A Caenorhabditis elegans Pheromone Antagonizes Volatile Anesthetic Action Through a Go-Coupled Pathway

Corresponding author: C. Michael Crowder, Box 8054, Washington University School of Medicine, 660 S. Euclid Ave., St. Louis, MO 63110., crowderm{at}morpheus.wustl.edu (E-mail)

Communicating editor: B. J. MEYER

	ABSTRACT

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION LITERATURE CITED

Volatile anesthetics (VAs) disrupt nervous system function by an ill-defined mechanism with no known specific antagonists. During the course of characterizing the response of the nematode C. elegans to VAs, we discovered that a C. elegans pheromone antagonizes the VA halothane. Acute exposure to pheromone rendered wild-type C. elegans resistant to clinical concentrations of halothane, increasing the EC₅₀ from 0.43 ± 0.03 to 0.90 ± 0.02. C. elegans mutants that disrupt the function of sensory neurons required for the action of the previously characterized dauer pheromone blocked pheromone-induced resistance (Pir) to halothane. Pheromone preparations from loss-of-function mutants of daf-22, a gene required for dauer pheromone production, lacked the halothane-resistance activity, suggesting that dauer and Pir pheromone are identical. However, the pathways for pheromone's effects on dauer formation and VA action were not identical. Not all mutations that alter dauer formation affected the Pir phenotype. Further, mutations in genes not known to be involved in dauer formation completely blocked Pir, including those altering signaling through the G proteins Go and Gq. A model in which sensory neurons transduce the pheromone activity through antagonistic Go and Gq pathways, modulating VA action against neurotransmitter release machinery, is proposed.

THROUGH an unknown mechanism, volatile anesthetics (VAs) disrupt the behavior of all metazoans. In humans, VAs block memory formation, consciousness, and volitional movement, thereby forming the basis for most surgical anesthesia. Identifying VA targets and the mechanism whereby they alter nervous system function has been a longstanding and difficult effort. A major limitation in anesthetic mechanism research has been the lack of genetic or pharmacologic inhibitors of anesthetic potency in vivo. Mutations or drugs that produce high-level resistance to VAs have not to our knowledge been described in vertebrates. Screens in Drosophila have isolated mutant stains that are modestly VA resistant to some anesthetic endpoints (KRISHNAN and NASH 1990 ; TINKLENBERG et al. 1991 ; LEIBOVITCH et al. 1995 ; GAMO et al. 1998 ); however, highly resistant mutants have not thus far been uncovered. In Caenorhabditis elegans, several mutants have been found to be markedly resistant to clinical concentrations of VAs, and the implicated genes have been placed in a pathway that regulates neurotransmitter release in C. elegans and in higher organisms (VAN SWINDEREN et al. 1999 , VAN SWINDEREN et al. 2001 ). At this point it is unclear if the molecular mechanisms being defined genetically in Drosophila and C. elegans will overlap. However, at least at a cellular level VA mechanisms in the two organisms may be similar, given that two of the halothane-resistant Drosophila mutants have been shown to partially antagonize the effects of halothane on glutamate release at the Drosophila larval neuromuscular junction (NISHIKAWA and KIDOKORO 1999 ).

The fact that in C. elegans single gene mutations can confer high-level resistance to clinical concentrations of VAs indicates that one major mechanism is acting at these concentrations in C. elegans. Thus, pharmacological antagonism of VA potency would be biologically possible in C. elegans if the proper drug were identified. In this study, we describe the serendipitous discovery of a C. elegans pheromone that is capable of antagonizing VAs in C. elegans. Importantly, the pheromone is not a stimulant of the behavior that VAs disrupt. In other words, the pheromone alters the effects of the drug on behavior, not the behavior itself. This pheromone antagonizes VA action through mechanisms previously implicated genetically to regulate dauer formation and VA sensitivity in the absence of pheromone in C. elegans. Our results confirm the importance of these gene products in VA mechanisms and demonstrate that pharmacologic antagonism of general anesthetics is possible in vivo. Further, they show that a pheromone can modulate nervous system function in adult C. elegans and that the pheromone is likely to be the same pheromone that controls dauer formation in C. elegans.

	MATERIALS AND METHODS

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION LITERATURE CITED

Nematode strains and conditions:
C. elegans mutant strains were obtained from the Caenorhabditis Genetics Center and from several laboratories whose research is referenced in this work. All assays were performed on well-fed young adult animals (1 day post-L4 stage) at room temperature (22°–24°). Strains were grown on uncrowded conditions (100–300 animals per plate) as described previously (BRENNER 1974 ) on nematode growth media (NGM) agar plates seeded with OP50 bacteria.

Behavioral assays:
VAs were delivered to C. elegans as described previously (CROWDER et al. 1996 ). Halothane or isoflurane were injected as liquids onto the tops of sealed glass chambers containing the assay plates. Gas-phase VA concentrations were measured by gas chromatography.

The effect of VAs on locomotion was quantified by the dispersal assay (CROWDER et al. 1996 ). Briefly, animals were washed off NGM plates into 1.5-ml polypropylene tubes, rinsed twice with S-basal, rinsed once with water, and then resuspended in 100 µl of water. Ten-microliter aliquots containing 50–100 worms were placed onto the center of dispersal assay plates (10-cm NGM plates seeded with a narrow ring of OP50 Escherichia coli along the edge of the plate). The plates were immediately placed into glass chambers, to which anesthetic was added. As soon as the worm-filled water drop dried (usually 2–5 min), the chambers were briefly shaken until the nematodes were induced to unclump and begin dispersing. After 45 min, the fraction of animals reaching the bacterial ring divided by the total number of worms was scored as the dispersal index. Incubation-induced resistance to VAs was measured by the dispersal assay with one difference. Instead of aliquoting the nematodes to dispersal plates immediately after the washing steps, the nematodes were allowed to remain in the 100 µl distilled water for 30 min prior to aliquoting onto dispersal plates. Sensitivity to VA-induced chemotaxis defects was measured as described previously (CROWDER et al. 1996 ). The behavioral assay is similar to the dispersal assay except that after the final wash animals are placed on chemotaxis plates, which are agar plates spotted with a chemoattractant, near the edge of the plate. The chemotaxis index was defined as the number of animals present after 2 hr within 0.5 cm of the attractant—the number of animals at an opposing control spot divided by the total number of animals on the plate. Sensitivity to VA-induced mating defects was measured as described previously (CROWDER et al. 1996 ). Ten young adult males and two dpy-11(e224) hermaphrodites were placed on each of five 3-cm plates seeded with a small spot of bacteria. The five plates were placed into a chamber along with a given concentration of VA for 24 hr after which the males were removed. The mating index for a given chamber was calculated as the fraction of plates with cross-progeny. Concentration/response data were fit by nonlinear regression to the VA EC₅₀ (the VA concentration where the effect is half maximal), which is used as the measure of VA sensitivity. Significant difference between EC₅₀'s was determined by simultaneous curve fitting as described previously (VAN SWINDEREN et al. 1997 ).

Pheromone extract and assays:
A crude C. elegans pheromone extract was prepared as described for dauer pheromone (GOLDEN and RIDDLE 1982 ). The substance was resuspended in distilled water, producing an oily yellow liquid, and stored at -20°. Dauer studies have calibrated the potency of the extract by measuring the dose required to induce 100% dauer larvae formation in the wild-type strain N2 at 20° (GOLDEN and RIDDLE 1984 ). However, even at 10% concentration our pheromone preparations induced only 50% dauer formation, suggesting that it may have been dilute.

Pheromone-induced resistance (Pir) to VAs was measured by the dispersal and chemotaxis assays as follows. Ten microliters of pheromone extract was diluted in each 990 µl of wash (three S-Basal and one distilled water), thus producing a 1% pheromone extract during the standard nematode washing steps prior to the dispersal assay. Following the washes, the nematodes were resuspended in a 50- to 100-µl aliquot of the same 1% pheromone extract in distilled water and immediately aliquoted to dispersal or chemotaxis plates, and subsequently the respective assays were as described above. For Pir in the mating assay, pheromone was added to OP50 bacteria to produce a 5% pheromone concentration, and the mating assay plates were seeded with a thin lawn of pheromone-containing bacteria 1 day prior to the assay.

	RESULTS

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION LITERATURE CITED

Incubation-induced resistance to halothane:
During the course of developing a high-throughput assay to quantitate VA-induced locomotion defects in C. elegans, we found significant variability in the VA sensitivity of the wild-type C. elegans strain N2. The assay, called the dispersal assay, measures the ability of a population of animals to disperse from the center of an agar plate to the edge. Dispersal, like other behaviors requiring coordinated locomotion, is particularly sensitive to VAs and is abolished at concentrations similar to those that anesthetize humans (CROWDER et al. 1996 ). The variability in VA sensitivity as measured by the dispersal assay was not random; rather, it was systematic. That is, for a particular dispersal assay, the sensitivity of all animals was consistent and well fit by a single sigmoidal concentration/response curve. However, between assays VA EC₅₀'s varied as much as 50%, and the distribution of the EC₅₀'s was bimodal. Ultimately, we hypothesized that the variability in VA sensitivities was due to the length of time the animals were allowed to remain in liquid prior to being aliquotted onto assay plates. To test this hypothesis, we compared the VA sensitivity of animals allowed to remain in liquid for 30 min prior to aliquotting to those immediately aliquotted (Fig 1). The EC₅₀ of incubated animals for the VA halothane was 0.79 ± 0.02 vol%, nearly double that of the nonincubated controls (0.42 ± 0.03 vol%).

View larger version (19K): In this window In a new window Download PPT slide   Figure 1. Incubation-induced resistance to halothane. (A) Incubation of wild-type N2 animals conferred resistance to halothane by the dispersal assay. This effect was significant by both simultaneous curve-fitting algorithms (WAUD 1972 ; DELEAN et al. 1978 ) and analysis of variance for 10 separate EC50 estimates for each condition (P < 0.05). Because of a large number of points, data for nonincubated N2 are pooled and averaged (±SEM). (B) Dispersal indices in the presence of equal concentrations of halothane (EC50 for no incubation condition) were scored under four conditions: (1) 0 hr—spotting the worms immediately after resuspension in water; (2) 4 hr—after incubation of the worms for 4 hr in water; (3) Sup—immediate spotting of worms resuspended in conditioned supernatant of other worms incubated for 4 hr; (4) Refresh—removing the supernatant of worms incubated for 4 hr and replacing with fresh water prior to spotting the animals on the dispersal plates. Shown are mean ± SEM dispersal indices (n = 3 experiments). *, significant resistance compared to the "0 hr" condition by ANOVA (P < 0.05); #, significant reversal of resistance compared to the "4 hrs" condition by ANOVA (P < 0.05).

We hypothesized that the VA resistance was conferred by a soluble product secreted by C. elegans into the water during the 30-min incubation step. To test this possibility, we added the supernatant of animals incubated for 4 hr to a nonincubated population of worms. This conditioned supernatant added to freshly washed animals produced a small but insignificant level of halothane resistance (Fig 1B). We also did the complementary experiment of replacing (refreshing) the supernatant of animals incubating for 4 hr with a final wash of fresh water. A 4-hr incubation conferred resistance to halothane, and when the 4-hr incubated animals were refreshed with their final wash, their halothane sensitivity returned to normal nonincubation levels. These experiments were consistent with the hypothesis that the incubation-induced resistance resulted from a secreted substance. Alternative explanations included starvation or hypoxia-induced VA resistance. The normalization of the VA sensitivity of 4-hr-incubated animals by replacement of the supernatant was inconsistent with starvation as the cause. However, these experiments did not rule out hypoxia.

A pheromone extract confers resistance to halothane:
To directly test the hypothesis that a secreted pheromone antagonizes VAs in C. elegans, we tested the effects of a pheromone extract on VA sensitivity. We prepared the pheromone extract according to the previously described protocol for isolation of dauer pheromone, which promotes larval diapause in C. elegans (GOLDEN and RIDDLE 1982 ). For testing of its effects on VA sensitivity, the pheromone was diluted to a 1% concentration into the dispersal assay wash buffers. This pheromone concentration was initially chosen on the basis of the published potency of dauer pheromone for inducing dauer formation and proved to be maximally effective at antagonizing VA sensitivity (Fig 2C). The dispersal assays were performed exactly like the no-incubation assays except they used the 1% pheromone buffers for the washing steps prior to transferring the animals onto the assay plates. The pheromone induced significant halothane resistance, increasing the dispersal EC₅₀ from 0.42 ± 0.03 to 0.95 ± 0.02 vol% (Fig 2A). The pheromone had no effect on locomotion in the absence of anesthetic (Fig 2B), indicating that the resistance was not merely secondary to making the animals hyperactive. The pheromone-induced resistance was dose dependent with a maximal effect achieved at a 1% concentration (Fig 2C). A similar extract from media containing only bacteria but no C. elegans did not induce halothane resistance (data not shown). Thus, the resistance activity requires and presumably is secreted by the worms. We refer to this phenotype as Pir (pheromone-induced resistance to volatile anesthetics).

View larger version (52K): In this window In a new window Download PPT slide   Figure 2. Pheromone-induced resistance to halothane. (A) N2 animals were exposed to 1% C. elegans pheromone extract immediately prior to placement onto dispersal assay plates (see MATERIALS AND METHODS). The raw data for pheromone-treated N2 animals are shown in comparison to averaged data for untreated N2. Resistance was significant by curve-fitting algorithms (WAUD 1972 ; DELEAN et al. 1978 ) and ANOVA of EC50's (P < 0.05). (B) Pheromone extract did not produce significant differences in N2 dispersal in the absence of halothane. Dispersal index was plotted against time for the length of the dispersal assay (45 min). Two separate experiments for each treatment are combined. A best-fit curve of the data showed that it takes 20 min for animals to perform half-maximally for both treatments. The curves were not significantly different. (C) N2 wild-type animals were exposed to different doses of pheromone. The data shown here are EC50's ± SE for the dispersal endpoint. The 0.5, 1.0, and 5.0% pheromone concentrations produced significant halothane resistance relative to the no pheromone control. (D) The pheromone extract induced significant resistance to halothane action against male mating behavior but not chemotaxis toward a volatile odorant. For male mating, 50 µl of pheromone extract was diluted into 1 ml of OP50 bacteria. This 5% OP50 solution was seeded onto male mating assay plates; control plates were seeded with straight OP50. The fraction of plates with successful mating after a 24-hr period was scored as the mating efficiency and plotted against halothane concentration to determine halothane EC50's. In the chemotaxis assay, the method for exposure to pheromone was identical to that for the dispersal assay. The fraction of animals at the odorant spot minus the fraction at the control was scored as the chemotaxis index and plotted against halothane concentration to determine EC50's. +P, pheromone treatment; -P, no pheromone control.

Pheromone extract also conferred resistance to halothane's effects on male mating behavior (Fig 2D). This anesthetic assay involves a completely different behavior and set of neurons and is disrupted by halothane with an EC₅₀ = 0.52 ± 0.02 vol% for the N2 strain (Fig 2D). Male mating plates seeded with an E. coli (OP50) solution mixed with pheromone extract significantly increased the halothane EC₅₀ to 0.91 ± 0.06 vol% against male mating. However, chemotaxis, a third behavior abolished by clinical concentrations of halothane (CROWDER et al. 1996 ), was unaffected by pheromone extract even though the chemotaxis assay involves washing procedures identical to those used for the dispersal assay. We have previously concluded, on the basis of the odorant dependence of the sensitivity of chemotaxis to anesthetics, that VAs do not disrupt chemotaxis through their effects on locomotion (CROWDER et al. 1996 ). The lack of effect of pheromone on VA-induced chemotaxis defects is further evidence that VA mechanisms acting on locomotion and chemotaxis behaviors are distinct.

Dauer pheromone pathway:
To define the mechanism underlying the pheromone's antagonism of halothane, we first tested strains carrying mutations in genes in the dauer formation pathway. Dauers are an alternative larval form whose formation is promoted by heat, starvation, and dauer pheromone (RIDDLE and ALBERT 1997 ). We speculated that the dauer pheromone and its transduction pathway were responsible for pheromone-induced halothane resistance. The most upstream component of the dauer pathway (Fig 3A) is daf-22, which is required for the biosynthesis of functional dauer pheromone (GOLDEN and RIDDLE 1985 ). If daf-22 is also required for the pheromone regulating VA sensitivity, then a pheromone preparation from daf-22 loss-of-function mutants should lack Pir activity. Indeed, daf-22 (m130lf) pheromone does not induce halothane resistance (Fig 3B). As expected for a gene thought to be involved in the biosynthesis of the pheromone, daf-22(m130lf) is not insensitive to wild-type pheromone-induced halothane resistance (Table 1). Thus, daf-22 is required for the activity but not response to both the dauer and Pir pheromones, consistent with these two pheromones being synthesized by a DAF-22-dependent mechanism and being the same pheromone.

View larger version (33K): In this window In a new window Download PPT slide   Figure 3. Effect of dauer-formation mutants on Pir. (A) The genetic pathway for control of dauer formation (THOMAS et al. 1993 ). The daf-2 branch of the pathway is not shown because it was not tested for effects on Pir. (B) Comparison of the effects of no pheromone, N2 pheromone, and pheromone made from daf-22(m130lf) on halothane sensitivity. daf-22(m130lf) disrupts production of dauer pheromone (GOLDEN and RIDDLE 1985 ). Halothane sensitivity was measured by halothane concentration/response curves against the dispersal behavior.

Dauer pheromone is detected by a set of sensory organs called amphids, which contain specialized ciliated sensory neurons exposed to the environment through a pore in the cuticle (ALBERT et al. 1981 ; BARGMANN and HORVITZ 1991 ). We tested mutants in genes required for the function of the amphid neurons and found that, as for dauer formation, these mutants are Pir defective (Table 1). Thus, the Pir activity requires the normal function of the amphid neurons, which likely detect the pheromone. daf-11 and daf-21 code for homologs of transmembrane guanylyl cyclase and HSP-90, respectively, both of which function upstream of the cilium structure genes to regulate dauer formation (BIRNBY et al. 2000 ). Both daf-11 and daf-21 mutants are Pir defective (Table 1), a result consistent with the proposed role of these genes in the function of amphid neurons. Reduction-of-function mutants in the TGFß-SMAD arm (daf-1, daf-4, daf-3, and so on; PATTERSON and PADGETT 2000 ) of the dauer pathway were normally responsive to pheromone-induced resistance to halothane (Table 1). However, mutants reducing TGß signaling (daf-1, daf-7, and daf-8) did increase native halothane sensitivity (i.e., in the absence of pheromone), and daf-3, daf-5, and daf-12 mutants, which suppress the Daf-c phenotypes of daf-1, daf-7, and daf-8, were significantly halothane resistant. Thus, while pheromone-induced VA resistance is not particularly sensitive to alterations in TGFß-SMAD signaling, native VA sensitivity appears to be.

goa-1 signaling pathway:
Various clues suggested the C. elegans Go pathway as a candidate for a mediator of pheromone's antagonism of anesthetics. G-protein -subunits transduce pheromone action in yeast (SONG and DOHLMAN 1996 ; DAVIS and DAVEY 1997 ; LEBERER et al. 1997 ). We have previously found that a dominant-negative mutation in the goa-1 gene, which codes for the -subunit of Go (MENDEL et al. 1995 ; SEGALAT et al. 1995 ), and gain-of-function mutations in egl-10, which codes for an RGS protein negatively regulating GOA-1 (KOELLE and HORVITZ 1996 ), confer a twofold resistance to halothane, similar in magnitude to that produced by pheromone (VAN SWINDEREN et al. 2001 ). We tested mutants in the goa-1 pathway to determine their role, if any, in pheromone signaling. We found that all mutations that reduce GOA-1 signaling are Pir defective, including the normally halothane-sensitive goa-1(null) alleles, the already halothane-resistant dominant-negative goa-1(sy192) allele, and egl-10(gf) alleles (Table 2). Interestingly, egl-10(lf) mutants are also Pir defective. Thus, both diminished and enhanced Go signaling disrupt pheromone's halothane antagonism. The regulation of C. elegans locomotion by GOA-1 is dependent on the normal function of diacyl glycerol kinase, coded for by sag-1 (suppressor of activate goa-1), also known as dgk-1 (HAJDU-CRONIN et al. 1999 ). We tested a sag-1(rf) mutant to determine if pheromone signaling also utilized this pathway. Indeed, sag-1(sy428) was found to be Pir defective (Table 2). Like goa-1(null) mutants, sag-1(sy428) is not halothane resistant in the absence of pheromone, indicating that pheromone does not produce halothane resistance solely by inhibiting the GOA-1/DGK-1 pathway. Go acts antagonistically with Gq to regulate transmitter release, Go negatively regulating release and Gq positively regulating it (HAJDU-CRONIN et al. 1999 ; LACKNER et al. 1999 ; MILLER et al. 1999 ; NURRISH et al. 1999 ). eat-16(sy438) is a reduction-of-function mutant isolated as a suppressor of goa-1(gf) (HAJDU-CRONIN et al. 1999 ). eat-16 codes for an RGS protein that functions to negatively regulate Gq and positively regulate Go. Thus, we predicted that eat-16(rf) mutants should phenocopy the Pir-defective phenotype of goa-1(lf) and egl-10(gf). Indeed, eat-16(sy438) is Pir defective. Moreover, like the dominant-negative goa-1(sy192) and egl-10(gf), but unlike goa-1(lf), sy438 is halothane resistant in the absence of pheromone. These data all support a role for Go and Gq in pheromone's antagonism of halothane.

View this table: In this window In a new window   Table 2. Effect of Go pathway mutations on Pir

Neurotransmitter/receptor pathways:
What might be the neurotransmitter/receptor systems through which pheromone antagonizes halothane? We tested neurotransmitter/receptor mutants that might reasonably affect Pir (Table 3). Multiple lines of evidence suggest that VAs may act in part in the vertebrate nervous system by enhancing GABAergic signaling (JONES et al. 1992 ; FRANKS and LIEB 1994 , FRANKS and LIEB 1998 ; MIHIC et al. 1997 ; KOLTCHINE et al. 1999 ; JENKINS et al. 2001 ). We hypothesized that pheromone might produce resistance by modulating the effect of VAs on GABAergic signaling. To examine this hypothesis, we measured the Pir phenotypes of unc-25(lf) and unc-49(lf), which each abolish GABA neurotransmission in C. elegans neurons by defects in GABA synthesis and in the GABA_A receptor, respectively (MCINTIRE et al. 1993 ; BAMBER et al. 1999 ; RICHMOND and JORGENSEN 1999 ). We found that neither unc-25 (e156lf) nor unc-49(e382lf) altered Pir nor were these mutants halothane resistant in the absence of pheromone (Table 3). These results eliminate the possibility of the GABA_A receptor mediating halothane action against locomotion in C. elegans. Another reasonable candidate for a neurotransmitter pathway either mediating or modulating Pir is the serotonergic pathway. In C. elegans, serotonin signals through GOA-1 and could be the neurotransmitter that pheromone is modulating (MENDEL et al. 1995 ; SEGALAT et al. 1995 ; NURRISH et al. 1999 ). However, bas-1(ad446), which lacks serotonin immunoreactivity, cat-2(e1112), which is defective in dopamine synthesis, and cat-4(e1141), which disrupts both dopamine and serotonin-mediated signaling (LOER and KENYON 1993 ), were all wild type for Pir (Table 3). Thus, the neurotransmitter system through which pheromone modulates halothane's action on locomotion is unclear.

Presynaptic machinery:
Several lines of evidence indicate that VAs inhibit the release of neurotransmitters in both vertebrates and C. elegans (ZORYCHTA and CAPEK 1978 ; TAKENOSHITA and TAKAHASHI 1987 ; KULLMANN et al. 1989 ; MIAO et al. 1995 ; PEROUANSKY et al. 1995 ; SCHLAME and HEMMINGS 1995 ; MACIVER et al. 1996 ; VAN SWINDEREN et al. 1999 , VAN SWINDEREN et al. 2001 ; NISHIKAWA and MACIVER 2000 ). In C. elegans, mutations that reduce transmitter release are VA hypersensitive and those that increase release are resistant (VAN SWINDEREN et al. 1999 , VAN SWINDEREN et al. 2001 ). The extreme exception is unc-64(md130), which has reduced transmitter release yet is markedly resistant to halothane and other VAs, more so than that produced by pheromone treatment or by goa-1(rf) mutants (VAN SWINDEREN et al. 1999 ). These large allelic differences that cannot be explained by an indirect effect on transmitter release implicate syntaxin or syntaxin-binding proteins as an essential, perhaps binding, component of the VA mechanism. We investigated the effect of mutations in genes involved in presynaptic release of neurotransmitter on the Pir phenotype (Table 4). Both the VA-resistant unc-64 syntaxin allele md130 and the VA hypersensitive allele js21 responded to pheromone by increasing their halothane EC₅₀'s (Table 4). A double-mutant strain carrying both goa-1(null) and unc-64(md130) combined the properties of the two mutants. The strain was halothane resistant in the absence of pheromone like unc-64(md130) but was unresponsive to pheromone like goa-1(n363). These results indicate that unlike goa-1(lf) mutations, the mechanism whereby syntaxin mutations alter VA sensitivity does not disrupt pheromone's mechanism, and the resistance of unc-64 (md130) does not depend on pheromone signaling. Likewise, reduction-of-function mutations in ric-4 and snb-1, which code for the syntaxin-binding SNARE proteins SNAP-25 and VAMP (Rand AND NONET 1997 ; NONET et al. 1998 ), respectively, do not block Pir.

To test whether pheromone itself alters transmitter release, we measured pheromone's effects on aldicarb sensitivity. Aldicarb is an acetylcholinesterase inhibitor that is routinely used to assess the effect of mutations on transmitter release (Rand AND NONET 1997 ). Mutations or drugs that confer resistance to aldicarb generally reduce neurotransmitter release at the neuromuscular junction while aldicarb hypersensitivity is found in mutants with increased cholinergic neurotransmission. We have previously shown that VAs induce aldicarb resistance, indicating that VAs inhibit acetylcholine release; unc-64(md130) but not goa-1(lf) blocks VA-induced aldicarb resistance, suggesting that VAs act downstream of GOA-1 but upstream of UNC-64 to inhibit transmitter release (VAN SWINDEREN et al. 1999 , VAN SWINDEREN et al. 2001 ). Pheromone might produce VA resistance by increasing transmitter release and thereby indirectly antagonizing VAs. However, pheromone had no effect on native aldicarb sensitivity (Fig 4A) nor did it alter the potency of VAs to induce aldicarb resistance (Fig 4B). Thus, as for locomotion, pheromone does not appear to alter indirectly the effects of halothane on cholinergic neurotransmitter release in the absence of VAs.

View larger version (13K): In this window In a new window Download PPT slide   Figure 4. Effect of pheromone on aldicarb action. Young adult N2 were placed on plates containing 0.5 mM aldicarb that either did or did not contain, in addition, 5% pheromone. (A) 30–50 animals per condition were scored for paralysis (lack of movement in response to touch) immediately and at 1, 2, 3, 4, and 5 hr after placement on the plates. The percentage paralyzed represents mean ± SEM for two independent experiments for each condition. The percentage paralyzed on pheromone-containing plates was not significantly different from the percentage without pheromone at any timepoint. (B) N2 were preincubated for 4 hr on aldicarb plates with or without pheromone and then moved to a 0.5-cm-diameter circle marked on the plates. The plates were then placed into chambers containing either no halothane or 0.2–0.5 vol% halothane, and the fraction of animals (n = 20–40/plate) moving out of the circle after 1 hr was scored as the movement index. Halothane produced a significant increase in the movement index (*P < 0.05 by ANOVA). The movement indices for the pheromone-containing plates were not significantly different from the no-pheromone plates.

	DISCUSSION

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION LITERATURE CITED

The mechanism of volatile anesthetics has long been postulated to be nonspecific. Nonspecific theories of anesthesia propose that volatile anesthetics imbed into membranes and disrupt membrane structure, thereby altering the function of numerous membrane-associated proteins. These theories predict that high-level resistance to anesthetics cannot be achieved by altering a single mechanism. The lack of specific antagonism of volatile anesthetics by a drug has heretofore been a pharmacologic argument for nonspecific theories of anesthesia. Pheromone's antagonism of halothane now demonstrates that pharmacologic antagonism of VAs is biologically possible.

A pathway for pheromone's effects on halothane action consistent with the genetic data is shown in Fig 5. The pathway is drawn to summarize and discuss the data, but given the lack of testable null mutants in syntaxin and syntaxin-interacting proteins as well as other issues discussed below, the pathway should be considered a working model at this point. Pheromone-induced halothane resistance is dependent on the normal function of the amphid sensory neurons. If pheromone constitutively antagonized VAs through the amphid neurons, then the cilium structure mutants that disrupt amphid function would be expected to have abnormal VA sensitivities. However, the cilium structure mutants have normal halothane sensitivities in the absence of applied pheromone, indicating that amphid neurons regulate VA sensitivity only in the presence of high pheromone concentrations. At least from an anatomical standpoint, the amphid neurons might reasonably modulate VA effects on locomotion, given that they synapse onto interneurons that coordinate locomotion (CHALFIE et al. 1985 ; WHITE et al. 1986 ).

View larger version (11K): In this window In a new window Download PPT slide   Figure 5. Proposed pathway for pheromone-induced VA resistance. Transduction of the pheromone signal requires and is presumably sensed by the amphid sensory neurons. Pheromone signaling through the amphid neurons or in the neuronal pathway downstream of amphid neurons requires the normal activity of both Go- and Gq-coupled pathways. The proposal of a positive modulation by amphid neurons on Gq activity, coded for by egl-30, and an inhibitory modulation on Go signaling is based on the VA resistance of goa-1(lf) and egl-30(gf) (VAN SWINDEREN et al. 2001 ). GOA-1, EGL-30, and their RGS proteins, EGL-10 and EAT-16, respectively, have previously been shown to modulate VA action against locomotion and transmitter release (VAN SWINDEREN et al. 2001 ). On the basis of the high-level resistance of a dominant-negative form of syntaxin (VAN SWINDEREN et al. 1999 ), VAs are shown to inhibit directly the function of a syntaxin-interacting protein that positively regulates transmitter release; however, the data are also consistent with VAs enhancing the function of a negative regulator of release.

We have previously shown that goa-1(null) mutants are isoflurane but not halothane resistant, whereas sy192, a dominant-negative goa-1 allele, and mutants in the RGS-protein-coding genes, egl-10 and eat-16, are both halothane and isoflurane resistant (VAN SWINDEREN et al. 2001 ). Pheromone also produces both halothane (Fig 2) and isoflurane resistance (data not shown). How do pheromone, sy192, and egl-10(gf) produce halothane resistance when goa-1(null) mutations do not? Given that loss-of-function mutants in eat-16, which normally negatively regulates egl-30, are both halothane and isoflurane resistant and pheromone unresponsive, we speculate that pheromone, sy192, and egl-10(gf) alter both goa-1 and egl-30 signaling. A mechanism whereby this might occur has recently been delineated through the characterization of mutants of gpb-2, which codes for Gß₅-like protein. GPB-2 was shown to interact with both EGL-10 and EAT-16 (CHASE et al. 2001 ; ROBATZEK et al. 2001 ; VAN DER LINDEN et al. 2001 ) and thereby enhance the GTPase activity of both RGS proteins. EGL-10 overexpression not only inhibits the function of GOA-1 but also enhances the function of EGL-30 by sequestering GPB-2 away from EAT-16 (CHASE et al. 2001 ; ROBATZEK et al. 2001 ; VAN DER LINDEN et al. 2001 ). Likewise, eat-16(lf) enhances egl-30 function while inhibiting that of goa-1. In light of these new findings, the stronger anesthetic and locomotion phenotypes of goa-1 (sy192) compared to goa-1(null) can be explained by a dominant-negative action against GPB-2 and thereby enhancement of EGL-30 function. Thus, we propose that the halothane resistance of pheromone, sy192, and the RGS mutants is produced primarily by increasing the activity of Gq and that the goa-1(null) mutants block the effect of pheromone by increasing Gq activity to a level that cannot be further increased by pheromone.

An instructive aspect of pheromone's action is its lack of effect on the behavior and synapses that are disrupted by VAs. Importantly, this specificity indicates that pheromone does not antagonize VAs by indirectly enhancing locomotion and transmitter release. However, it is puzzling how pheromone does this. Two explanations are reasonable. One possibility is that pheromone actually binds to VA targets and directly inhibits the binding of VAs without altering the normal function of the VA target. While certainly appealing, this explanation seems unlikely, given that the Pir is abolished by goa-1 null mutations, which themselves are not halothane resistant; therefore, GOA-1 cannot be the primary target for halothane. For pheromone to be a direct antagonist, one would have to postulate that the binding of pheromone is dependent on the activity of goa-1. The fact that mutants that disrupt sensory neurons are Pir defective would suggest that the pheromone acts primarily there, whereas motor neurons are the most likely cellular site for VA effects against locomotion and cholinergic neurotransmission (VAN SWINDEREN et al. 1999 ). We have previously shown that the high-level halothane and isoflurane resistance produced by unc-64(md130) is partially diminished by a goa-1(null) mutant (VAN SWINDEREN et al. 2001 ). However, pheromone-induced resistance is additive to md130's. Like the goa-1(null); unc-64(md130) data, this result shows that the resistance activity of the md130 product can be modulated. These data also show that pheromone does not simply inhibit goa-1 function [i.e., pheromone does not phenocopy goa-1(null)]. Our current working hypothesis is that pheromone, Go, and Gq signaling alter the structure (perhaps by changing the phosphorylation state) or abundance of a protein or proteins to which VAs and the dominant-negative truncated syntaxin bind. Investigations are underway to identify this VA target.

	ACKNOWLEDGMENTS

We thank the Caenorhabditis Genetics Center, funded by the National Institutes of Health (NIH)–National Center for Research Resources, and members of the C. elegans community, whose work is referenced herein, for providing many of the strains tested. This work was supported by NIH grants RO1GM-55832 and RO1GM-059781 to C.M.C. from the National Institute of General Medical Sciences.

Manuscript received December 28, 2001; Accepted for publication February 7, 2002.

	LITERATURE CITED

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION LITERATURE CITED

ALBERT, P. S., S. J. BROWN, and D. L. RIDDLE, 1981  Sensory control of dauer larva formation in Caenorhabditis elegans.. J. Comp. Neurol. 198:435-451[Medline].

BAMBER, B. A., A. A. BEG, R. E. TWYMAN, and E. M. JORGENSEN, 1999  The Caenorhabditis elegans unc-49 locus encodes multiple subunits of a heteromultimeric GABA receptor. J. Neurosci. 19:5348-5359[Abstract/Free Full Text].

BARGMANN, C. I. and H. R. HORVITZ, 1991  Control of larval development by chemosensory neurons in Caenorhabditis elegans. Science 251:1243-1246[Abstract/Free Full Text].

BIRNBY, D. A., E. M. LINK, J. J. VOWELS, H. TIAN, and P. L. COLACURCIO et al., 2000  A transmembrane guanylyl cyclase (DAF-11) and Hsp90 (DAF-21) regulate a common set of chemosensory behaviors in Caenorhabditis elegans. Genetics 155:85-104[Abstract/Free Full Text].

BRENNER, S., 1974  The genetics of Caenorhabditis elegans. Genetics 77:71-94[Abstract/Free Full Text].

CHALFIE, M., J. E. SULSTON, J. G. WHITE, E. SOUTHGATE, and J. N. THOMSON et al., 1985  The neural circuit for touch sensitivity in Caenorhabditis elegans. J. Neurosci. 5:956-964[Abstract].

CHASE, D. L., G. A. PATIKOGLOU, and M. R. KOELLE, 2001  Two RGS proteins that inhibit Galpha(o) and Galpha(q) signaling in C. elegans neurons require a Gbeta(5)-like subunit for function. Curr. Biol. 11:222-231[Medline].

CROWDER, C. M., L. D. SHEBESTER, and T. SCHEDL, 1996  Behavioral effects of volatile anesthetics in Caenorhabditis elegans. Anesthesiology 85:901-912[Medline].

DAVIS, K. and J. DAVEY, 1997  G-protein-coupled receptors for peptide hormones in yeast. Biochem. Soc. Trans. 25:1015-1021[Medline].

DELEAN, A., P. J. MUNSON, and D. RODBARD, 1978  Simultaneous analysis of families of sigmoidal curves: application to bioassay, radioligand assay, and physiological dose-response curves. Am. J. Physiol. 235:E97-102[Abstract/Free Full Text].

FRANKS, N. P. and W. R. LIEB, 1994  Molecular and cellular mechanisms of general anaesthesia. Nature 367:607-614[Medline].

FRANKS, N. P. and W. R. LIEB, 1998  Which molecular targets are most relevant to general anaesthesia? Toxicol. Lett. 100–101:1-8.

GAMO, S., K. DODO, H. MATAKATSU, and Y. TANAKA, 1998  Molecular genetical analysis of Drosophila ether sensitive mutants. Toxicol. Lett. 100–101:329-337.

GOLDEN, J. W. and D. L. RIDDLE, 1982  A pheromone influences larval development in the nematode Caenorhabditis elegans. Science 218:578-580[Abstract/Free Full Text].

GOLDEN, J. W. and D. L. RIDDLE, 1984  The Caenorhabditis elegans dauer larva: developmental effects of pheromone, food, and temperature. Dev. Biol. 102:368-378[Medline].

GOLDEN, J. W. and D. L. RIDDLE, 1985  A gene affecting production of the Caenorhabditis elegans dauer-inducing pheromone. Mol. Gen. Genet. 198:534-536[Medline].

HAJDU-CRONIN, Y. M., W. J. CHEN, G. PATIKOGLOU, M. R. KOELLE, and P. W. STERNBERG, 1999  Antagonism between G(o)alpha and G(q)alpha in Caenorhabditis elegans: the RGS protein EAT-16 is necessary for G(o)alpha signaling and regulates G(q)alpha activity. Genes Dev. 13:1780-1793[Abstract/Free Full Text].

JENKINS, A., E. P. GREENBLATT, H. J. FAULKNER, E. BERTACCINI, and A. LIGHT et al., 2001  Evidence for a common binding cavity for three general anesthetics within the GABAA receptor. J. Neurosci. 21:RC136.

JONES, M. V., P. A. BROOKS, and N. L. HARRISON, 1992  Enhancement of gamma-aminobutyric acid-activated Cl-currents in cultured rat hippocampal neurones by three volatile anaesthetics. J. Physiol. 449:279-293[Abstract/Free Full Text].

KOELLE, M. R. and H. R. HORVITZ, 1996  EGL-10 regulates G protein signaling in the C. elegans nervous system and shares a conserved domain with many mammalian proteins. Cell 84:115-125[Medline].

KOLTCHINE, V. V., S. E. FINN, A. JENKINS, N. NIKOLAEVA, and A. LIN et al., 1999  Agonist gating and isoflurane potentiation in the human gamma-aminobutyric acid type A receptor determined by the volume of a second transmembrane domain residue. Mol. Pharmacol. 56:1087-1093[Abstract/Free Full Text].

KRISHNAN, K. S. and H. A. NASH, 1990  A genetic study of the anesthetic response: mutants of Drosophila melanogaster altered in sensitivity to halothane. Proc. Natl. Acad. Sci. USA 87:8632-8636[Abstract/Free Full Text].

KULLMANN, D. M., R. L. MARTIN, and S. J. REDMAN, 1989  Reduction by general anaesthetics of group Ia excitatory postsynaptic potentials and currents in the cat spinal cord. J. Physiol. 412:277-296[Abstract/Free Full Text].

LACKNER, M. R., S. J. NURRISH, and J. M. KAPLAN, 1999  Facilitation of synaptic transmission by EGL-30 Gqalpha and EGL-8 PLCbeta: DAG binding to UNC-13 is required to stimulate acetylcholine release. Neuron 24:335-346[Medline].

LEBERER, E., D. Y. THOMAS, and M. WHITEWAY, 1997  Pheromone signalling and polarized morphogenesis in yeast. Curr. Opin. Genet. Dev. 7:59-66[Medline].

LEIBOVITCH, B. A., D. B. CAMPBELL, K. S. KRISHNAN, and H. A. NASH, 1995  Mutations that affect ion channels change the sensitivity of Drosophila melanogaster to volatile anesthetics. J. Neurogenet. 10:1-13[Medline].

LOER, C. M. and C. J. KENYON, 1993  Serotonin-deficient mutants and male mating behavior in the nematode Caenorhabditis elegans. J. Neurosci. 13:5407-5417[Abstract].

MACIVER, M. B., A. A. MIKULEC, S. M. AMAGASU, and F. A. MONROE, 1996  Volatile anesthetics depress glutamate transmission via presynaptic actions. Anesthesiology 85:823-834[Medline].

MCINTIRE, S. L., E. JORGENSEN, and H. R. HORVITZ, 1993  Genes required for GABA function in Caenorhabditis elegans. Nature 364:334-337[Medline].

MENDEL, J. E., H. C. KORSWAGEN, K. S. LIU, Y. M. HAJDU-CRONIN, and M. I. SIMON et al., 1995  Participation of the protein Go in multiple aspects of behavior in C. elegans. Science 267:1652-1655[Abstract/Free Full Text].

MIAO, N., M. J. FRAZER, and C. LYNCH, III, 1995  Volatile anesthetics depress Ca²⁺ transients and glutamate release in isolated cerebral synaptosomes. Anesthesiology 83:593-603[Medline].

MIHIC, S. J., Q. YE, M. J. WICK, V. V. KOLTCHINE, and M. D. KRASOWSKI et al., 1997  Sites of alcohol and volatile anaesthetic action on GABA(A) and glycine receptors. Nature 389:385-389[Medline].

MILLER, K. G., M. D. EMERSON, and J. B. RAND, 1999  Goalpha and diacylglycerol kinase negatively regulate the Gqalpha pathway in C. elegans. Neuron 24:323-333[Medline].

NISHIKAWA, K. and Y. KIDOKORO, 1999  Halothane presynaptically depresses synaptic transmission in wild-type Drosophila larvae but not in halothane-resistant (har) mutants. Anesthesiology 90:1691-1697[Medline].

NISHIKAWA, K. and M. B. MACIVER, 2000  Excitatory synaptic transmission mediated by NMDA receptors is more sensitive to isoflurane than are non-NMDA receptor-mediated responses. Anesthesiology 92:228-236[Medline].

NONET, M. L., O. SAIFEE, H. ZHAO, J. B. RAND, and L. WEI, 1998  Synaptic transmission deficits in Caenorhabditis elegans synaptobrevin mutants. J. Neurosci. 18:70-80[Abstract/Free Full Text].

NURRISH, S., L. SEGALAT, and J. M. KAPLAN, 1999  Serotonin inhibition of synaptic transmission: Galpha(0) decreases the abundance of UNC-13 at release sites. Neuron 24:231-242[Medline].

PATTERSON, G. I. and R. W. PADGETT, 2000  TGF beta-related pathways. Roles in Caenorhabditis elegans development. Trends Genet. 16:27-33[Medline].

PEROUANSKY, M., D. BARANOV, M. SALMAN, and Y. YAARI, 1995  Effects of halothane on glutamate receptor-mediated excitatory postsynaptic currents. Anesthesiology 83:109-119[Medline].

RAND, J. B., and M. L. NONET, 1997 Synaptic transmission, pp. 611–643 in C. elegans II, edited by D. L. RIDDLE, T. BLUMENTHAL, B. J. MEYER and J. R. PRIESS. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.

RICHMOND, J. E. and E. M. JORGENSEN, 1999  One GABA and two acetylcholine receptors function at the C. elegans neuromuscular junction. Nat. Neurosci. 2:791-797[Medline].

RIDDLE, D. L., and P. S. ALBERT, 1997 Genetic and environmental regulation of dauer larva development, pp. 739–768 in C. elegans II, edited by D. L. RIDDLE, T. BLUMENTHAL, B. J. MEYER and J. R. PRIESS. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.

ROBATZEK, M., T. NIACARIS, K. STEGER, L. AVERY, and J. H. THOMAS, 2001  eat-11 encodes GPB-2, a Gbeta(5) ortholog that interacts with G(o)alpha and G(q)alpha to regulate C. elegans behavior. Curr. Biol. 11:288-293[Medline].

SCHLAME, M. and H. C. HEMMINGS, JR., 1995  Inhibition by volatile anesthetics of endogenous glutamate release from synaptosomes by a presynaptic mechanism. Anesthesiology 82:1406-1416[Medline].

SEGALAT, L., D. A. ELKES, and J. M. KAPLAN, 1995  Modulation of serotonin-controlled behaviors by Go in Caenorhabditis elegans. Science 267:1648-1651[Abstract/Free Full Text].

SONG, J. and H. G. DOHLMAN, 1996  Partial constitutive activation of pheromone responses by a palmitoylation-site mutant of a G protein alpha subunit in yeast. Biochemistry 35:14806-14817[Medline].

TAKENOSHITA, M. and T. TAKAHASHI, 1987  Mechanisms of halothane action on synaptic transmission in motoneurons of the newborn rat spinal cord in vitro. Brain Res. 402:303-310[Medline].

THOMAS, J. H., D. A. BIRNBY, and J. J. VOWELS, 1993  Evidence for parallel processing of sensory information controlling dauer formation in Caenorhabditis elegans. Genetics 134:1105-1117[Abstract].

TINKLENBERG, J. A., I. S. SEGAL, T. Z. GUO, and M. MAZE, 1991  Analysis of anesthetic action on the potassium channels of the Shaker mutant of Drosophila. Ann. NY Acad. Sci. 625:532-539[Medline].

VAN DER LINDEN, A. M., F. SIMMER, E. CUPPEN, and R. H. PLASTERK, 2001  The G-protein ß-subunit GPB-2 in Caenorhabditis elegans regulates the G(o)-G(q) signaling network through interactions with the regulator of G-protein signaling proteins EGL-10 and EAT-16. Genetics 158:221-235[Abstract/Free Full Text].

VAN SWINDEREN, B., D. R. SHOOK, R. H. EBERT, V. A. CHERKASOVA, and T. E. JOHNSON et al., 1997  Quantitative trait loci controlling halothane sensitivity in Caenorhabditis elegans. Proc. Natl. Acad. Sci. USA 94:8232-8237[Abstract/Free Full Text].

VAN SWINDEREN, B., O. SAIFEE, L. SHEBESTER, R. ROBERSON, and M. L. NONET et al., 1999  A neomorphic syntaxin mutation blocks volatile-anesthetic action in Caenorhabditis elegans. Proc. Natl. Acad. Sci. USA 96:2479-2484[Abstract/Free Full Text].

VAN SWINDEREN, B., L. B. METZ, L. D. SHEBESTER, J. E. MENDEL, and P. W. STERNBERG et al., 2001  Go regulates volatile anesthetic action in Caenorhabditis elegans.. Genetics 158:643-655[Abstract/Free Full Text].

WAUD, D. R., 1972  On biological assays involving quantal responses. J. Pharmacol. Exp. Ther. 183:577-607[Abstract/Free Full Text].

WHITE, J., E. SOUTHGATE, J. THOMSON, and S. BRENNER, 1986  The structure of the nervous system of Caenorhabditis elegans.. Philos. Trans. R. Soc. Lond. B Biol. Sci. 314:1-340.

ZORYCHTA, E. and R. CAPEK, 1978  Depression of spinal monosynaptic transmission by diethyl ether: quantal analysis of unitary synaptic potentials. J. Pharmacol. Exp. Ther. 207:825-836[Abstract/Free Full Text].

This article has been cited by other articles:

				B. T. Carroll, G. R. Dubyak, M. M. Sedensky, and P. G. Morgan Sulfated Signal from ASJ Sensory Neurons Modulates Stomatin-dependent Coordination in Caenorhabditis elegans J. Biol. Chem., November 24, 2006; 281(47): 35989 - 35996. [Abstract] [Full Text] [PDF]

				A. H. Hawasli, O. Saifee, C. Liu, M. L. Nonet, and C. M. Crowder Resistance to Volatile Anesthetics by Mutations Enhancing Excitatory Neurotransmitter Release in Caenorhabditis elegans Genetics, October 1, 2004; 168(2): 831 - 843. [Abstract] [Full Text] [PDF]

				P. Nagele, L. B. Metz, and C. M. Crowder Nitrous oxide (N2O) requires the N-methyl-D-aspartate receptor for its action in Caenorhabditis elegans PNAS, June 8, 2004; 101(23): 8791 - 8796. [Abstract] [Full Text] [PDF]

				K. M. Nolan, T. R. Sarafi-Reinach, J. G. Horne, A. M. Saffer, and P. Sengupta The DAF-7 TGF-beta signaling pathway regulates chemosensory receptor gene expression in C. elegans Genes & Dev., December 1, 2002; 16(23): 3061 - 3073. [Abstract] [Full Text] [PDF]