An Aversive Response to Osmotic Upshift in Caenorhabditis elegans

C. elegans aversively responds to osmotic upshifts

The osmolarity of the standard culturing medium for C. elegans is ∼150 mOsm. Exposure to solutions of ≥1 Osm osmolarity induces shrinkage of the worm body and immobilization in a few minutes. A long-term exposure to similar osmolarity induces adaption and resistance due to increased accumulation of glycerol in the body (Lamitina et al., 2004; Solomon et al., 2004). C. elegans acutely avoids solutions of high osmolarity (≥1 Osm) by generating reversals within a few seconds of a direct encounter, and the avoidance requires the C. elegans homolog of the mammalian TRPV channels in the sensory neuron ASH. In contrast, on direct contact C. elegans does not generate a similar reversing response to hyperosmotic stimuli ranging from 200 mOsm to 1 Osm (Colbert et al., 1997; Liedtke et al., 2003; Hilliard et al., 2004, 2005). Because solutions that have osmolarity within this range also cause shrinkage of the body within a few minutes of exposure and induce a long-term adaption (Liedtke et al., 2003; Solomon et al., 2004), we examined whether C. elegans displayed a different behavioral response to hyperosmotic stimuli of this range. We used an automated droplet assay that allowed the nematode to swim in 6 µl drops of a solution and measured the rate of reversals and large body bends (Fig. 1A; Luo et al., 2008; Ha et al., 2010). Previously, we have shown that during swimming C. elegans generates continuous “C-shaped” body swings that are occasionally interrupted by turns that include reversals and large body bends and are followed by reorienting movements. Attractive stimuli suppress the turning rate, while removal of attractive stimuli or exposure to repulsive cues increases it (Bargmann et al., 1993; Clark et al., 2007; Luo et al., 2008; Srivastava et al., 2009; Ha et al., 2010). Therefore, an increase in the number of turns within a given amount of time represents an aversive response (Fig. 1A). We found that the worms cultivated on the standard NGM, which had an osmolarity of ∼150 mOsm, displayed a stable turning rate in a glycerol solution of 150 mOsm. In comparison, when exposed to a glycerol solution of 400 mOsm these animals gradually and significantly increased their turning rate and reached the highest turning rate after 5 min (Fig. 1B). It is conceivable that the worms generated the behavioral response as part of the response to dehydration. Consistently, we found that the worms that were cultivated under the condition of 150 mOsm shrank their bodies to (mean ± SEM) 97.5% ± 1.1% or 77.6% ± 2.1% (n = 7 each), respectively, of the area that their bodies occupied on a flat image after swimming in a glycerol solution of 200 or 400 mOsm for 8 min (Materials and Methods). In addition, we measured how fast the worms thrashed their bodies in the 150 and 400 mOsm solutions (Materials and Methods) and found that they thrashed significantly less in the 400 mOsm glycerol solution (Fig. 1C), suggesting that the increase in the turning rate does not result from a general increase in locomotion. We also examined the behavioral response to glycerol solutions of higher osmolarity, 600 and 800 mOsm, and found that both stimuli increased the turning rate (Fig. 1D,E) within a few minutes after exposure. To address whether the aversive response was induced by an increase in osmolarity or by an increase in the concentration of glycerol, we examined the animals that were exposed to a NaCl solution or a sorbitol solution of similar osmolarity. We found that exposure to either a NaCl solution of 400 mOsm or a sorbitol solution of 400 mOsm similarly induced an increase in the turning rate (Fig. 1F,G). Together, these results indicate that with a continuous exposure C. elegans aversively responds to osmotic upshifts by gradually increasing reorienting movements. For the rest of the study, we used glycerol solutions to characterize the behavioral response to osmotic upshifts.

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Figure 1.

C. elegans generates an aversive response to mild osmotic upshifts. A, A sample image of the droplet assay (Materials and Methods). Each 6 µl droplet of buffer contains one worm, and the arrow denotes a large body bend. B, WT animals that are cultivated under the standard osmotic condition (∼150 mOsm) display a stable turning rate in a glycerol solution of 150 mOsm but gradually and significantly increase the turning rate in a hyperosmotic glycerol solution of 400 mOsm. n = 55 each. C, WT animals that are cultivated under the standard osmotic conditions (∼150 mOsm) display a significantly lower thrashing speed in a hyperosmotic glycerol solutions of 400 mOsm than in a glycerol solution of 150 mOsm. n = 55 each. D, E, WT animals that are cultivated under the standard osmotic condition (∼150 mOsm) also gradually and significantly increase the turning rate in the hyperosmotic glycerol solutions of 600 mOsm (D; n = 27 each) or 800 mOsm (E; n = 12 each). Note that in E the turning rate starts to drop after 3 min in the 800 mOsm solution, due to decreased mobility of the animals possibly caused by server dehydration. Therefore, the statistical test is performed on the turning rates in the 4th minute in E. F, G, WT animals cultivated under the standard osmotic condition (∼150 mOsm) display an increased turning rate in a 400 mOsm hyperosmotic solution that is balanced by either NaCl (F; n = 24 each) or sorbitol (G; n = 24 each) in comparison with the turning rate in the 150 mOsm solution. For B–G, the turning rates in solutions of different osmolarities in the 5th minute (in the 4th minute in E) are compared using the Student’s t test. H, WT animals that are cultivated under the standard osmotic condition (∼150 mOsm) display a stable turning rate on a NGM plate of 150 mOsm but gradually and significantly increase the turning rate on a NGM plate of 400 mOsm. The osmolarity of the plates is balanced by sorbitol. The turning rates on plates of different osmolarity are compared using the Student’s t test. n = 7 animals each. For all, ***p < 0.001, **p < 0.01. Values are reported as the mean ± SEM. WT, wild type.

Next, we asked whether C. elegans responded to osmotic upshifts on a solid substrate in a manner similar to that in a liquid environment. We measured reversals and turns in animals that freely moved on a NGM agar plate that had osmolarity of ∼150 or 400 mOsm. We found that the animals on the 400 mOsm NGM plate generated significantly more turns and reversals than the animals on the 150 mOsm NGM plate within a 5 min exposure (Fig. 1H). Note that the rate of turns and reversals during swimming is higher than that during crawling, which is consistent with the faster locomotory dynamics previously characterized for swimming in comparison with those of crawling (Pierce-Shimomura et al., 1999; Gray et al., 2005; Korta et al., 2007; Srivastava et al., 2009; Ha et al., 2010). It has been shown that when crawling forward on a solid substrate, C. elegans generates sinusoidal body waves that are randomly interrupted by reorienting movements, mainly reversals and turns; attractive and aversive environmental conditions suppress or increase the reorienting movements, respectively (Pierce-Shimomura et al., 1999). Together, these results indicate that C. elegans responds to the upshift of osmolarity by gradually increasing reorienting maneuvers during either swimming or crawling. This navigational response takes a few minutes to occur, which is different from an acute escape-like avoidance behavior.

Because maintaining a stable intracellular osmolarity is a homeostatic drive, we examined whether the aversive response to osmotic upshifts is associated with homeostatic responses. Because homeostatic responses are evoked by deviations from a baseline condition (Craig, 2003; Critchley et al., 2004), we tested animals with different internal osmolarity. Previous studies show that C. elegans adapts to prolonged exposure to hyperosmotic conditions by increasing the intracellular osmolarity through increased synthesis of glycerol (Lamitina et al., 2004). Thus, we predicted that adapting the animals to an external osmolarity of 400 mOsm would prevent them from increasing their turning rate in response to exposure to an osmolarity of 400 mOsm. Consistently, we found that animals that were cultivated on the NGM plate of 400 mOsm displayed similar turning rates when exposed to 400 or 200 mOsm (Fig. 2A). In addition, these results also show that an osmotic downshift from 400 to 200 mOsm did not induce a significant change in the turning rate. We speculate that a potential behavioral response to osmotic downshifts might be displayed with other locomotory parameters or might occur too slowly to be detected within a few minutes of exposure. We also tested osr-1(rm1) and dpy-2(e8) mutant animals. Previous studies have shown that a prolonged exposure to hyperosmotic conditions (1 Osm) decreases the body volume and reduces mobility in C. elegans due to the loss of water. Both the osr-1(rm1) and dpy-2(e8) mutants harbored mutations that generated resistance to the water loss induced by hyperosmotic conditions (Solomon et al., 2004; Wheeler and Thomas, 2006). We found that neither the osr-1(rm1) mutants nor the dpy-2(e8) mutants increased their turning rates in response to the osmotic upshift from 150 to 400 mOsm (Fig. 2B,C). Previous studies show that the osr-1(rm1) mutant animals generate normal avoidance to nose touch or the repellent 1-octanol (Solomon et al., 2004). We found that the dpy-2(e8) mutant animals normally avoided (WT, 85.3% ± 2.0% avoidance; dpy-2, 86.7% ± 1.9% avoidance, n = 30 animals each; p = 0.625, Student’s t test, mean ± SEM) gentle touches to the anterior part of the body (Chalfie et al., 1985). Therefore, these two mutants are not generally defective in generating avoidance response. Together, these results suggest that the aversive response to osmotic upshifts is part of the physiologic response to the hyperosmotic stress.

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Figure 2.

The increase in the turning rate is part of the physiologic response to hyperosmotic stresses. A, Wild-type animals cultivated under the standard NGM plates of 150 mOsm display a higher turning rate in a hyperosmotic solution of 400 mOsm (NGM_400 mOsm, n = 32) than in a 200 mOsm solution (NGM_200 mOsm, n = 28); but animals that are cultivated on the plates of 400 mOsm display a turning rate in the solution of 400 mOsm (adapted_400 mOsm, n = 32) that is similar to that in the solution of 200 mOsm (adapted_200 mOsm, n = 32). Significant interaction between cultivating conditions and osmolarity is tested by two-way ANOVA. B, The osr-1(rm1) mutant animals do not display an increased turning rate in the hyperosmotic solution of 400 mOsm (n = 29) compared with the turning rate in 150 mOsm (n = 28). Both the mutant and WT animals are cultivated under the standard osmotic condition of 150 mOsm. n = 29 and 31 for wild-type animals tested in solutions of 150 and 400 mOsm, respectively. C, The dpy-2(e8) mutant animals do not display an increased turning rate in the hyperosmotic solution of 400 mOsm (n = 16) compared with the turning rate in 150 mOsm (n = 16). Both the mutant and wild-type animals are cultivated under the standard osmotic condition of 150 mOsm. n = 16 for WT animals tested in each condition. For B and C, significant interaction between genotype and osmolarity is tested by two-way ANOVA. For all, ***p < 0.001. Values are reported as the mean ± SEM. WT, wild type.

The aversion to osmotic upshift requires the cGMP-gated channel subunit TAX-2

To characterize the aversive response to the osmotic upshift, we sought the underlying sensory mechanisms by examining a pool of candidate genes.

First, we examined aquaporins, which are a family of membrane proteins that are conserved throughout the animal kingdom and regulate the transport of water and small-molecule solutes, such as glycerol, cross the cell membrane (Agre et al., 2002). Eleven aquaporin homologs have been identified in C. elegans (Park and Saier, 1996; Ishibashi and Sasaki, 1998; Morishita et al., 2004). We tested the effect of all available mutations in aqp-1, -2, -3, -4, -6, -7, -8, -9, -10, and -11 (Table 1). Among all the mutations tested, only the deletion mutation in aqp-9 generated a significant but small reduction in the increased turning rate induced by an osmotic upshift from 150 to 400 mOsm. These results suggest that the aquaporin water channel AQP-9 may play a role in the sensorimotor response to the osmotic upshift.

Next, we tested the mutations in a few mechanical sensory channels, because mechanical responses are implicated in sensing osmotic stimuli (Zhang et al., 2007). We tested mutations in asci-1 that encoded a DEG/ENaC/ASIC mechanosensory channel (Russell et al., 2014), in trp-4 that encoded the C. elegans homolog of the mechanosensitive TRPN channel (Li et al., 2006), as well as in mec-4 that encoded an amilioride-sensitive mechanosensory channel subunit (O’Hagan and Chalfie, 2006). None of the tested mutations generated a detectable defect in response to the upshift of osmolarity from 150 to 400 mOsm (Table 1). Therefore, the increased turning rate evoked by exposing to an upshift of osmolarity in C. elegans is likely different from the mechanosensory responses that are mediated by these channels.

To further explore the neuronal mechanisms underlying the sensory response of osmotic upshifts, we examined mutations in channels that were known to regulate responses to different chemosensory cues. Previous studies have shown that C. elegans acutely avoids solutions of a high concentration of sodium and generates reversals in a few seconds after a direct contact with the salt. This response requires the function of a sodium-sensitive channel transmembrane channel-like 1 (TMC-1) in the polymodal sensory neurons ASH (Chatzigeorgiou et al., 2013). A recent study shows that TMC-1 regulates an acute avoidance of alkaline pH in ASH (Wang et al., 2016). C. elegans generates a similar acute avoidance of osmotic shocks produced by solutions of high osmolarity, such as 1 m glycerol or 4 m fructose, and the avoidance depends on the function of a C. elegans TRPV channel, OSM-9, in the ASH sensory neurons (Colbert et al., 1997; Liedtke et al., 2003; Hilliard et al., 2004). We found that mutations in osm-9, tmc-1, or tmc-2, which encoded a homolog of tmc-1, did not disrupt the aversive behavioral response to the osmotic upshift (Fig. 3A, Table 1). In contrast, two canonical mutations, p691 and p671, in tax-2 that encoded a cGMP-gated channel subunit and was known to regulate multiple sensory responses, including olfaction (Bargmann, 2006), severely disrupted the increased turning rate in response to the osmotic upshift from 150 to 400 mOsm (Fig. 3B,C). The defects in both alleles were rescued by a genomic fragment that contained the regulatory and the coding regions of tax-2 (Fig. 3D,E). These results indicate that the cGMP-gated channel subunit TAX-2 is required for the sensorimotor response to the osmotic upshift and that the gradually generated aversive response to the osmotic upshift defines a new sensory-evoked behavior that is different from the avoidance of a high salt solution or osmotic shocks.

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Figure 3.

The cGMP-gated channel subunit TAX-2 acts in the AQR, PQR, and URX neurons to regulate the aversive response to osmotic upshifts. A, The mutant animals osm-9(ky10) cultivated under the standard condition display an increased turning rate in the hyperosmotic solution of 400 mOsm that is similar to that of WT animals. n = 31 and 29 WT animals tested in 150 and 400 mOsm, respectively; n = 32 and 31 osm-9 animals tested in 150 and 400 mOsm, respectively. B, C, Two mutation alleles of tax-2, p691 and p671, are severely defective in increasing the turning rate in response to the osmotic upshift from 150 to 400 mOsm, compared with WT animals. For B, n = 24 WT animals were tested in either 150 or 400 mOsm and n = 20 and 24 p691 animals were tested in 150 and 400 mOsm, respectively; for C, n = 23 and 24 WT animals were tested in 150 and 400 mOsm, respectively, and n = 21 and 27 p671 animals were tested in 150 and 400 mOsm, respectively. D, E, Expressing the genomic DNA fragment containing the coding region and regulatory sequences of tax-2 (Ptax-2::tax-2) rescues the defects in both tax-2 alleles. For D, n = 32 and 31 WT animals tested in 150 and 400 mOsm, respectively; n = 29 and 31 transgenic animals expressing Ptax-2::tax-2 tested in 150 and 400 mOsm, respectively; and n = 28 and 26 p691 mutant sibling animals tested in 150 and 400 mOsm, respectively; for E, n = 31 and 32 WT animals were tested in 150 and 400 mOsm, respectively; n = 31 and 32 transgenic animals expressing Ptax-2::tax-2 were tested in 150 and 400 mOsm, respectively; and n = 30 p671 mutant sibling animals were tested in either 150 or 400 mOsm. F, Expressing the wild-type tax-2 cDNA in the body cavity sensory neurons AQR, PQR, URX (Pgcy-36::tax-2) rescues the defects in the tax-2(p691) mutants in generating an increased turning rate in response to the osmotic upshift from 150 to 400 mOsm. n = 24 and 22 WT animals tested in 150 and 400 mOsm, respectively, n = 22 and 24 transgenic animals expressing Pgcy-36::tax-2 tested in 150 and 400 mOsm, respectively, n = 23 and 24 p691 mutant sibling animals tested in 150 and 400 mOsm, respectively. G, Expressing cell death-promoting molecule EGL-1 in the body cavity sensory neurons AQR, PQR, and URX (Pgcy-36::egl-1) abolished the increased turning rate in response to the osmotic upshift from 200 to 400 mOsm. n = 36 and 44 WT animals were tested in 200 and 400 mOsm, respectively; n = 40 and 44 transgenic animals expressing Pgcy-36::eg-1 were tested in 200 and 400 mOsm, respectively. H, The heat map of the calcium signals in the URX neuron in individual animals that were exposed to 150 mOsm and switched to 400 mOsm and in individual animals that were exposed to 150 mOsm and switched to 150 mOsm. The calcium signal was presented as the percentage change in GCaMP6 signal using the average GCaMP6 signal in the first 60 s as the baseline [(F − F_base)/F_base * 100%; see Materials and Methods]. I, The average change in the GCaMP6 signal in URX is significantly higher when animals are switched from 150 to 400 mOsm (n = 15) than when animals are switched from 150 to 150 mOsm (n = 13). Box plot shows the first and third quartile, median, and the maximum and minimum. The Student’s t test was used for comparison. For A–C and G, significant interaction between genotype (wild-type vs mutant animals) and osmolarity is tested with two-way ANOVA; for D–F, significant interaction between genotype (transgenic vs nontransgenic siblings) and osmolarity is tested with two-way ANOVA. For all, animals are cultivated under the standard NGM plates with an osmotic condition of 150 mOsm. ***p < 0.001, **p < 0.01, *p < 0.05. n.s., Not significant. Values are reported as the mean ± SEM. WT, wild type.

TAX-2 regulates osmotic response in the sensory neurons that are exposed to the body fluid

Several sensory neurons in C. elegans mediate sensorimotor responses through the cGMP-gated channel TAX-2/TAX-4 (Bargmann, 2006). Next, we asked in which sensory neurons tax-2 regulated the aversive response to osmotic upshifts.

First, we tested the effect of several mutations that generated defects in the development of subsets of the tax-2-expressing sensory neurons. We tested the ttx-1(p676) and the che-1(p679) mutants, which were defective in the development and differentiation of the sensory neurons AFD and ASE, respectively (Satterlee et al., 2001; Uchida et al., 2003). We also examined the transgenic animals that expressed a gain-of-function isoform of potassium channel twk-18 in the sensory neuron ADF [Psrh-142::twk-18(gf)] (Qin et al., 2013). The expression of twk-18(gf) hyperpolarizes and inhibits the activity of the expressing neurons (Kunkel et al., 2000). None of the transgenic animals or the mutant animals exhibited a detectable defect in their response to the osmotic upshift (Table 2). These results suggest that the sensory neurons ADF, AFD, or ASE are not required for the increased turning rate evoked by the upshift of osmolarity from 150 to 400 mOsm.

Next, to identify the tax-2-expressing sensory neurons that regulate the aversive response to the osmotic upshift, we expressed the wild-type tax-2 cDNA selectively in a few sensory neurons in the tax-2(p691) mutants and examined the potential rescuing effects. Because we found that exposure to the osmotic upshift from 150 to 400 mOsm for a few minutes generated a significant shrinkage of the worm body, we hypothesized that osmotic upshifts might alter the osmolarity of the body fluid. Thus, we expressed tax-2 in AQR, PQR, and URX (Pgcy-36::tax-2), which comprised a set of sensory neurons that were exposed to the body cavity. We found that the expression of tax-2 in AQR, PQR, and URX fully rescued the defect of the tax-2(p691) mutant animals in responding to the osmotic upshift from 150 to 400 mOsm (Fig. 3F). In contrast, expressing tax-2 in the main olfactory sensory neurons AWB and AWC using the odr-1 promoter (Podr-1::tax-2; Qin et al., 2013) or in the sensory neuron ASI (Pstr-3::tax-2) had no detectable rescuing effect in the aversive behavioral response (Table 3). These results together show that the function of TAX-2 in the body cavity neurons are sufficient to regulate the aversive response to the osmotic upshift. The ciliated endings or the somata of the AQR, PQR, and URX neurons are directly exposed to the body fluid and are referred to as the body cavity sensory neurons (White et al., 1986). To further examine the role of the body cavity neurons, we tested the transgenic animals that expressed the cell death-promoting molecule EGL-1 (Conradt and Horvitz, 1998) selectively in these neurons (Chang et al., 2006). We found that the transgenic animals displayed a behavioral response significantly different from that of wild-type animals. The animals without AQR, PQR, and URX generated a higher turning rate at 200 mOsm and did not significantly increase the turning rate on exposure to 400 mOsm (Fig. 3G). These results suggest that the body cavity neurons regulate the turning rate at 200 mOsm, which may prevent a further increase in the turning rated at the higher osmolarity. Nevertheless, the result reveals an important role of the body cavity neurons in regulating the turning rate under either the standard or higher osmotic condition or both. Together, these results indicate that the body cavity sensory neurons mediate the aversive response to the osmotic upshift via the function of the cGMP-gated channel subunit TAX-2.

To characterize the mechanisms underlying the function of the body cavity sensory neurons in mediating the aversive response to the osmotic upshift, we examined the intracellular calcium transients in the sensory neuron URX under the standard condition and during the osmotic upshift. We focused on the URX neuron because the function of URX in regulating the sensorimotor response to another environmental cue, oxygen, has been well characterized (Cheung et al., 2004; Gray et al., 2004; Zimmer et al., 2009; Witham et al., 2016). We used the transgenic lines that expressed a genetically encoded fluorescent reporter, GCaMP6 (Chen et al., 2013), in URX and a microfluidic device (Chronis et al., 2007) that allowed the head of the transgenic animal exposed to a stream of solution. The solution could be switched between two alternatives that were controlled by an air flow. We first exposed the transgenic animal to a solution of 150 mOsm for 60 s and then switched the stimulating fluidic stream to a solution of either 150 or 400 mOsm. We recorded the GCaMP6 signal in the cell body of the URX neuron and quantified the change of GCaMP6 signals in response to the switch of the solution (Materials and Methods). We found that URX generated calcium signals with higher amplitudes during the exposure to 400 mOsm than during the exposure to 150 mOsm (Fig. 3H,I). Although the increase in the URX calcium signal is moderate, it is significant. Previous studies show that URX responds to increased environmental oxygen concentrations by transiently increasing the intracellular calcium signals (Cheung et al., 2004; Gray et al., 2004; Zimmer et al., 2009; Witham et al., 2016) and that ectopically activating URX evokes aversive behavior (Busch et al., 2012). Together, these results demonstrate that the osmotic upshift from 150 to 400 mOsm activates the body cavity neuron URX to mediate the aversive behavioral response.

Previous studies have shown that guanylyl cyclases act downstream of G-protein-coupled receptors to produce cGMP that regulate the function of the cGMP-gated TAX-2/TAX-4 channels (Bargmann, 2006). Next, we examined the Gα subunits and the guanylyl cyclases that were expressed in the AQR, PQR, and URX neurons. The C. elegans genome encodes a large number of Gα proteins, GPA-1 to GPA-16, ODR-3, GOA-1, and ELG-30 (Jansen et al., 1999). We tested mutations in gpa-8 and gpa-13 that were expressed in AQR, PQR, and URX (Table 4). We found that neither of the mutations generated a significant defect in responding to the osmotic upshift. These results suggest that these Gα proteins are not required for the osmotic response or that they act in a redundant manner. We also examined mutations in the genes that encoded guanylyl cyclases and were shown to be expressed in the AQR, PQR, and URX neurons (Table 4). The pool included gcy-28, which encoded a receptor guanylyl cyclase and regulated an innate olfactory preference and integration (Tsunozaki et al., 2008; Shinkai et al., 2011); gcy-33, 35 and gcy-36, all of which encoded soluble guanylyl cyclases that mediated responses to oxygen (Cheung et al., 2004; Gray et al., 2004); as well as several other guanylyl cyclases whose functions were less well characterized (Yu et al., 1997; Ortiz et al., 2006). Again, we found that none of the mutations significantly altered the behavioral response to the osmotic upshift (Table 4). These results show that the tested guanylyl cyclases are not required for the response to the osmotic upshift or they act redundantly.

The neural circuit underlying navigational locomotion mediates the behavioral aversion to osmotic upshifts

To further understand how the osmotic upshift generates an aversive behavioral response, we probed the downstream neural circuit. The interneurons AIB, AIY, and AIZ are the first layer of interneurons that connect the sensory input with the downstream motor system (White et al., 1986). To test the potential role of these interneurons, we expressed the gain-of-function isoform of the potassium channel twk-18(gf) (Kunkel et al., 2000) to inhibit the activity of all or subsets of these interneurons (Luo et al., 2014) and examined the resulting effect on behavior. We found that expressing twk-18(gf) in either AIY alone [Pttx-3::twk-18(gf)] or primarily in AIB [Pinx-1::twk-18(gf)] generated no obvious phenotype in the aversive response evoked by the osmotic upshift (Table 5). In contrast, combining the two transgenes [Fig. 4A, Pttx-3::twk-18(gf); Pinx-1::twk-18(gf)] significantly reduced the increase in the turning rate in response to the osmotic shift from 150 to 400 mOsm. In addition, expressing twk-18 in AIB, AIY, and AIZ together [Fig. 4B, Pttx-3::twk-18(gf); Podr-2b(3a)::twk-18(gf)] similarly suppressed the aversive behavior, but expressing twk-18 mainly in AIB and AIZ [Podr-2b(3a)::twk-18(gf)] was not sufficient to generate a detectable defect (Table 5). Together, these results show that the interneurons AIB and AIY together play a significant role in mediating the behavioral aversion to the osmotic upshift.

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Figure 4.

The neural circuit underlying navigation regulates the aversive response to osmotic upshifts. A, Inhibiting the activity of the interneurons AIB and AIY by expressing twk-18(gf) produces significant defects in generating an increased turning rate in response to the osmotic upshift from 150 to 400 mOsm. n = 40 and 38 transgenic animals expressing twk-18(gf) in AIB and AIY were tested in 150 and 400 mOsm, respectively; and n = 40 and 38 WT sibling animals were tested in 150 and 400 mOsm, respectively. B, Inhibiting AIB, AIY, and AIZ by expressing twk-18(gf) produces significant defects in generating an increased turning rate in response to the osmotic upshift from 150 to 400 mOsm. n = 40 and 33 transgenic animals expressing twk-18(gf) in AIB, AIY, and AIZ were tested in 150 and 400 mOsm, respectively; n = 40 and 39 WT sibling animals were tested in 150 and 400 mOsm, respectively. C, Blocking the neural transmission of several command interneurons and motor neurons by expressing the tetanus toxin with the glr-1 promoter produces significant defects in generating an increased turning rate in response to the osmotic upshift from 150 to 400 mOsm. n = 24 animals in all conditions. D, Blocking the neural transmission of several motor neurons by expressing the tetanus toxin with the lad-2 promoter produces significant defects in generating an increased turning rate in response to the osmotic upshift from 150 to 400 mOsm. n = 23 transgenic animals expressing the tetanus toxin with the lad-2 promoter were tested in either 150 or 400 mOsm; and n = 22 and 23 WT sibling animals were tested in 150 and 400 mOsm, respectively. For all, animals are cultivated on the standard NGM plates with an osmotic condition of 150 mOsm, and the significant interaction between genotype (transgenic vs nontransgenic siblings) and osmolarity is tested using two-way ANOVA, ***p < 0.001, **p < 0.01. Values are reported as the mean ± SEM. WT, wild type.

Previous findings show that AIB and AIY interneurons act upstream of several command interneurons and motor neurons to regulate reversals and turns (Tsalik and Hobert, 2003; Gray et al., 2005; Ha et al., 2010). Thus, we further examined the locomotory circuit. We found that expressing the tetanus toxin (Schiavo et al., 1992) with the glr-1 promoter to block presynaptic release from several command interneurons and motor neurons (Hendricks et al., 2012) completely abolished the increase in the turning rate induced by the osmotic shift (Fig. 4C). The glr-1 promoter is expressed in several command interneurons, including AVA, which is known to regulate reversals, and a few motor neurons, such as SMD and RIM that regulate head bending and turns during escape responses (Chalfie et al., 1985; Brockie et al., 2001; Gray et al., 2005; Ha et al., 2010). We also used the lad-2 promoter (Wang et al., 2008) to block neurotransmission from several motor neurons that regulate head bending, including SMD (Hendricks et al., 2012). We found that the mutation also generated a significant defect in the osmotic upshift-induced increase in the turning rate (Fig. 4D). Together, these results indicate that the aversive response to the osmotic upshift uses the neural circuit that underlies the head bendings and turns to regulate navigational locomotion.