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date: 19 January 2018

Caenorhabditis elegans Olfaction

Summary and Keywords

To survive, animals must properly sense their surrounding environment. The types of sensation that allow for detecting these changes can be categorized as tactile, thermal, aural, or olfactory. Olfaction is one of the most primitive senses, involving the detection of environmental chemical cues. Organisms must sense and discriminate between abiotic and biogenic cues, necessitating a system that can react and respond to changes quickly. The nematode, Caenorhabditis elegans, offers a unique set of tools for studying the biology of olfactory sensation.

The olfactory system in C. elegans is comprised of 14 pairs of amphid neurons in the head and two pairs of phasmid neurons in the tail. The male nervous system contains an additional 89 neurons, many of which are exposed to the environment and contribute to olfaction. The cues sensed by these olfactory neurons initiate a multitude of responses, ranging from developmental changes to behavioral responses. Environmental cues might initiate entry into or exit from a long-lived alternative larval developmental stage (dauer), or pheromonal stimuli may attract sexually mature mates, or repel conspecifics in crowded environments. C. elegans are also capable of sensing abiotic stimuli, exhibiting attraction and repulsion to diverse classes of chemicals. Unlike canonical mammalian olfactory neurons, C. elegans chemosensory neurons express more than one receptor per cell. This enables detection of hundreds of chemical structures and concentrations by a chemosensory nervous system with few cells. However, each neuron detects certain classes of olfactory cues, and, combined with their synaptic pathways, elicit similar responses (i.e., aversive behaviors). The functional architecture of this chemosensory system is capable of supporting the development and behavior of nematodes in a manner efficient enough to allow for the genus to have a cosmopolitan distribution.

Keywords: olfaction, chemosensation, connectomics, GPCRs, neural circuits

In its simplest form, olfaction requires a receptor to detect a chemical and transduce a signal within the cell detecting the cue. The evolution of eukaryotic nervous systems has led to the production of specialized cells, the olfactory neurons, which sense, process, and communicate chemical changes sensed in the environment and transmit information to the rest of the organism.

Studies of olfaction in humans and other mammals have found that the olfactory neurons are extremely specialized, and express only one odorant receptor per cell, following a “one neuron, one receptor” rule (Bargmann, 2006b; Bear, Lassance, Hoekstra, & Datta, 2016; Chess, Simon, Cedar, & Axel, 1994; Serizawa et al., 2003). However, these receptors are not specific themselves, and may detect multiple odorants. While much can be learned studying these systems, researchers also investigate evolutionarily distinct olfactory systems, using the fruit fly Drosophila melanogaster and the nematode Caenorhabditis elegans. Both species offer useful benefits, such as nervous systems that are easily manipulated. The nervous system of Drosophila contains approximately 150,000 neurons (Jenett et al., 2012), compared to approximately 86 billion neurons found in human brains (Azevedo et al., 2009). The C. elegans nervous system, meanwhile, has only 302 neurons (White, Southgate, Thomson, & Brenner, 1986) whose lineal origins and connectivity are known, and therefore is an extremely compact and experimentally tractable nervous system. In C. elegans, approximately 36 neurons contribute to olfaction, and the “one neuron, one receptor” rule is not followed; multiple olfactory receptors are present in individual neurons. Despite this, the molecular mechanisms of olfaction remain conserved, with G protein-coupled receptors, receptor guanylate cyclases, and intracellular signaling cascades sensing and initiating responses to many chemical cues in the environment (Bargmann, 2006a). Studying the C. elegans olfactory network will allow for generation of an in-depth understanding of a complex neural coding strategy: How does this small number of olfactory neurons sense and integrate a myriad of olfactory signals to generate the robust behaviors observed?

Caenorhabditis elegans

The nematode Caenorhabditis elegans is eutelic—every animal has the same number of somatic cells. As such, every cell lineage has been mapped, from fertilized zygote to fully developed adult (Sulston & Horvitz, 1977). As such, the structure of the entire nervous system, comprised of 302 neurons, has been determined using electron micrographs (Ward, Thomson, White, & Brenner, 1975; Ware, Clark, Crossland, & Russell, 1975; White, Southgate, et al., 1986). In recent years, researchers have started characterizing the network’s functional connections(Azulay, Itskovits, & Zaslaver, 2016; Bargmann, 2012; Bargmann & Marder, 2013; Hong & Park, 2016; Rengarajan & Hallem, 2016; Sohn, Choi, Ahn, Lee, & Jeong, 2011; Towlson, Vertes, Ahnert, Schafer, & Bullmore, 2013).

A small proportion of C. elegans are male (0.01%–0.02%) (Hodgkin, 1983), which are morphologically distinct from hermaphrodites. Males do not carry eggs and exhibit a fan-shaped tail (Barr & Garcia, 2006). Besides these gross morphological differences, the male nervous system is comprised of 385 neurons, 89 of which are sex-specific (Sammut et al., 2015; Sulston, Albertson, & Thomson, 1980). Only a small number of these sex-specific neurons are thought to function in olfaction. How the addition of these sex-specific neurons changes the connectome of the nervous system has not yet been fully explained (Fagan & Portman, 2014; Garcia & Portman, 2016; M. P. Hart & Hobert, 2015; Portman, 2017; Serrano-Saiz et al., 2017).

C. elegans has become a leading model for understanding olfactory nervous systems, as it has a fully mapped hermaphroditic nervous system, is transparent, and is amenable to genetic manipulations. The nematode allows for the study of mechanisms underlying olfaction using genetically encoded calcium indicators (GECIs) to measure real-time neural dynamics in live animals (Chokshi, Bazopoulou, & Chronis, 2010; Hilliard, Apicella, Kerr, Suzuki, Bazzicalupo, & Schafer, 2005; Schrodel, Prevedel, Aumayr, Zimmer, & Vaziri, 2013; Tatro, 2014; Touhara & Vosshall, 2009). Combining the robust behavioral responses of nematodes with the ease of generating genetic mutants, understanding olfactory mechanisms has dramatically increased through the use of this roundworm.

In their natural milieu—usually rotting fruits and vegetables (Félix & Braendle, 2010; Schulenburg & Felix, 2017; Teotonio, Estes, Phillips, & Baer, 2017)—C. elegans individuals are exposed to a large variety of chemical cues, which signal information about their environment. These cues range from gases and volatiles, to water-soluble compounds, and from abiotic to biogenic cues.

Olfactory Neuronal Anatomy

Caenorhabditis elegans OlfactionClick to view larger

Figure 1. C. elegans Olfactory Anatomy. (A) The Chemosensory Amphid Olfactory Sensilla: The ciliated amphid olfactory neurons develop as bilateral pairs with three distinct cilia morphologies: single-rod (blue), double-rod (white), and winged (orange). The dendritic extensions for the left-side neurons are shown in matching colors. The CEM (brown) are male-specific neurons that exhibit radial symmetry, with both dorsal and ventral left-right pairs. (B) The Oxygen and Carbon Dioxide Sensing Amphid Sensilla: The BAG neurons (pink) sense carbon dioxide and develop bag-shaped cilia (also shown in pink). The URX neurons (yellow) contribute to oxygen sensation, with dendrites that do not develop cilia and that terminate in the pseudocoelomic fluid. URX sensation of oxygen is aided by the AQR neuron (blue), which develops as a single neuron, with a dendrite that wraps around the pharyngeal tube. The IL2 neurons (green) develop as three bilateral pairs (dorsal, lateral, and ventral), with cilia exposed to the environment. (C) The Phasmid Olfactory Sensilla. The PQR neuron (blue) contributes to the gas sensing network, alongside the URX and AQR neurons located in the amphid region. The two chemosensory phasmid neurons, PHA (green) and PHB (orange), extend their cilia out to the external environment, and modulate turns initiated by amphid olfactory neurons.

Although olfaction occurs at both ends of the nematode (Hilliard, Bargmann, & Bazzicalupo, 2002), most research has focused on the neurons within major anterior sensilla, the amphids. The amphids are a pair of channels that each contain sensory dendrites of 12 sensory neurons (Ward et al., 1975; Ware et al., 1975; White et al., 1986) (Figure 1A). Eleven of the amphid neuron-types are chemosensory. In addition to amphid olfactory neurons, the head contains other olfactory neurons: BAG and URX, which are involved in sensing carbon dioxide and oxygen, respectively (Figure 1B), with the aid of the AQR and PQR neurons (Gray et al., 2004).

The IL2 neurons, located anterior to the amphid olfactory neurons, are not present as a bilateral pair, but instead are present as a set of six neurons, with dorsal, ventral, and lateral pairs (Figure 1B). The lateral neurons display a connectivity different from that of the remaining four neurons, further complicating the elucidation of their role (Wang, Schwartz, & Barr, 2010). There are neurons in the tail (phasmids) that sense odorants, though the number of chemosensory neurons in the tail is drastically lower than that seen in the anterior, with only two pairs of phasmid neurons being present that contain cilia exposed to the external environment (Figure 1C).

The Male Nervous System

Of the 89 sex-specific neurons present in the male nervous system, only four located in the amphid region participate in olfaction: the cephalic male (CEM) neurons (White, Nicholas, Gritton, Truong, Davidson, & Jorgensen, 2007). These radially symmetric neurons exhibit dorsal/ventral as well as left/right symmetry, with cilia exposed to the external environment alongside the hermaphroditic amphid olfactory neurons (Figure 1A). The clearly distinguishable male-tail is heavily involved in the mating process and includes the majority of the male-specific neurons. Thirty-six ray neurons innervate this structure (Liu & Sternberg, 1995; White et al., 2007).

The Nature of Amphid Olfactory Neurons

Winged Cilia Olfactory Neurons

Although not exposed to the external environment, winged ciliated neurons are involved in the sensation of volatile cues, which diffuse across the cuticle (Wes & Bargmann, 2001; White et al., 1986). C. elegans is able to distinguish between seven classes of these odors. Most of these volatiles were found to be products of bacterial metabolism, suggesting a biologically relevant role for AWA and AWC—food chemotaxis (Bargmann, Hartwieg, & Horvitz, 1993). Most neurons in C. elegans are present as anatomically symmetrical, bilateral pairs. However, AWC has been shown to exhibit stochastic asymmetry in its gene expression and sensing abilities (Cochella et al., 2014; Pierce-Shimomura, Morse, & Lockery, 1999; Troemel, Kimmel, & Bargmann, 1997; Wes & Bargmann, 2001; Yu, Avery, Baude, & Garbers, 1997).

One of the repulsion-driving olfactory neurons, AWB, also senses volatiles, such as 1-octanol (Troemel, Kimmel, & Bargmann, 1997). The biological relevance of 1-octanol in natural environments can be debated, as it has only been found in the extracts from some enteric Gram negative bacteria (Elgaali, Hamilton-Kemp, Newman, Collins, Yu, & Archbold, 2002). However, given that the volatile metabolites emitted by bacteria are complex mixtures (Hamilton-Kemp, Newman, Collins, Elgaali, Yu, & Archbold, 2005), and the natural food sources of nematodes are incompletely known, 1-octanol serves as a reliable stimulus for AWB in experimental conditions.

The Single- and Double-Rod Ciliated Amphid Neurons

Water-soluble attractants and repellents are sensed by the single- and double-rod ciliated amphid neurons (Table 1). These neurons contain rod-shaped cilia which extend through the amphid sheath and cuticle to sense odorants in the external environment (Bargmann, 2006a; Ward et al., 1975). Like AWC, ASE is asymmetric, but it is more consistent in its asymmetry than AWC, with the right neuron (ASER) always sensing Cl- and K+ ions, and ASEL sensing Na+ (Pierce-Shimomura et al., 1999).

ASH is unique among the amphid sensilla in that it is polymodal and acts as the key nociceptor in C. elegans (Chatzigeorgiou, Bang, Hwang, & Schafer, 2013; de Bono & Maricq, 2005; Hukema, Rademakers, Dekkers, Burghoorn, & Jansen, 2006; Komuniecki, Harris, Hapiak, Wragg, & Bamber, 2012; Walker, Vázquez-Manrique, Gower, Gregory, Schafer, & Baylis, 2009). While most neurons sense an odorant and elicit a general response downstream, ASH’s response is more complex, as the downstream signaling exhibits specificity to the stimulus sensed. For example, while OCR-2 and OSM-9 (see Downstream Ion Channels Involved in Olfaction section) are required for all responses within the ASH Neurons, the g-proteins utilized for different GPCRs varies (i.e., either GPA-3 or ODR-3 can function to generate a neuronal response) (Hilliard, Bergamasco, Arbucci, Plasterk, & Bazzicalupo, 2004; Walker et al., 2009).

Largely involved in developmental processes, ASI assists in controlling entry into dauer. ASI is the sole source of DAF-7 and TGF-β‎ in C. elegans, which act to prevent dauer entry (Meisel, Panda, Mahanti, Schroeder, & Kim, 2014; Ren, Lim, Johnsen, Albert, Pilgrim, & Riddle, 1996; Schackwitz, Inoue, & Thomas, 1996). This signaling pathway is regulated by the availability of food, population density, carbon dioxide levels, the presence of dauer pheromone, and temperature, while more recent work has expanded this list to include mRNA decay pathways (Androwski, 2017; Borbolis, Flessa, Roumelioti, Diallinas, Stravopodis, & Syntichaki, 2017). ASI is also required for proper dauer exit and resumption of normal development after stress conditions are mitigated (Ren et al., 1996), as well as being required for withdrawal from noxious stimuli(Mills et al., 2016). The olfactory regulation of dauer control arises through ASI sensation of the majority of daumone constituents: ascr#2, ascr#3, and ascr#5 (Kim et al., 2009; McGrath, Xu, Ailion, Garrison, Butcher, & Bargmann, 2011; Park et al., 2012).

The double-ciliated neuron, ADL, has been shown to sense ascr#3 as well. However, instead of initiating dauer entry or avoidance behaviors, ADL regulates body fat content (Hussey et al., 2017).

The ASK neuron functions in driving both avoidance and attractive behaviors. Removal of ASH through laser ablation experiments results in ASK gaining the ability to sense many aversive stimuli (Hilliard et al., 2002; Hukema et al., 2006; Sambongi et al., 1999). ASK plays a major role in sensing attractive biogenic cues. Icas#1, #3, and #9, indolated derivatives of ascr#1, #3, and #9, respectively, were shown to attract C. elegans of both sexes (Srinivasan et al., 2012). However, this attraction is extremely concentration-dependent across the sexes, with males no longer attracted to low concentrations. ASK has also been shown to sense ascr#3, which is repulsive to hermaphrodites at concentrations that attract males. This likely arises from the combined output of ASK in males with the chemoattraction that is driven by the male-specific CEM neurons (Narayan et al., 2016; Pungaliya et al., 2009; Srinivasan et al., 2008; White & Jorgensen, 2012).

ADF is unique in that it is the only serotonergic sensory neuron in C. elegans hermaphrodites (Sze, Victor, Loer, Shi, & Ruvkun, 2000). Little is known about what ADF may be sensing in the surrounding environment, although it has been shown to contribute to dauer control, as animals lacking ADF display aberrant dauer repression (Schackwitz et al., 1996).

Little has been studied concerning the role of the ASG and ASJ neurons. ASJ is known to contribute to dauer entry and recovery, but what exactly it senses in the environment to initiate these developmental changes remains unknown. ASJ has also been shown to be involved in the sensation of pathogenic bacteria (Meisel et al., 2014). It is likely that, like ASI, these neurons sense a combination of pheromone cues, population density cues, and information about food availability. This, however, remains to be determined.

Male-Specific Ciliated Amphid Olfactory Neurons

Male C. elegans contain four extra ciliated neurons that function as olfactory neurons—the CEM neurons. Ascarosides #3 and #8, both of which contribute to the induction of dauer, also elicit repulsion of hermaphrodites, yet attract males, are sensed by these neurons (Jang et al., 2012; Macosko et al., 2009; Narayan et al., 2016; Pungaliya et al., 2009; Srinivasan et al., 2008).

Oxygen and Carbon Dioxide Sensing Neurons

Water soluble and volatile compounds are not the only stimuli sensed by olfactory neurons. Oxygen and carbon dioxide levels are sensed by a small subset of sensory neurons not included in the classical set of amphid olfactory neurons (Carrillo & Hallem, 2015).

The ciliated BAG neurons sense CO2 levels (Bretscher, Busch, & de Bono, 2008; Bretscher et al., 2011; Busch et al., 2012; Zimmer et al., 2009), and drive avoidance behaviors in situations in which there are elevated levels of the gas (Hallem & Sternberg, 2008). With help from ASE, and the canonical thermosensory neuron AFD, BAG is the main sensor of CO2 in the nematode (Bretscher et al., 2011).

The URX neurons, which are not ciliated, also develop in the amphid region and contain dendrites that are exposed to the pseudocoelomic fluid (Styer, Singh, Macosko, Steele, Bargmann, & Aballay, 2008). In combination with the AQR and PQR neurons, URX helps C. elegans aerotax towards ideal O2 levels (Chang, Chronis, Karow, Marletta, & Bargmann, 2006; Gray et al., 2004).

Other Amphid Olfactory Neurons

The IL2 neurons have cilia exposed to the external environment, but what they sense and what they communicate is unknown. In dauer larvae, IL2 cilia regulate a dispersal behavior, though it remains unknown how the cilia are activated to induce this output (Lee et al., 2012; Schroeder et al., 2013).

The Nature of Phasmid Olfactory Neurons

Although the roles of PHA and PHB have been determined (Barrios, Ghosh, Fang, Emmons, & Barr, 2012; Hilliard et al., 2002), what these neurons are sensing remains largely unknown. Many of the sex-specific neurons present in males are located in the tail, increasing the overall number of neurons exposed to the environment, increasing phasmid olfactory capability (White et al., 2007). Three pairs of ray neurons also contribute to an increase in the number of serotonergic olfactory neurons (Jafari, Xie, Kullyev, Liang, & Sze, 2011; White et al., 2007). It has not been yet determined exactly what these olfactory neurons sense.

The distal tips of the copulatory spicules contain the exposed cilia of the SPD and SPV neurons. Given the spicules’ role during mating, it is proposed that these neurons sense vulval pheromones to manage release of sperm into the correct location (LeBoeuf, Correa, Jee, & García, 2014; Liu & Sternberg, 1995; Schindelman, Whittaker, Thum, Gharib, & Sternberg, 2006). To date, there has been no identification of these proposed vulval pheromones.

Assays Used in the Study of C. elegans Olfaction

Behavioral Assays

The most ethologically relevant output of sensation to study is behavior. With obtainable results on short timescales, the effect of a stimulus on C. elegans behavior can be studied both in fine detail and in a high-throughput manner (Ghosh, Nitabach, Zhang, & Harris, 2017; Maruyama, 2016). The simplest of observable behaviors are attraction and repulsion. In fact, this chemotaxis was studied in C. elegans even before Brenner’s initial push for the use of the nematode as a standard model system (Ward, 1973).

The eutelic and transparent nature of C. elegans allows for reliable identification and laser ablation of specific cells. In these ablations, a laser is focused and pulsed at the nucleus of a neuron of interest, effectively killing the cell (Fang-Yen, Gabel, Samuel, Bargmann, & Avery, 2012). Nematodes undergo this laser surgery as larvae, and are then allowed to develop in the absence of the ablated cell, after which, the animal can be assayed, and changes in outputs observed.

In 1973, Ward et al. showed that C. elegans are able to respond to attractants by moving up a concentration gradient, accumulating in the area of the cue, and then habituating to the cue (Figure 2A) (Ward, 1973). In the now-canonical population chemotaxis assay developed by the Bargmann lab, the number of C. elegans at selected time points within specified attractant areas are counted to generate a chemotaxis index (Figure 2A) (Bargmann et al., 1993). Changes in this index amongst mutants or laser-ablated animals have helped identify neurons and neuromodulators that play roles in attractive olfaction.

Caenorhabditis elegans OlfactionClick to view larger

Figure 2. Behavioral Assays Used in Understanding Olfaction. (A) Attractive Gradient Assay: C. elegans are placed in a drop of an attractant chemical (shaded area). As the chemical diffuses, creating a gradient, animals reside in the ideal concentration of the cue. (B) Chemotaxis Assay: C. elegans are placed in the center of plate, with two drops of attractant at either end of the plate (shaded area). The number of animals within this region are scored over time and divided by the total number of worms on the plate to generate an Attraction Index. (C) Drop Avoidance Assay: A drop of aversive chemical is placed on the tail of a forward moving nematode. Capillary action draws the solution up to the head of the animal where it is sensed and initiates a reversal, and ultimately, a change of direction. The number of drops that cause animals to avoid is divided by the total number of drops applied to generate an Avoidance Index. (D) Population Level Avoidance Assay: C. elegans are placed on one side of an aversive chemical barrier (darker region). In order to reach a volatile attractant (shaded area), they must cross the aversive barrier. (E) Aerotaxis Assay: An agar field containing C. elegans is placed in a chamber, which is connected to inputs for oxygen and nitrogen. The inflow of gases is controlled to create an oxygen gradient across the chamber. Nematodes can aerotax to the region with the preferred oxygen content.

C. elegans utilize two independent mechanisms to chemotax towards ideal concentrations of odorants: pirouettes (klinokinesis) and weathervaning (klinotaxis). Characterization of changes in the uses of these methods of chemotaxis in mutant animals has helped to identify the underpinnings of how C. elegans reach an ideal environment (Chalasani et al., 2007; Lockery, 2011; Luo, Gabel, Ha, Zhang, & Samuel, 2008). The pirouette mechanism is comprised of “bouts of sharp turns,” usually including a reversal and/or ω‎ turn (Iino & Yoshida, 2009). These sharp turns are initiated upon a sensation of a decrease in the concentration of an attractant, such as salt (Pierce-Shimomura et al., 1999), although they also utilize this method in response to volatile odorants (Chalasani et al., 2007).

In contrast to pirouettes, C. elegans also employ a more gradual curving towards an attractant, termed weathervaning (Iino & Yoshida, 2009). This behavior was first proposed in 1973, but there was no further evidence supporting this behavior until over three decades later (Iino & Yoshida, 2009; Ward, 1973). To generate this curve, the amphid sensory neurons were found to sense changes in odorant concentrations at each apex of the head swing during the sinusoidal movement of the worm (Izquierdo & Lockery, 2010; Kato, Xu, Cho, Abbott, & Bargmann, 2014; Larsch, Flavell, Liu, Gordus, Albrecht, & Bargmann, 2015). Iino and Yoshida proposed that slightly larger head swings are then generated in the direction of an attractant, generating the curving motion (Iino & Yoshida, 2009).

Avoidance assays have also been established to aid in the understanding of how C. elegans senses chemical cues in its environment. A drop assay was developed by Hilliard et al., which exposes a forward moving animal to a drop of a soluble chemical cue (Figure 2C) (Hilliard et al., 2002; Hilliard, Bergamasco, Arbucci, Plasterk, & Bazzicalupo, 2004). An avoidance index is then calculated by determining the fraction of worms that initiate reversals upon exposure to the cue. There is also a population-level aversion assay in which worms are placed on one side of a line of aversive chemical. A volatile attractant is then placed on the opposite side of the aversive chemical line (Figure 2D). An aversion index is calculated by dividing the number of worms that cross the boundary in one hour by the total number of worms on the plate (Hart, 2006).

C. elegans senses oxygen and carbon dioxide levels through olfactory neurons. Aerotaxis studies involve specialized devices, in which a gas gradient can be generated by pumping opposing gases into a chamber housing an agar arena (Figure 2E) (Gray et al., 2004). A score is given based upon where animals are found within the gradient, and mutants can easily be determined by changes in this distribution.

Assays allowing for the study of more complex behaviors (such as aggregation, foraging, and dispersal) have also been developed, as advancements in worm tracking and technological capabilities have been made (Greene et al., 2016b; Milward, Busch, Murphy, de Bono, & Olofsson, 2011; Sambongi et al., 1999). With greater software and coding capabilities, worm tracking is now common, and even a marketable process. Many labs have developed their own tracking software (Albrecht & Bargmann, 2011; Buckingham, Partridge, & Sattelle, 2014; Chalasani et al., 2007; Cronin, Feng, & Schafer, 2006; Faumont et al., 2011; Feng, Cronin, Wittig, Sternberg, & Schafer, 2004; Kawano, Po, Gao, Leung, Ryu, & Zhen, 2011; Leifer, Fang-Yen, Gershow, Alkema, & Samuel, 2011; Stirman, Crane, Husson, Gottschalk, & Lu, 2012; Swierczek, Giles, Rankin, & Kerr, 2011; Tsechpenakis, Bianchi, Metaxas, & Driscoll, 2008; Tsibidis & Tavernarakis, 2007), and companies exist yhat offer user friendly software for purchase, such as WormLab—developed by MBF Bioscience. There are options available for the tracking of single worms or populations of worms, either on agar plates, in suspension, and more. Analyses with these trackers include parameters such as speed, turning frequencies, and bending angles, with multi-worm trackers even being able to distinguish between two worms that come into contact with each other. In-depth descriptions of available worm-trackers, with side-by-side comparisons are available (Husson, Costa, Schmitt, & Gottschalk, 2012).

Developmental Assays

As a result of processing information about their immediate environment and their developmental state, nematodes emit small molecules to communicate information to conspecifics. To do so, they utilize a class of biogenic, small-molecule pheromones, termed ascarosides. To date, ascarosides have been found to mediate entry into dauer, an alternative and environmentally persistent developmental state (the collective mix of ascarosides that signal dauer formation are termed “daumone” (Golden & Riddle, 1982)), foraging behavior, avoidance behavior, and mate recognition (Butcher, 2017a, 2017b; Chute & Srinivasan, 2014; Ludewig & Schroeder, 2013).

Caenorhabditis elegans OlfactionClick to view larger

Figure 3. Dauer Development Assays. (A) Dauer Formation Assay: A few adult C. elegans are allowed to lay eggs on a plate before being removed. These eggs are allowed to hatch and larvae grow under dauer promoting conditions (pheromone, temperature, etc.). The proportion of worms in the dauer state are determined in two ways: By visual scoring of worms that maintain dauer appearance and morphology, or by applying a solution of 1% SDS to the population, and scoring the animals still alive after as being in the dauer state. The number of animals that enter dauer are divided by the total number of animals on the plate to determine Percent Dauer Formation. (B) Dauer Recovery Assay: C. elegans are grown in liquid culture until entering dauer. Non-dauer animals are removed from the population, and dauer animals are plated. After three days, the number of nematodes that have exited dauer to resume normal development are scored and divided by the total number of plated animals to determine Percent Dauer Recovery.

In dauer-formation assays (Figure 3A), worms are allowed to develop on plates containing chemicals that may induce dauer development. Since dauer worms survive harsh conditions, the worms surviving after treatment can be scored and compared to the pre-treatment population to calculate the fraction of induced dauer larvae (Jeong et al., 2005). Alternatively, dauers can also be identified “based on size, shape, and lack of pharyngeal pumping” (Butcher, Fujita, Schroeder, & Clardy, 2007).

In dauer recovery assays (Figure 3B), worms are cultured in liquid media and allowed to form dauers. Non-dauer worms are removed from the culture, and the dauers are then placed on an agar plate containing chemicals that may induce exit from dauer. After two days, dauer and non-dauer worms are visually scored (Butcher et al., 2007).

Imaging of Neural Activity in Olfactory Neurons

With the development of genetically encoded calcium indicators (GECIs), visualization of neuronal activity in vivo has become possible (Nakai, Ohkura, & Imoto, 2001). These proteins allow for real-time readout of intra-cellular calcium levels. Original GECIs utilized Green Fluorescent Protein (GFP) as their fluorescent protein in the fusion (generating the GCaMP family of GECIs), although recent developments have incorporated red-shifted chromophores, such as RFP (RCaMPs) (Akerboom et al., 2012, 2009; Sun et al., 2013).

By expressing GECIs in olfactory neurons, and exposing C. elegans to an olfactory stimulus, the depolarization and hyperpolarization of sensory neurons can not only be visualized, but measured (Hilliard et al., 2005). Microfluidic devices allow for the containment of nematodes in a manner that allows for capture of the fluorescent dynamics of GECIs upon exposure to olfactory cues (Reilly, Lawler, Albrecht, & Srinivasan, 2017). Animals can be assayed while “trapped” in devices, or allowed to roam in “arenas” (Chronis, Zimmer, & Bargmann, 2007; Lagoy & Albrecht, 2015; Larsch, Ventimiglia, Bargmann, & Albrecht, 2013).

Mechanisms of Olfaction

Molecular and Cellular Mechanisms

G Protein-Coupled Receptors

The C. elegans genome encodes approximately 1,200 G protein-coupled receptors (GPCRs), over half of which are proposed to be chemosensory in nature. Only a handful of receptors have been directly linked to sensation of specific cues (Bastiani & Mendel, 2006). This has likely arisen due to the redundancy of olfactory neurons in the nematode. With only approximately three dozen neurons participating in olfaction, there is a remarkable amount of GPCR co-expression within olfactory neurons. As such, the redundant nature of these receptors has caused genetic screens to be ineffective in linking receptors to biological roles. In the two decades since the finding that ODR-10 senses diacetyl (Sengupta, Chou, & Bargmann, 1996), only a handful of GCPRs have been linked to specific targets: SRBC-64 and -66, DAF-37 and -38, SRG-36 and -37 sense dauer inducing ascarosides (Kim et al., 2009; McGrath et al., 2011; Park et al., 2012), SRX-43 and SRX-44 sense icas#9 to control foraging behaviors (Greene et al., 2016b; Greene, Dobosiewicz, Butcher, McGrath, & Bargmann, 2016a), while SRI-14 senses high concentrations of diacetyl alongside the low-concentration sensing ODR-10 (Sengupta et al., 1996; Taniguchi, Uozumi, Kiriyama, Kamizaki, & Hirotsu, 2014).

GPCRs can bind ligands present in the worm’s environment, and, in conjunction with associated G proteins, propagate signaling cascades. The remaining GPCRs, which are not expected to be chemosensory in nature, likely play other roles in neurotransmission (Bargmann, 1998; Bargmann, 2006a).

The original method of determining GPCR expression patterns was through the generation of transcriptional fusions, wherein the promoter of the GPCR gene of interest drives expression of GFP. These first tests showed that GPCRs in C. elegans are expressed in either single or multiple neuronal classes (Taniguchi et al., 2014; Troemel, Chou, Dwyer, Colbert, & Bargmann, 1995). As expected, it was confirmed that GCPRs, which function as chemoreceptors in these olfactory neurons, are localized to the cilia of these neurons (Dwyer, Troemel, Sengupta, & Bargmann, 1998; Troemel et al., 1995).

This has continued to hold true with the advent of ascaroside receptor discoveries, using both fusions and immunostaining (Greene et al., 2016b; Greene, Dobosiewicz, et al., 2016a; Kim et al., 2009; McGrath et al., 2011; Park et al., 2012). GPCRs have been shown to function as heterodimers, with two unique receptors involved in the sensation of each cue. In this sensation method, some receptors function as primary binders of specific targets, while the other member may function as a partner for multiple primary receptors in the sensation of stimuli (Park et al., 2012). Despite the relatedness of ascarosides in terms of core structure, the receptors identified to date are widespread across the GPCR families present in C. elegans; genes have been discovered to be responsible for ascaroside sensation in the srbc, srg, srw, and srx families (Greene et al., 2016b; Greene, Dobosiewicz, et al., 2016a; Kim et al., 2009; McGrath et al., 2011; Park et al., 2012).

Stimuli sensed by GPCRs need not be as structurally complex as ascaroside pheromones, however. In fact, the ketone diacetyl was found to be sensed by two different GPCRs, dependent on the concentration of the stimuli: ODR-10 for low concentrations and SRI-14 for high concentrations. Diacetyl elicits an attractive response when sensed by the AWA neurons and bound to ODR-10 (Sengupta et al., 1996). However, high concentrations are sensed by SRI-14, expressed in the nociceptive ASH neuron, and produce aversive behaviors (Taniguchi et al., 2014).

G protein-coupled receptors would not function properly without the related G proteins and downstream intracellular signaling cascades. Within the C. elegans genome, there are twenty Gα‎, two Gβ‎, and two Gγ‎ subunits (Jansen, Thijssen, Werner, van derHorst, Hazendonk, & Plasterk, 1999). Due to the presence of hundreds of GPCRs in the olfactory neurons, it is understandable that multiple subunits may be expressed in individual neurons. Because of this, the roles of individual Gα‎ subunits vary in importance (Bastiani & Mendel, 2006).

Downstream Ion Channels Involved in Olfaction

Downstream of GPCRs and their related G-protein complexes(Cuppen, van der Linden, Jansen, & Plasterk, 2003), C. elegans use two canonical intracellular signaling cascades to elicit depolarizations (Bargmann, 2006a): cyclic nucleotide-gated channels, and transient receptor potential (TRP) channels, which are utilized in two separate subsets of olfactory neurons (Table 1) (Coburn & Bargmann, 1996; Colbert & Bargmann, 1997; Komatsu, Mori, Rhee, Akaike, & Ohshima, 1996; Tobin et al., 2002).

Olfactory neurons utilizing cyclic nucleotide-gated channels to initiate depolarizations express TAX-2 and TAX-4 as a heterodimer in the cilia of the neuron (Table 1). This channel is downstream of receptor-type guanylate cyclases, which induce influxes of cGMP, and lead to a flood of sodium or calcium into the neuron (Ortiz et al., 2006). These channels are also functional in the oxygen and carbon dioxide sensing neurons, where they are activated by soluble guanylate cyclases (Bretscher et al., 2011; Carrillo, Guillermin, Rengarajan, Okubo, & Hallem, 2013; Cheung, Arellano-Carbajal, Rybicki, & de Bono, 2004; Gray et al., 2004; Hallem et al., 2011; Hallem & Sternberg, 2008; Yu et al., 1997).

While receptor-type guanylate cyclases (rGCs) are crucial toward generating cGMP flux downstream of GCPRs in TAX-2/TAX-4 neurons (Ortiz et al., 2006), there are a few instances wherein they act as chemoreceptors themselves. For example, GCY-9 functions to directly bind CO2 in the BAG neurons, while GCY-14 serves as a pH indicator in the left ASE neuron (Murayama & Maruyama, 2013; Murayama, Takayama, Fujiwara, & Maruyama, 2013; Smith, Martinez-Velazquez, & Ringstad, 2013). The large, nematode specific expansion of rGCs (Fitzpatrick, O’Halloran, & Burnell, 2006) suggests that there are more playing as yet unknown but direct roles in chemosensation.

Conversely, olfactory neurons can utilize heterodimeric TRP channels to provoke depolarizations in the sensory cilia. OSM-9/OCR-2 form a heterodimeric channel in these TRP-utilizing cilia (Table 1). In Table 1, there is no overlap between cells expressing both tax-2 and tax-4, and cells expressing both osm-9 and ocr-2, but there is some overlap between tax-2, tax-4 and osm-9 alone. In this signaling cascade, G proteins regulate lipid metabolism which, in turn, activate the TRP channel, leading to the desired depolarization (Kahn-Kirby et al., 2004).

Table 1. Olfactory Neurons and Cilium Structures




Olfactory Cues Sensed

Intracellular Components





osm-9, ocr-2










tax-2, tax-4, osm-9


Single Rod



tax-2, tax-4, osm-9


Single Rod

Dauer Control


tax-2, tax-4, osm-9


Single Rod


Aversive Stimuli

osm-9, ocr-2


Single Rod

Dauer Control/Foraging

Daumone, Icas#9

tax-2, tax-4, osm-9


Single Rod

Dauer Control


tax-2, tax-4, osm-9


Single Rod


Icas, ascr#3

tax-2, tax-4, osm-9


Double Rods

Dauer Control


osm-9, ocr-2


Double Rods


Water solubes, ascr#3, icas#9

osm-9, ocr-2





tax-2, tax-4




















Single Rod



osm-9, ocr-2


Male Attraction

ascr#3, ascr#8






Vulval Pheromones

Circuit-Level Analysis

Integration of Olfactory Inputs

Unlike other known olfactory systems studied in neuroscience, such as that of the mouse, C. elegans olfaction relies on only a small subset of neurons, requiring co-expression of receptors within individual neurons. This is in juxtaposition to the one-neuron-one-receptor system observed in mammals (Bargmann, 2006b; Chess et al., 1994; Serizawa et al., 2003). The majority of this limited number of olfactory neurons converges onto a small subset of interneurons, largely responsible for integration of sensation. Without this integration, proper olfactory responses would be impossible. This layer of neuronal networks is critical when investigating olfactory mechanisms.

Caenorhabditis elegans OlfactionClick to view larger

Figure 4. Network Regulation of Olfaction. The first two layers of olfactory sensation integration: The majority of amphid olfactory neurons synapse onto at least one of the “First Layer Interneurons.” These recurrently connected interneurons then synapse and transmit information onto motor neurons (not shown) and the Command Interneurons: AVB, PVC, AVA, AVD, and AVE. The Command Interneurons control the forward and reverse locomotory circuitries. Black arrows denote physical, not functional connections.

With the exception of ASJ and URX, all of the amphid olfactory neurons synapse onto at least one member of the “first-layer interneurons”: AIA, AIB, AIY, and AIZ (Ward et al., 1975; Ware et al., 1975; White, Southgate, Thomson, et al., 1986). It is these neurons that are speculated to integrate the information from the upstream neurons, and initiate proper behavioral or developmental responses. These “first-layer interneurons” are responsible for summing and integrating sensory input, before passing the information downstream onto either motor neurons or the command interneurons which control forward and reverse locomotory circuitries (Figure 4) (Chalfie et al., 1985). The complex and recurrent connectivity among these four neurons allows for fine-tuning and integration of sensory input prior to initiating downstream signaling.

Caenorhabditis elegans OlfactionClick to view larger

Figure 5. Functional Circuits Involved in C. elegans’ Olfaction. (A) The RMG interneuron acts as a hub for olfactory input from four chemosensory neurons, the oxygen-carbon dioxide sensing URX neuron, and the IL2 neurons. Integrated input is them passed downstream onto the command interneuron. (B) The AWA and ASH neurons synapses onto the RIM interneuron to control initiation of forward and backwards locomotion. (C) Icas#9, an indolated ascaroside is sensed by three chemosensory neurons and two receptors to control roaming behavior. (D) The AWC-AIA-AIY circuit functions in an “Odor-OFF” fashion. When odor is present (left), the AWC neuron inhibits AIY, and is inhibited by AIA. When the odor is removed (right), the AWC neurons inhibit the inhibitory AIY, and activate AIB. The removal of AIY inhibition contributes to AIY activity as well. (E) Nociceptive avoidance is fine-tuned by ASI and ASH. A “reciprocal inhibition” is observed, with ASI activating ADF inhibition of ASH. Meanwhile, ASH activates RIC inhibition of ASI.

Interneurons such as RMG have are thought to serve as hubs for integration of olfactory input. RMG receives inputs from six amphid olfactory neurons, and modulates aggregation and other social behaviors (Figure 5A) (Macosko et al., 2009). Likewise, neurons such as AWA and ASH modulate the RIM interneuron to initiate backward and inhibit forward movements (Figure 5B) (Ghosh et al., 2016). While they are not directly responsible for sensing olfactory stimuli in the environment, it is critical to include these synaptic partners in studies of olfaction, as the regulatory roles of these neurons contribute immensely to the complexity of olfactory circuits.

A Functional Foraging Circuit

Icas#9, a weak inducer of dauer, promotes foraging behavior. Work by Greene, Dobosiewicz, et al. (2016a) unveiled a circuit that senses icas#9 through three olfactory neurons: ASI, ASJ, and ADL (Figure 5C) (Greene et al., 2016a, 2016b). ASI promotes roaming when sensing icas#9 in opposition to ADL (Figure 5C). ASJ senses icas#9 through a different receptor than that present in ASI, yet drives the same behavior (Greene, Dobosiewicz, et al., 2016a). ADL, utilizing the same receptor found in ASJ, represses the foraging behavior promoted by ASI and ASJ.

Responding to Volatile Odors

The AWC-AIA-AIY circuit is a complex circuit that tunes responses to volatile odors. AWC functions in an “odor-off” fashion; when odors are removed, AWC inhibition of the downstream interneuron AIY is relieved (Figure 5D) (Chalasani et al., 2007). Upon removal of the odor, glutamatergic inhibition of AIY resumes. In turn, the response of AWC to these odors is modulated by the AIA interneuron (Figure 5D) (Chalasani et al., 2010).

Nociception and Avoidance

ASI and ASH both sense copper to drive avoidance behaviors. The “reciprocal inhibition” observed in these neurons tunes this avoidance behavior. ASI activates ADF, which, in turn, functions to inhibit ASH (Figure 5E). ASH, meanwhile, activates the RIC interneuron, which then works to inhibit ASI signaling (Guo et al., 2015).

ASH has also been shown to sense 1-octanol and to activate the AIB interneuron and the AVA motor neuron, which drives backwards locomotion (Figure 4). AWC also contributes towards the rate of spontaneous reversals by activating the AIB interneuron (Figure 5D). However, AWC is inhibited by both 1-octanol and food, preventing it from activating the AIB interneuron. The removal of food alters internal 5-HT levels (Harris et al., 2011, 2009), which in turn also inhibits the AIB interneuron. This cumulative inhibition of the AIB interneuron results in inhibition of reversals and, upon completion of a reversal, re-initiation of forward movement (Summers, Layne, Ortega, Harris, Bamber, & Komuniecki, 2015).

Male Ascaroside Sensation

Many functional connectomes remain to be understood. Male-specific CEM neurons sense ascarosides #3 and #8 to attract males to receptive hermaphrodites (Srinivasan et al., 2008). However, while the class of neurons is required for proper response to these cues, not all four CEM neurons elicit the same changes in calcium levels (Narayan et al., 2016). A subset of CEM neurons depolarize, others hyperpolarize, and others seem to not respond at all. These subsets vary from animal to animal, suggesting that the cumulative output of the CEM neurons is more important than individual CEM neuron response. Given that the four neurons respond differently, it is likely that there is cross-talk occurring between the four neurons, possibly similar to the ASI-ASH “reciprocal inhibition.”

Future Directions

Given the complexity of the olfactory system in C. elegans, genetic and molecular approaches need to be complemented with functional circuit studies. Identifying receptors responsible for the sensation of olfactory stimuli coupled with elucidating the functional connections of the nematode, will improve the understanding of the olfactory connectome.

Functional Connectomics

The initial wave of C. elegans neuronal circuit dissections focused heavily on understanding the physical connectome. Many studies have since shown that neurons communicate heavily outside of synaptic connections suggesting a much bigger picture of how neural circuits function (Bargmann, 2012; Bentley et al., 2016; Rajendran et al., 2014). Canonically, only neuropeptide signals have been known to function in a long-range, extra-synaptic fashion (Leinwand & Chalasani, 2014; Mills et al., 2012). However, recent studies have shown that the monoaminergic nervous system communicates in context-dependent and extra-synaptic manners (Hardaway et al., 2015; Komuniecki, Hapiak, Harris, & Bamber, 2014). Also, extracellular vesicles are playing increasingly important roles in extra-synaptic cell-to-cell signaling and circuit regulation (Rajendran et al., 2014; Wang et al., 2015).

Increasingly, the physiological state or age of C. elegans is being found to influence the functional aspects of the connectome (Chao, Komatsu, Fukuto, Dionne, & Hart, 2004; Ghosh et al., 2016; Leinwand, Yang, Bazopoulou, Chronis, Srinivasan, & Chalasani, 2015). To completely elucidate the functional connectome, factors such as age, sex, physiological state, and life history, must be taken into account and assayed independently. A likely outcome of these studies is that many overlapping functional connectomes will be generated providing deeper insights into the role of neuroregulators.

Receptor Identification

To fully understand olfaction, the mechanistic underpinnings of the process must be revealed. To do this, the receptors that physically sense and bind these cues need to be identified. Due to the inherent complexity that arises from co-expression (Hostettler, Grundy, Kaser-Pebernard, Wicky, Schafer, & Glauser, 2017; Troemel et al., 1995) and heterodimerization (Park et al., 2012) of GPCRs, fully identifying receptor complexes will require an immense amount of investigation. With multiple receptors being required for sensing different concentrations of the same cue, the mechanisms of olfaction have become even more complex than initially believed (Taniguchi et al., 2014).

There are many approaches currently available for identifying receptors, ranging from RNAi knockdowns to quantitative trait locus mapping for identification of chromosomal regions responsible for proper olfactory response (Greene et al., 2016b; Greene, Dobosiewicz, et al., 2016a; Taniguchi et al., 2014). The development of biochemical probes increases the scope by which identifying olfactory receptors can be achieved. These probes are modified cues, containing functional groups that can be photo-activated to covalently bind to the receptor, allowing for direct confirmation of binding. These probes can also be constructed to allow for FRET analysis following covalent binding, removing the need for purification to identify the receptor (Park et al., 2012). Recently, genomic and transcriptomic investigations have been used to match receptor expression profiles with ligand targets (Greene et al., 2016b; Greene, Dobosiewicz, et al., 2016a; Greer et al., 2016).

While much has been discovered about molecular and circuit mechanisms of olfaction, much remains to be understood. C. elegans’ experimental amenability and defined connectome offers an ideal tool for addressing these gaps by providing a “systems-level” analysis of olfaction. Increases in neuronal imaging capabilities allows for unrestrained live imaging with minimal alterations being made to the natural state of the animal. This allows for circuit-level read-outs to be deciphered. Recent studies have focused on understanding global control of olfaction (Kato et al., 2015; Nguyen et al., 2016; Venkatachalam et al., 2016). Future work will likely focus on uncovering the regulation of olfactory circuits that integrate olfactory stimuli along with targeted identification of individual GPCRs necessary for olfactory sensation.


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