappe, M. E., Amrein, H., & Vosshall, L. B. (2004). Or83b encodes a broadly expressed odorant receptor essential for Drosophila olfaction. Neuron, 43, 703–714.Find this resource:
Laughlin, J. D., Ha, T. S., Jones, D. N. M., & Smith, D. P. (2008). Activation of pheromone-sensitive neurons is mediated by conformational activation of pheromone-binding protein. Cell, 133, 1255–1265.Find this resource:
Leal, W. S. (2012). Odorant reception in insects: Roles of receptors, binding proteins, and degrading enzymes. Annual Review of Entomology, 58, 373–391.Find this resource:
Leal, W. S., Chen, A. M., Ishida, Y., Chiang, V. P., Erickson, M. L., Morgan, T. I., & Tsuruda, J. M. (2005). Kinetics and molecular properties of pheromone binding and release. Proceedings of the National Academy of Sciences USA, 102, 5386–5391.Find this resource:
LeDue, E. E., Chen, Y.-C., Jung, A. Y., Dahanukar, A., & Gordon, M. D. (2015). Pharyngeal sense organs drive robust sugar consumption in Drosophila. Nature Communications, 6, 1–18.Find this resource:
Lee, J. K., & Strausfeld, N. J. (1990). Structure, distribution, and number of surface sensilla and their receptor cells on the antennal flagellum of the male sphinx moth Manduca sexta. Journal of Neurocytology, 19, 519–538.Find this resource:
Lee, Y., Kang, M. J., Shim, J., Cheong, C. U., Moon, S. J., & Montell, C. (2012). Gustatory receptors required for avoiding the insecticite L-canavanine. Journal of Neuroscience, 32, 1429–1435.Find this resource:
Lee, Y., Kim, S. H., & Montell, C. (2010). Avoiding DEET through insect gustatory receptors. Neuron, 67, 555–561.Find this resource:
Lee, Y., Moon, S. J., & Montell, C. (2009). Multiple gustatory receptors required for the caffeine response in Drosophila. Proceedings of the National Academy of Sciences USA, 106, 4495–4500.Find this resource:
Lei, H., Riffell, J. A., Gage, S. L., and Hildebrand, J. G. (2009). Contrast enhancement of stimululs intermittency in a primary olfactory network and its behavioral significance. Journal of Biology, 8(21), 1–15.Find this resource:
Li, Q., & Liberles, S. D. (2015). Aversion and attraction through olfaction. Current Biology, 25(3), F120–R129. doi:10.1016/j.cub.2014.11.044Find this resource:
Liman, E. R., Zhang, Y. V., & Montell, C. (2014). Peripheral coding of taste. Neuron, 81, 984–1000.Find this resource:
Liu, L., Johnson, W. A., & Welsh, J. J. (2003). Drosophila DEG/ENaC pickpocket genes are expressed in the tracheal system, where they may be involved in liquid clearance. Proceedings of the National Academy of Sciences USA, 100, 2128–2133.Find this resource:
Liu, J., Ward, A., Gao, J., Dong, Y., Nishio, N., Inada, H., . . . Xu, X. Z. (2010). C. elegans phototransduction requires a G protein-dependent cGMP-pathway and a taste receptor homolog. Nature Neuroscience, 13, 715–722.Find this resource:
Lundin, C., Käll, L., Kreher, S. A., Kapp, K., Sonnhammer, S. L., Carson, J. R., Heijne, G., & Nilsson, I. (2007). Membrane topology of the Drosophila OR83b odorant receptor. FEBS Letters, 581, 5601–5604.Find this resource:
Martin, J. P., Beyerlein, A., Dacks, A. M., Reisenman, C. E., Riffell, J. A., Lei, H., & Hildebrand, J. G. (2011). The neurobiology of insect olfaction: sensory processing in a comparative context. Progress in Neurobiology, 95(3), 427–447.Find this resource:
Masek, P., & Keene, A. C. (2013). Drosophila fatty acid taste signal through the PLC pathway in sugar-sensing neurons. PLoS Genetics, 9(9), e1003710. doi: 10.1371/journal.pgen.1003710.Find this resource:
McClintock, T. S., Ache, B. W., & Derby, C. D. (2006). Lobster olfactory genomics. Integrative and Comparative Biology, 46, 940–947.Find this resource:
Mishra, D., Miyamoto, T., Tezenom, Y. H., Broussard, A., Yavuz, A., Slone, J., Russell, D. H., & Amrein, H. (2013). The molecular basis of sugar sensing in Drosophila larvae. Current Biology, 23, 1466–1471.Find this resource:
Miyamoto, T., & Amrein, H. (2014). Diverse roles for the Drosophila fructose sensor Gr43a. Fly, 8(1), e27241, 1–7.Find this resource:
Miyamoto, T., Slone, J., Song, X., & Amrein, H. (2012). A fructose receptor functions as a nutrient sensor in the Drosophila brain. Cell, 151(5), 1113–1125.Find this resource:
Miyamoto, T., Wright, G., & Amrein, H. (2014). Nutrient sensors. Current Biology, 23, 369–373.Find this resource:
Mizunami, M., Hanmanaka, Y., & Nishino, H. (2015). Toward elucidating diversity of neural mechanisms underlying insect learning. Zoological Letters, 1(8). doi:10.1186/s40851–014-0008–6.Find this resource:
Moon, S. J., Kottgen, M., Jiao, Y., Xu, H., & Montell, C. (2006). A taste receptor required for the caffeine response in vivo. Current Biology, 16, 1812–1817.Find this resource:
Moon, S. J., Lee, Y., Jiao, Y., & Montell, C. (2009). A Drosophila gustatory receptor essential for aversive taste and inhibiting male-to male courtship. Current Biology, 19, 1623–1627.Find this resource:
Murlis, J., & Jones, C. (1981). Fine-scale structure of odour plumes in relation to insect orientation to distant pheromone and other attractant sources. Physiology and Entomology, 6, 71–86.Find this resource:
Nadasdy, Z. (2010). Binding by asynchrony: the neuronal phase code. Frontiers in Neuroscience, 4, 1–11.Find this resource:
Nakagawa, T., & Vosshall, L. (2009). Controversy and consensus: noncanonical signaling mechanisms in the insect olfactory system. Current Opinion Neurobiology, 19, 284–292.Find this resource:
Neuhaus, E. M., Gisselmann, G., Zhang, W., Dooley, R., Stortkuhl, K., & Hatt, H. (2005). Odorant receptor heterodimerization in the olfactory system of Drosophila melanogaster. Nature Neuroscience, 8(1), 15–17.Find this resource:
Ni, L., Bronk, P., Chang, E., Lowell, A. M., Flam, J. O., Panzano, V. C., Theobald, D. L., Griffith, L. C., & Garrity, P. A. (2013). A gustatory receptor paralogue controls rapid warmth avoidance in Drosophila. Nature, 500, 580–584.Find this resource:
(p. 363) Nolte, A., Funk, N. W., Mukunda, L., Gawalek, P., Werckenthin, A., Hansson, B. S., Wicher, D., & Stengl, M. (2013). In situ tip-recordings found no evidence for an Orco-based ionotropic mechanism of pheromone-transduction in Manduca sexta. PLoS One, 8(5): e62648. doi:10.1371/journal.pone.0062648.Find this resource:
Park, I. J., Hein, A. M., Bobkov, Y. V., Reidenbach, M. A., Ache, B. W., & Principe, J. C. (2016). Neurally encoding time for olfactory navigation. PLoS Computational Biology, 12(1), e1004742.Find this resource:
Pelosi, P., Iovinella, I., Felicioli, A., & Dani, F. R. (2014). Soluble proteins of chemical communication: an overview across arthropods. Frontiers in Physiology, 5, 320. doi: 10.3389/fphys.2014.00320.Find this resource:
Pelosi, P., Zhou, J. J., Ban, L. P., & Calvello, M. (2006). Soluble proteins in insect chemical communication. Cellular and Molecular Life Science, 63(14), 1658–1676.Find this resource:
Peñalva-Arana, D. C., Lynch, M., & Robertson, H. M. (2009). The chemoreceptor genes of the waterflea Daphnia pulex: many Grs but no Ors. BMC Evolutionary Biology, 9, 79.Find this resource:
Persuy, M.-A., Sanz, G., Tromelin, A., Thomas-Danguin, T., Gibrat, J.-F., & Pajot-Augy, E. (2015). Chapter one- mammalian olfactory receptors: Molecular mechanisms of odorant detection, 3D-modelling, and structure-activity relationships. Progress in Molecular Biology and Translational Science, 130, 1–36.Find this resource:
Pichlo, M., Bungert-Plümke, S., Weyand, I., Seifert, R., Bönigk, W., Strünker, T., . . . Collienne, U. (2014). High density and ligand affinity confer ultrasensitive signal detection by a guanylyl cyclase chemoreceptor. Journal of Cell Biology, 206, 541–557.Find this resource:
Pikielny, C. W. (2012). Sexy DEG/ENaC channels involved in gustatory detection of fruit fly pheromones. Science Signaling, 5, pe48.Find this resource:
Pophof, B. (2004). Pheromone-binding proteins contribute to the activation of olfactory receptor neurons in the silkmoths Antheraea polyphemus and Bombyx mori. Chemical Senses, 29(2), 117–125.Find this resource:
Pravin, S., & Reidenbach, M. A. (2013). Simultaneous sampling of flow and odorants by crustaceans can aid searches within a turbulent plume. Sensors (Basel), 13, 16591–16610.Find this resource:
Ram, J. L., Fei, X., Danaher, S. M., Lu, S., Breithaupt, T., & Hardege, J. D. (2008). Finding females: Pheromone-guided reproductive tracking behavior by male Nereis succinea in the marine environment. Journal of Experimental Biology, 211(Pt 5), 757–765.Find this resource:
Reidenbach, M. A., & Koehl, M. A. R. (2011). The spatial and temporal patterns of odors sampled by lobsters and crabs in a turbulent plume. Journal of Experimental Biology, 214, 3138–3153.Find this resource:
Robertson, H. M. (2015). The insect chemoreceptor superfamily is ancient in animals. Chemical Senses, 40, 609–614.Find this resource:
Robertson, H. M., Warr, C. G., & Carlson, J. R. (2003). Molecular evolution of the insect chemoreceptor gene superfamily in Drosophila melanogaster. Proceedings of the National Academy of Sciences USA, 100, 14537–14542.Find this resource:
Rogers, M. E., Sun, M., Lerner, M. R., & Vogt, R. G. (1997). SNMP-1, a noval membrane protein of olfactory neurons of the silk moth Antheraea polyphemus with homology to the CD36 family of membrane proteins. Journal of Biological Chemistry, 272, 14792–14799.Find this resource:
Rogers, M. E., Steinbrecht, R. A., & Vogt, R. G. (2001a). Expression of SNMP-1 in olfactory neurons and sensilla of male and female antennae of the silkmoth Antheraea polyphemus. Cell Tissue Research, 303, 433–446.Find this resource:
Rogers, M. E., Krieger, J., & Vogt, R. G. (2001b). Antennal SNMPs (sensory neuron membrane proteins) of Lepidoptera define a unique family of invertebrate CD36-like proteins. Journal of Neurobiology, 49, 47–61.Find this resource:
Rytz, R., Croset, V., & Benton, R. (2013). Ionotropic receptors (IRs): chemosensory ionotropic glutamate receptors in Drosophila and beyond. Insect Biochemistry and Molecular Biology, 43, 888–897.Find this resource:
Saina, M., Busengdal, H., Sinigaglia, C., Petrone, L., Oliveri, P., Rentzsch, R., & Benton, R. (2015). A cnidarians homologue of an insect gustatory receptor functions in developmental body patterning. Nature Communications, 6(6243), 1–24. doi:10.1038/ncomms7243.Find this resource:
Sakurai, T., Nakagawa, T., Mitsuno, H., Mori, H., Endo, Y., Tanoue, S., Yasukochi, Y., Touhara, K., & Nishioka, T. (2004). Identification and functional characterization of a sex pheromone receptor in the silkmoth Bombyx mori. European Journal of Neuroscience, 21, 2167–2176.Find this resource:
Sargsyan, V., Getahun, M. N., Llanos, S. L., Olsson, S. B., Hansson, B. S., & Wicher, D. (2011). Phosphorylation via PKC regulates the function of the Drosophila odorant co-receptor. Frontiers in Cell Neuroscience, 5(5). doi:10.3389/fncel.2011.00005.Find this resource:
Sato, K., Pellegrino, M., Nakagawa, T., Nakagawa, T., Vosshall, L. B., & Touhara, K. (2008). Insect olfactory receptors are heteromeric ligand-gated ion channels. Nature, 452, 1002–1006.Find this resource:
Sato, K., Tanaka, K., & Touhara, K. (2011). Sugar-regulated cation channel formed by an insect gustatory receptor. Proceedings of the National Academy of Sciences, 108(28), 11680–11685.Find this resource:
Schmitt, B. C., & Ache, B. W. (1979). Olfaction: Responses of a decapod crustacean are enhanced by flicking. Science, 205(4402), 204–206.Find this resource:
Schmidt, M., & Gnatzy, W. (1984). Are the funnel-canal organs the “campaniform sensilla” of the shore crab, Carcinus maenas (Decapoda, Crustacea)? II. Ultrastructure. Cell Tissue Research, 237(1), 81–93.Find this resource:
Scholes, J. (1965). Discontinuity of the excitation process in locust visual cells. Cold Spring Harbor Symposium in Quantative Biology, 30, 517–527.Find this resource:
Scott, K., Brady, R. Jr., Cravchik, A., Morozov, P., Rzhetsky, A., Zuker, C., & Axel, R. (2001). A chemosensory gene family encoding candidate gustatory and olfactory receptors in Drosophila. Cell, 104, 661–673.Find this resource:
Shen, K., Tootoonian, S., Laurent, G. (2013). Encoding of mixtures in a simple olfactory system. Neuron, 80(5), 1248–1262.Find this resource:
Slone, J., Daniels, J., & Amrein, H. (2007). Sugar receptors in Drosophila. Current Biology, 17, 1809–1816.Find this resource:
Smart, R., Kiely, A., Beale, M., Vargas, E., Carraher, C., Kraliecek, A. V., . . . Warr, C. G. (2008). Drosophila odorant receptors are novel seven transmembrane domain proteins that can signal independently of heterotrimeric G proteins. Insect Biochemistry and Molecular Biology, 38, 770–780.Find this resource:
Steinbrecht, R. A., Laue, M., & Ziegelberger, G. (1995). Immunocytochemistry of odorant-binding protein and general binding protein in olfactory sensilla of the silk moths Antheraea and Bombyx. Cell Tissue Research, 282, 203–217.Find this resource:
Stengl, M. (1993). Intracellular-messenger-mediated cation channels in cultured olfactory receptor neurons. Journal of Experimental Biology, 178, 125–147.Find this resource:
(p. 364) Stengl, M. (1994). Inositol-trisphosphate-dependent calcium currents precede cation currents in insect olfactory receptor neurons. Journal of Comparative Physiology A, 174, 187–194.Find this resource:
Stengl, M. (2010). Pheromone transduction in moths. Frontiers in Cell Neuroscience, 4, 133.Find this resource:
Stengl, M., & Funk, N. W. (2013). The role of the coreceptor Orco in insect olfactory transduction. Journal of Comparative Physiology A, 199, 897–909.Find this resource:
Stengl, M., & Hildebrand, J. G. (1990). Insect olfactory neurons in vitro: Morphological and immunocytochemical characterization of male-specific antennal receptor cells from developing antennae of male Manduca sexta. Journal of Neuroscience, 10, 837–847.Find this resource:
Stengl, M., Zintl, R., de Vente, J., & Nighorn, A. (2001). Localization of cGMP-immunoreactivity and of soluble guanylyl cyclase in antennal sensilla of the hawkmoth Manduca sexta. Cell Tissue Research, 304, 409–421.Find this resource:
Stengl, M., Zufall, F., Hatt, H., & Hildebrand, J. G. (1992). Olfactory receptor neurons from antennae of developing male Manduca sexta respond to components of the species-specific sex pheromone in vitro. Journal of Neuroscience, 12, 2523–2531.Find this resource:
Stephanyan, R., Day, K. Urban, J., Harding, D. L., Shetty, R. S., Derby, C. D., Ache, B. W., & McClintock, T. S. (2006). Gene expression and specificity in the mature zone of the lobster olfactory organ. Physiology and Genomics, 25(2), 224–233.Find this resource:
Stocker, R. F. (1994). The organization of the chemosensory system in Drosophila melanogaster: A review. Cell Tissue Research, 275, 3–26.Find this resource:
Stocker, R. F. (2001). Drosophila as a focus in olfactory research: mapping of olfactory sensilla by fine structure, odor specificity, odorant receptor expression, and central connectivity. Microscopy Research and Technique, 55(5), 284–296.Find this resource:
Stopfer, M., Jayaraman, V., & Laurent, G. (2003). Intensity versus identity coding in an olfactory system. Neuron, 39(6), 991–1004.Find this resource:
Szyszka, P., Gerkin, R. C., Galizia, C. G., & Smith, B. H. (2014). High-speed odor transduction and pulse tracking by insect olfactory receptor neurons. Proceedings of the National Academy of Sciences USA, 111(47), 16925–16930.Find this resource:
Talluri, S., Bhatt, A., & Smith, D. P. (1995). Identification of a Drosophila G protein alpha subunit (dGq alpha-3) expressed in chemosensory cells and central neurons. Proceedings of the National Academy of Sciences USA, 92, 11475–11479.Find this resource:
Taniguchi, G., Uozumi, T., Kiriyama, K., Kamizaki, T., & Hirotsu, T. (2014). Screening of odor-receptor pairs in Caenorhabditis elegans reveals different receptors for high and low odor concentrations. Science Signaling, 7(323), ra39. doi:10.1126/scisignal.2005136.Find this resource:
Thorne, N., & Amrein, H. (2008). Atypical expression of Drosophila gustatory receptor genes in sensory and central neurons. Journal of Comparative Neurology, 506, 548–568.Find this resource:
Tripathy, S., Peters, O. J., Staudacher, E. M., Kalwar, F. R., Hatfield, M. N., & Daly, K. C. (2010). Odors pulsed at wing beat frequencies are tracked by primary olfactory networks and enhance odor detection. Frontiers in Cellular Neuroscience, 16(4), 1.Find this resource:
Tsitoura, P., Andronopoulou, E., Tsikou, D., Agalou, A., Papakonstantinou, M. P., Kotzia, G. A. . . . Iatrou, K. (2010). Expression and membrane topology of Anopheles gambiae odorant receptors in lepidopteran insect cells. PLoS One, 5(11), e15428Find this resource:
Tumlinson, J. H., Brennan, M. M., Doolittle, R. E., Mitchell, E. R., Brabham, A., Mazomemos, B. E., . . . Jackson, D. M. (1989). Identification of a pheromone blend attractive to Manduca sexta (L.) males in a wind tunnel. Archives in Insect Biochem. Physiology, 10, 255–271.Find this resource:
Ueno, K., & Kidokoro, Y. (2008). Adenylyl cyclase encoded by AC78C participates in sugar perception in Drosophila melanogaster. European Journal of Neuroscience, 28, 1956–1966.Find this resource:
Ueno, K., Kohatsu, S., Clay, C., Forte, M., Isono, K., & Kidokoro, Y. (2006). Gsα is involved in sugar perception in Drosophila melanogaster. Journal of Neuroscience, 26(23), 6143–6152.Find this resource:
Ueno, K., Ohta, M., Morita, H., Mikuni, Y., Nakajima, S., Yamamoto, K., & Isono, K. (2001). Trehalose sensitivity in Drosophila correlates with mutations in and expression of the gustaotry receptor gene Gr5a. Current Biology, 11(18), 1451–1455.Find this resource:
Vermehren, A., Langlais, K. K., & Morton, D. B. (2006). Oxygen-sensitive guanylyl cyclases in insects and their potential roles in oxygen detection and in feeding behaviors. Journal of Insect Physiology, 52, 340–348.Find this resource:
Vermehren-Schmaedick, A., Scudder, C., Timmermans, W., & Morton, D. B. (2011). Drosophila gustatory preference behaviors require the atypical soluble guanylyly cyclases. Journal of Comparative Physiology A, 197, 717–727.Find this resource:
Vickers, N. J. (2000). Mechanisms of animal navigation in odor plumes. Biology Bulletin, 198(2), 203–212.Find this resource:
Vickers, N. J. (2006). Winging it: Moth flight behavior and responses of olfactory neurons are shaped by pheromone plume dynamics. Chemical Senses, 31, 155–166.Find this resource:
Vickers, N. J., & Baker, T. C. (1992). Male Heliothis virescens maintain upwind flight in response to experimentally pulsed fiaments of their sex pheromone (Lepidoptera: Noctuidae). Journal of Insect Behavior, 5, 669–687.Find this resource:
Vickers, N. J., & Baker, T. C. (1994). Reiterative responses to single strands of odor promote sustained upwind flight and odor source location by moths. Proceedings of the National Academy of Sciences USA, 91, 5756–5760.Find this resource:
Vickers, N. J., Christensen, T. A., Baker, T. C., & Hildebrand, J. G. (2001). Odour-plume dynamics influence the brains´s olfactory code. Nature, 410, 466–470.Find this resource:
Vogt, R. G., & Riddiford, L. M. (1981). Pheromone binding and inactivation by moth antennae. Nature, 293, 161–163.Find this resource:
Vogt, R. G., & Riddiford, L. M. (1986). Pheromone reception: A kinetic equilibrium. In T. L. Payne, M. C. Birch, & E. J. Kennedy (eds.)., Mechanisms in insect olfaction (pp. 201–208). Oxford, UK: Clarendon.Find this resource:
Vogt, R. G., Miller, N. E., Litvack, R., Fandino, R. A., Sparks, J., Staples, J., . . . Dickens, J. C. (2009). The insect SNMP gene family. Insect Biochemistry and Molecular Biology, 39(7), 448–456.Find this resource:
Vosshall, L. B., Amrein, H., Morozov, P. S., Rzhetsky, A., & Axel, R. (1999). A spatial map of olfactory receptor expression in the Drosophila antenna. Cell, 96, 725–736.Find this resource:
Vosshall, L. B., & Hansson, B. S. (2011). A unified nomenclature system for the insect olfactory coreceptor. Chemical Senses, 36(6), 497–498.Find this resource:
Weiss, L. A., Dahanukar, A., Kwon, J. Y., Banerjee, D., & Carlson, J. R. (2011). The molecular and cellular basis of bitter taste in Drosophila. Neuron, 69(2), 258–272.Find this resource:
Wetzel, C. H., Behrendt, H. J., Gisselmann, G., Stortkuhl, K. F., Hovemann, B., & Hatt, H. (2001). Functional expression (p. 365) and characterization of a Drosophila odorant receptor in a heterologous cell system. Proceedings of the National Academy of Sciences USA, 98(16), 9377–9380.Find this resource:
Whittaker, R. H., & Feeny, P. P. (1971). Allelochemics: Chemical interactions between species. Science, 171(3973), 757–770.Find this resource:
Wicher, D. (2015). Olfactory signaling in insects. Progress in Molecular Biology and Translation Sciences, 130, 37–54.Find this resource:
Wicher, D., Schäfer, R., Bauernfeind, R., Stensmyr, M. C., Heller, R., Heinemann, S. H., & Hansson, B. S. (2008). Drosophila odorant receptors are both ligand-gated and cyclic-nucleotide-activated cation channels. Nature, 452, 1007–1011.Find this resource:
Willis, M. A., & Baker, T. C. (1984). Effects of intermittent and continuous pheromone stimulation on the flight behavior of the oriental fruit moth, Grapholita molesta. Physiology and Entomology, 9, 341–354.Find this resource:
Wistrand, M., Kall, L., & Sonnhammer, E. L. (2006). A general model of G protein-coupled receptor sequences and its application to detect remote homologs. Protein Sciences, 15(3), 509–521.Find this resource:
Xiang, Y., Yuan, Q., Vogt, N., Looger, L. L., Jan, L. Y., & Jan, Y. N. (2010). Light-avoidance mediating phtotoreceptors tile the Drosophila larval body wall. Nature, 468, 921–926.Find this resource:
Xu, P. X., Atkinson, R., Jones, D. N. M., & Smith, D. P. (2005). Drosophila OBP LUSH is required for activity of pheromone-sensitive neurons. Neuron, 45, 193–200.Find this resource:
Xu, W., Papanicolau, A., Liu, N. Y., Dong, S. L., & Anderson, A. (2014). Chemosensory receptor genes in the oriental tobacco budworm Helicoverpa assulta. Insect Molecular Biology, 24(2), 253–263.Find this resource:
Yao, C. A., & Carlson, J. R. (2010). Role of G-proteins in odor-sensing and CO2-sensing neurons in Drosophila. Journal of Neuroscience, 30(13), 4562–4572.Find this resource:
Yau, K. W., & Hardie, R. C. (2009). Phototransduction motifs and variations. Cell, 139, 246–264.Find this resource:
Zhang, H. J., Anderson, A. R., Trowell, S. C., Luo, A. R., Xiang, Z. H., & Xia, Q. Y. (2011). Topological and functional characterization of an insect gustatory receptor. PLoS One, 6(8): e24111.Find this resource:
Zhang, J., Liu, Y., Walker, W. B., Dong, S.-L., & Wang, G.-R. (2015). Identification and localization of two sensory neuron membrane proteins from Spodoptera litura (Lepidoptera: Noctuidae). Insect Sciences, 22, 399–408.Find this resource:
Zhang, Y. V., & Montell, C. (2013). The molecular basis for attractive salt-taste coding in Drosophila. Science, 340, 1334–1338.Find this resource:
Zhang, Y. V., Raghuwanshi, R. P., Shen, W. L., & Montell, C. (2013). Food experience-induced taste desensitization modulated by the Drosphila TRPL channel. Nature Neuroscience, 16, 1468–1476.Find this resource:
Ziegelberger, G. (1995). Redox-shift of the pheromone-binding protein in the silkmoth Antheraea ployphemus. European Journal of Biochemistry, 232, 706–711.Find this resource:
Ziegelberger, G., Van den Berg, M. J., Kaissling, K.-E., Klumpp, S., & Schultz, J. E. (1990). Cyclic nucleotide levels and guanylate cyclase activity in pheromone-sensitive antennae of the silkmoths Antheraea polyphemus and Bombyx mori. Journal of Neuroscience, 10, 1217–1225. (p. 366) Find this resource: