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. Author manuscript; available in PMC: 2012 Nov 11.
Published in final edited form as: Cell. 2011 Nov 11;147(4):922–933. doi: 10.1016/j.cell.2011.08.053

The neural circuits and synaptic mechanisms underlying motor initiation in C. elegans

Beverly J Piggott 1,2,4, Jie Liu 1,4, Zhaoyang Feng 3,4, Seth A Wescott 1, X Z Shawn Xu 1,2
PMCID: PMC3233480  NIHMSID: NIHMS333420  PMID: 22078887

Summary

C. elegans is widely used to dissect how neural circuits and genes generate behavior. During locomotion, worms initiate backward movement to change locomotion direction spontaneously or in response to sensory cues; however, the underlying neural circuits are not well defined. We applied a multidisciplinary approach to map neural circuits in freely-behaving worms by integrating functional imaging, optogenetic interrogation, genetic manipulation, laser ablation, and electrophysiology. We found that a disinhibitory circuit and a stimulatory circuit together promote the initiation of backward movement, and that circuitry dynamics is differentially regulated by sensory cues. Both circuits require glutamatergic transmission but depend on distinct glutamate receptors. This dual mode of motor initiation control is found in mammals, suggesting that distantly related organisms with anatomically distinct nervous systems may adopt similar strategies for motor control. Additionally, our studies illustrate how a multidisciplinary approach facilitates dissection of circuit and synaptic mechanisms underlying behavior in a genetic model organism.

Introduction

One of the ultimate goals of neuroscience research is to understand how neural circuits and genes generate behavior. Despite the great diversity of their overall anatomy, the basic building blocks of the nervous systems (i.e. structural motifs/modules of neural networks) display similarity across phylogeny (Reigl et al., 2004; Sporns and Kotter, 2004). As such, genetically tractable organisms have emerged as promising models to decode the neural and genetic basis of behavior (de Bono and Maricq, 2005).

The nematode C. elegans possesses complex behaviors ranging from motor, sensory, mating, social, sleep and drug-dependence behaviors to learning and memory (de Bono and Bargmann, 1998; de Bono and Maricq, 2005; Feng et al., 2006; Liu and Sternberg, 1995; Mori and Ohshima, 1995; Raizen et al., 2008). Interestingly, such a complex array of C. elegans behaviors, some of which were once thought to be present only in higher organisms, are mediated by a surprisingly small nervous system with merely 302 neurons and ~7,000 synapses (White et al., 1986). C. elegans also represents the only organism whose entire nervous system has been completely reconstructed by electron microscopy (EM) (White et al., 1986). These features in conjunction with its amenability to genetic manipulation make C. elegans an attractive model for decoding the neural and genetic basis of behavior. However, even for such a simple model organism as C. elegans, it remains largely mysterious as to how the nervous system is functionally organized to generate behaviors.

One of the most prominent behaviors in C. elegans is its locomotion behavior (de Bono and Maricq, 2005). Locomotion forms the foundation of most, if not all, C. elegans behaviors (e.g. sensory, social, mating, sleep and drug-dependent behaviors, and learning and memory), as these behaviors all involve locomotion and are, to varying degrees, manifested at the locomotion level. During locomotion, worms often initiate backward movement (i.e. reversals) to change the direction of locomotion either spontaneously or in response to sensory cues (de Bono and Maricq, 2005). Previous work from a number of labs has identified several key components in the neural circuitry that controls the initiation of reversals (Alkema et al., 2005; Gray et al., 2005; Hart et al., 1995; Kaplan and Horvitz, 1993; Maricq et al., 1995; Zheng et al., 1999). In particular, a group of command interneurons (AVA, AVD and AVE) were found to be essential for the initiation of reversals, as laser ablation of the precursors to both AVA and AVD rendered worms incapable of moving backward (Chalfie et al., 1985). Based on the structural map, these command interneurons receive inputs directly from sensory neurons and also from upstream intereneurons (1st and 2nd layer interneurons), and send outputs to ventral cord motor neurons (A/AS type) that drive reversals (Chalfie et al., 1985; White et al., 1986). Activation of sensory neurons by sensory cues would directly or indirectly excite these command interneurons, leading to the initiation of reversals (de Bono and Maricq, 2005). This constitutes a feed-forward stimulatory circuit (Figure 1A). However, it is not clear whether this circuit, though widely accepted, truly accounts for all of the reversal events seen in this organism.

Figure 1. The current model of the locomotion circuitry that controls the initiation of backward movement.

Figure 1

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(A) In this model, the command interneurons AVA/AVD/AVE receive input from sensory neurons and interneurons (1st layer: AIB/AIA/AIY/AIZ; 2nd layer: RIM/RIA/RIB) and directly synapse onto downstream motor neurons (A/AS, not drawn) to drive backward locomotion.

(B) A schematic drawing of the CARIBN system that enables simultaneous imaging of neuronal activity and behavioral states in freely-behaving worms.

See also Figure S1.

In this study, we applied a multidisciplinary approach to map neural circuits in freely-behaving animals. Using this approach, we interrogated the locomotion circuitry and found that our current view on the circuitry needs to be significantly revised. We identified a disinhibitory circuit acting in concert with the command interneuron-dependent stimulatory circuit to control the initiation of reversals. Interestingly, the activity patterns of these two circuits are differentially regulated by sensory cues. Notably, such a dual mode of motor initiation control has also been identified in mammals, suggesting that morphologically distinct nervous systems from distantly related organisms may adopt similar strategies to control motor output. Our study also highlights the value of applying a multidisciplinary approach to dissect the neural and genetic basis of behavior.

Results

Role of command interneurons in the initiation of reversals during spontaneous locomotion

The current model is that the command interneurons AVA, AVD and AVE, particularly AVA, mediate the initiation of reversals (Figure 1A). As a first step, we imaged the calcium activity of AVA during spontaneous locomotion by expressing in AVA a transgene encoding G-CaMP3.0, a genetically-encoded calcium sensor (Tian et al., 2009). DsRed was co-expressed with G-CaMP3.0 to enable ratiometric imaging. To reliably correlate behavior and neuronal activity, we developed an automated calcium imaging system that allows simultaneous imaging of behavior and neuronal calcium transients in freely-behaving animals (Figure 1B and S1). We name it CARIBN (Calcium Ratiometric Imaging of Behaving Nematodes) system.

We used the CARIBN system to perform imaging experiments on worms moving on the surface of an NGM (nematode growth media) plate in an open environment without any physical restraint, which is the standard laboratory condition under which nearly all behavioral analyses in C. elegans are conducted. Consistent with previous results obtained with a similar system (Ben Arous et al., 2010), we found that AVA exhibited an increase in calcium level during reversals (Figure 2A–B), indicating that AVA is involved in controlling backward movement during spontaneous locomotion.

Figure 2. The RIM neuron acts to inhibit the initiation of backward locomotion and relieving such inhibition triggers backward locomotion.

Figure 2

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(A) AVA exhibits an increase in calcium level during spontaneous reversals. The bar on top of the trace denotes the time window during which the worm underwent backward movement.

(B) Peak percentage change in the ratio of G-CaMP/DsRed fluorescence in AVA during reversals (n=40). Control: transgenic worms expressing YFP and DsRed under the same promoter.

(C) Laser of ablation of AVA, AVD, AVE and RIM. n≥5.

(D) RIM is inhibited during reversals.

(E) Peak percentage change in the ratio of G-CaMP/DsRed fluorescence in RIM during reversals (n=37).

(F) Inhibition of RIM by NpHR triggers reversals. NpHR was expressed as a transgene specifically in RIM. Control worms (transgene-free siblings) showed a basal level of spontaneous reversals. **p<0.0001 (t test). n=10.

(G) Inhibition of RIM by NpHR triggers reversals by turning on a parallel pathway. ChR2 was expressed as a transgene specifically in RIM and was turned on with a flash of blue light (2.5–5 mW/mm2). n≥5. **p<0.0001 (ANOVA with the Bonferroni test). All error bars: SEM.

See also Supplemental movie 1.

Command interneurons are not essential for in the initiation of reversals

To further evaluate the role of the command interneurons AVA/AVD/AVE in reversal initiation, we ablated these neurons individually and in combination. While worms lacking AVA exhibited a reduced reversal frequency, ablation of AVD or AVE did not result in a notable defect in reversal frequency (Figure 2C), consistent with the view that AVA plays a more important role in triggering reversals than do AVD and AVE (Gray et al., 2005; Zheng et al., 1999). Surprisingly, worms lacking AVA, AVD and AVE all together can still efficiently initiate reversals, albeit at a reduced frequency (Figure 2C and Suppl. movie 1). These results demonstrate that while the command interneurons AVA/AVD/AVE are important for initiating reversals, they are not essential for this motor program. Thus, there must be some unknown circuits that act in parallel to the command interneuron-mediated circuit to regulate the initiation of reversals during locomotion.

RIM inhibits the initiation of reversals and its activity is suppressed during reversals

To identify such circuits, we first examined the wiring pattern of the worm nervous system. RIM, RIA and RIB are classified as the “second-layer” interneurons that are suggested to act upstream of the command interneurons in the locomotion circuitry (Figure 1A) (Gray et al., 2005). In particular, the inter/motor neuron RIM sits at a unique position. It receives input from a number of interneurons and also sends output to downstream head motor neurons and neck muscles (White et al., 1986). Importantly, consistent with previous reports (Alkema et al., 2005; Gray et al., 2005; Zheng et al., 1999), laser ablation of RIM greatly increased reversal frequency (Figure 2C). This suggests that RIM inhibits the initiation of reversals during locomotion. By contrast, laser ablation of RIA and RIB do not show a significant effect on reversal frequency during spontaneous locomotion ((Gray et al., 2005) and data not shown), though these neurons regulate certain sensory behaviors (Mori and Ohshima, 1995).

We therefore imaged the activity of RIM during spontaneous locomotion using the CARIBN system. If RIM suppresses the initiation of reversals as suggested above, one would predict that each reversal event should be accompanied by a down-regulation of RIM activity. Indeed, RIM activity was down-regulated during reversals (Figure 2D–E). This result is consistent with the model that RIM inhibits reversal initiation, implying that relieving such inhibition by suppressing RIM activity should trigger reversals.

Suppression of RIM activity can initiate reversals independently of AVA/AVD/AVE

To test this, we took an optogenetic approach by expressing halorhodopsin (NpHR) as a transgene specifically in RIM. NpHR is a light-gated chloride pump, and its activation by light suppresses neuronal activity (Zhang et al., 2007). Inhibition of RIM by NpHR effectively triggered reversals in freely-moving worms (Figure 2F), suggesting that RIM tonically suppresses reversals during locomotion and relieving such suppression triggers reversals.

To ascertain whether the role of RIM in reversal initiation depends on the command interneurons AVA/AVD/AVE, we checked worms lacking AVA/AVD/AVE. Inhibition of RIM by NpHR can still initiate reversals in AVA/AVD/AVE-ablated worms (Figure 2G). Thus, suppression of RIM activity can trigger reversals independently of the AVA/AVD/AVE-mediated stimulatory circuit. This finding reveals the presence of an RIM-mediated parallel circuit in promoting reversals.

As a control, we performed the converse experiment. If inhibition of RIM can turn on the parallel circuit, stimulation of RIM should not. To test this, we expressed ChR2, a light-gated cation channel (Boyden et al., 2005; Nagel et al., 2005), as a transgene specifically in RIM. To specifically interrogate the role of the parallel circuit, we killed AVA/AVD/AVE to eliminate the stimulatory circuit as it could be artificially turned on by its connections with RIM (Guo et al., 2009). In these worms, stimulation of RIM by ChR2 cannot trigger reversals (Figure 2G). This is in sharp contrast to the observation that inhibition of RIM by NpHR can trigger reversals in the same type of worms (Figure 2G). Thus, RIM inhibition, rather than stimulation, can turn on the parallel circuit to initiate reversals.

Collectively, the above data suggest that RIM acts in a circuit in parallel to the command interneurons AVA/AVD/AVE to tonically suppress reversals during forward movement, and inhibition of RIM relieves such suppression, leading to reversal initiation.

AIB acts upstream of RIM to trigger reversals

We next asked which neurons act upstream of RIM to initiate reversals. The wiring map of C. elegans nervous system reveals that though over a dozen of neurons synapse onto RIM, most of them merely form sparse connections with RIM. Among them, AIB is quite unique in that it is a first-layer interneuron and forms unusually dense synaptic connections with RIM by sending over 30 synapses to RIM (Wormatlas.org and (White et al., 1986)). In addition, AIB regulates reversals in olfactory behavior (Chalasani et al., 2007). Laser ablation of AIB suppressed the reversal frequency to a level similar to that of AVA/AVD/AVE-ablated worms (Figure 3I). These observations raise the possibility that AIB may regulate reversal initiation by modulating RIM activity. We thus imaged AIB activity during reversals using the CARIBN system. AIB activity increased during reversals (Figure 3A–B), suggesting a role for AIB in promoting the initiation of reversals during spontaneous locomotion.

Figure 3. The AIB neuron promotes the initiation of backward locomotion by inhibiting the activity of RIM.

Figure 3

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(A) AIB fires during reversals. G-CaMP and DsRed were co-expressed as a transgene specifically in AIB.

(B) Peak percentage ratio change in G-CaMP/DsRed fluorescence in AIB during reversals (n=21). Control worms express YFP and DsRed under the same promoter.

(C) Stimulation of AIB by ChR2 triggers reversals. A flash of blue light was used to trigger reversals in worms expressing ChR2 as a transgene specifically in AIB. Control worms (transgene-free siblings) showed a basal level of spontaneous reversals. **p<0.0001 (t test). n=10.

(D) AIB acts upstream of RIM to promote the initiation of reversals. As RIM suppresses reversals, worms lacking RIM showed a higher basal level of spontaneous reversals. **p<0.0001 (t test). n≥7.

(E) AIB triggers reversals in an AVA/AVD/AVE-independent manner. **p<0.0001 (t test). n≥9.

(F–G) Calcium imaging of RIM shows that RIM is inhibited by stimulation of AIB. The dotted lines in the traces represent those few missing frames with low image quality, which are refractory to image processing.

(H) Peak percentage change in RIM calcium level in response to AIB stimulation by ChR2. n≥6.

(I) Simultaneous ablation of AVA/AVD/AVE and AIB abolished nearly all reversal events during spontaneous locomotion. AVA/AVD/AVE data is a duplicate from figure 2C. *p<0.03, **p<0.0001 (ANOVA with the Bonferroni test). n≥5. All error bars: SEM.

If AIB promotes reversal initiation, then stimulating AIB should trigger reversals. To test this, we expressed ChR2 as a transgene specifically in AIB. Stimulation of AIB by ChR2 effectively triggered reversals, providing further evidence for a role of AIB in promoting reversal initiation (Figure 3C).

The fact that AIB extensively synapses onto RIM suggests that AIB may act through RIM to promote the initiation of reversals. However, AIB also makes synaptic connections with other neurons, including AVA (White et al., 1986). Thus, the possibility that AIB acts through AVA rather than RIM to promote reversals cannot be ruled out. We thus repeated the ChR2 experiments on RIM-ablated worms and found that stimulation of AIB by ChR2 can no longer further stimulate reversals in these worms (Figure 3D). By contrast, worms with AVA/AVD/AVE ablated still initiated reversals in response to AIB stimulation by ChR2 (Figure 3E). These results suggest that under this condition, AIB acts through the RIM-dependent parallel circuit, rather than the AVA/AVD/AVE-dependent stimulatory circuit, to promote the initiation of reversals.

AIB triggers reversals by inhibiting RIM

We considered that AIB may inhibit RIM to trigger reversals. This model predicts that stimulation of AIB should result in inhibition of RIM. To test this, we recorded the activity of RIM in response to AIB stimulation by ChR2. Although optogenetics has been applied to stimulate neurons in freely-behaving worms (Leifer et al., 2011; Stirman et al., 2011), it has not been possible to simultaneously record neuronal activity in the same animal. The CARIBN system allows us to stimulate one neuron by optogenetics while recording the activity of another neuron on freely-behaving animals. Specifically, the blue light used to image G-CaMP calcium signals in RIM can also turn on ChR2 expressed in AIB, making it possible to image the activity of RIM in response to stimulation of AIB on freely-behaving worms. Upon light stimulation, RIM exhibited a sharp decrease in calcium level (Figure 3F–H). As predicted, worms initiated reversals (Figure 3F). The decrease in RIM activity depended on AIB stimulation, as no such response was observed in worms lacking the ChR2 transgene in AIB (Figure 3G–H). This data, together with the results from electrophysiological recordings (see below), strongly suggests that AIB triggers reversals by inhibiting RIM activity.

Taken together, our results suggest a model in which AIB acts upstream to inhibit RIM, an inter/motor neuron that tonically inhibits reversals during locomotion; activation of AIB suppresses RIM activity, which in turn relieves the inhibitory effect of RIM on backward movement, thereby triggering reversals. In other words, backward locomotion inhibited by RIM can be “disinhibited” by AIB. This would constitute a disinhibitory circuit that promotes the initiation of reversals (Figure 7I).

Figure 7. Electrophysiological characterization of the AIB-RIM synapse of the disinhibitory circuit in response to nose touch.

Figure 7

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(A–B) RIM is hyperpolarized in response to nose touch, which depends on eat-4. n≥9. Clamping current: 0 pA.

(C–D) AIB stimulation by ChR2 leads to inhibition of RIM. See AIB traces in figure S3A. n≥6. Clamping current: 0 pA.

(E–F) Glutamate (1mM) perfusion evokes a hyperpolarizing outward current in RIM, which was absent in avr-14(ad1302) mutant worms. Voltage clamp: 0 mV. The tiny inward current in avr-14(ad1302) mutant was carried by an unknown glutamate-gated cation channel whose activity was small and masked by the predominant anion channel AVR-14 in wild-type worms. n≥6.

(G) Glutamate-gated currents are carried by a Cl− channel. n=5.

(H) No IPSP signal was detected in RIM of avr-14(ad1302) mutant worms in response to nose touch. Clamping current: 0 pA.

(I) A schematic model illustrating the disinhibitory and stimulatory circuits. The dotted arrows in red indicate crosstalk between the two circuits. AIB, if over-stimulated by ChR2 (with >10× brighter blue light), also sends output to AVA (BJP, JL and XZSX, unpublished observation). The dotted arrows in black indicate that other unknown sensory neurons and interneurons may regulate the two circuits by sending output to AVA, AIB, and RIM.

All error bars in this figure: SEM. See also Figure S3.

The disinhibitory and stimulatory circuits together form the primary pathways promoting reversal initiation during spontaneous locomotion

Is this disinhibitory circuit important for the initiation of reversals during spontaneous locomotion? If so, then simultaneous elimination of both the disinhibitory and stimulatory circuits should result in a severe defect in reversal initiation. Indeed, while ablation of AVA/AVD/AVE or AIB only reduced reversal frequency, ablation of AVA/AVD/AVE and AIB together abolished nearly all reversal events during spontaneous locomotion (Figure 3I). These results suggest that the AIB-RIM-dependent disinhibitory circuit and the command interneurons AVA/AVD/AVE-dependent stimulatory circuit together form the primary pathways to control reversal initiation during spontaneous locomotion.

Both the disinhibitory and stimulatory circuits are recruited to promote the initiation of reversals in response to nose touch

We then wondered how sensory cues impinge on these two circuits. In addition to spontaneous reversals, worms initiate reversals in response to various sensory stimuli, particularly aversive cues. As a consequence, these animals are able to avoid unfavorable or hazardous environments, a behavioral response essential for their survival. We focused on nose touch behavior, one of the best characterized avoidance behaviors (Kaplan and Horvitz, 1993). In this behavior, touch delivered to the worm nose tip triggers reversals. The polymodal sensory neuron ASH is the primary sensory neuron detecting nose touch stimuli, as its ablation leads to a severe defect in nose touch behavior (Kaplan and Horvitz, 1993). In addition, nose touch can stimulate this neuron in calcium imaging assays (Hilliard et al., 2005). Notably, ASH sends synapses to both AIB and AVA (White et al., 1986), and nose touch can excite AVA in electrophysiological assays (Mellem et al., 2002). This suggests a model in which ASH may engage both the disinhibitory and stimulatory circuits in this avoidance behavior.

To test the above model, we first employed our CARIBN system to image the activity of the nose touch circuits. As this imaging system performs recording in an open environment, we were able to deliver touch stimuli directly to the nose tip of freely-moving worms while simultaneously monitoring their neuronal activities and behavioral states. Our model predicts that nose touch should stimulate AVA, but inhibit RIM via stimulating AIB. Indeed, upon nose touch, AVA showed an increase in calcium activity during reversals (Figure 4A and 4C). Similarly, nose touch also stimulated AIB during reversals (Figure 4B–C). By contrast, RIM was inhibited during reversals (Figure 4D and 4F). Importantly, in AIB-ablated worms, RIM was no longer inhibited during reversals, indicating that the inhibition of RIM requires AIB (Figure 4E–F). This is consistent with the model that sensory information flows to RIM via AIB. These observations suggest that nose touch may trigger reversals by recruiting both the disinhibitory and stimulatory circuits.

Figure 4. Worms employ both the disinhibitory and stimulatory circuits to trigger backward locomotion in nose touch avoidance behavior.

Figure 4

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(A) AVA is stimulated during reversals in nose touch behavior.

(B) AIB is stimulated during reversals in nose touch behavior. The dotted lines in the trace represent missing frames.

(C) Bar graph summarizing the data in (A) and (B). n≥11.

(D) RIM is inhibited during reversals in nose touch behavior.

(E) Inhibition of RIM depends on AIB.

(F) Bar graph summarizing the data in (D) and (E). n≥12.

(G) Simultaneous ablation of both the disinhibitory and stimulatory circuits abolished nearly all reversal events triggered by nose touch. n≥5. **p<0.0001 (ANOVA with the Bonferroni test). All error bars: SEM.

To provide additional evidence, we killed AIB, RIM and the command interneurons. Laser ablation of AIB, RIM or AVA/AVD/AVE all led to a significant reduction in reversal frequency (Figure 4G), indicating that both the disinhibitory and stimulatory circuits contribute to nose touch behavior. More importantly, simultaneous elimination of both circuits by killing AVA/AVD/AVE together with AIB or RIM virtually abolished all reversals triggered by nose touch (Figure 4G). Thus, the disinhibitory and stimulatory circuits together form the primary pathways through which worms initiate reversals to avoid nose touch cues.

The disinhibitory circuit cooperates with the stimulatory circuit to promote the initiation of reversals in response to osmotic shock

Similar to nose touch, osmotic shock delivered to the worm nose also triggers reversals by stimulating the same sensory neuron ASH (Hilliard et al., 2005). Notably, osmotic shock is known to be much more noxious than nose touch (Mellem et al., 2002), and unlike nose touch, a failure to avoid high osmolarity environment (e.g. 4M fructose) leads to death. As a result, osmotic shock suppressed head oscillations during reversals, while nose touch did not; nor was this phenomenon observed during spontaneous locomotion (Alkema et al., 2005) (Figure 5G). Suppression of head oscillations is believed to facilitate efficient escape from noxious cues such as osmotic shock, and this behavioral strategy requires stimulation of RIM (Alkema et al., 2005). As was the case with spontaneous locomotion and nose touch behavior, both AVA and AIB were stimulated by osmotic shock (Figure 5A–C); however, RIM was stimulated rather than inhibited by osmotic shock (Figure 5D and 5F), an observation distinct from that observed in the other two behaviors. This indicates that while the stimulatory circuit was clearly functional in osmotic avoidance behavior, the disinhibitory circuit was instead recruited to promote suppression of head oscillations in this behavior.

Figure 5. The role of the disinhibitory and stimulatory circuits in triggering backward locomotion in osmotic avoidance behavior.

Figure 5

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(A–C) AVA and AIB are stimulated during reversals in osmotic avoidance behavior. Stimulus: 2M glycerol. n≥11.

(D) RIM is stimulated during reversals triggered by osmotic shock.

(E) RIM is inhibited during reversals in worms lacking AVA/AVD/AVE. The dotted lines in this trace and in (A) represent missing frames.

(F) Bar graph summarizing the data in (D) and (E). n≥7.

(G) Head oscillations occur during reversals in spontaneous locomotion and nose touch behavior but are suppressed in osmotic avoidance behavior. n=5. **p<0.0001 (ANOVA).

(H) Simultaneous ablation of both the disinhibitory and stimulatory circuits abolished nearly all reversal events triggered by osmotic shock. n≥5. **p<0.0001 (ANOVA with the Bonferroni test). All error bars: SEM.

To further characterize the osmotic avoidance circuits, we performed laser ablation experiments. Worms lacking the disinhibitory circuit (AIB or RIM ablated) only exhibited a slight, but insignificant, reduction in reversal frequency in osmotic avoidance behavior (Figure 5H). As expected, worms with RIM ablated no longer suppressed head oscillations during reversals, consistent with the role of RIM in this function (Figure 5G). By contrast, worms lacking the stimulatory circuit (AVA/AVD/AVE-ablated) displayed a significant defect in osmotic avoidance behavior (Figure 5H); notably, osmotic shock can still trigger reversals in these worms, albeit at a reduced frequency, indicating that additional circuits are functional in the absence of the stimulatory circuit (Figure 5H).

We considered that the remaining reversal events in AVA/AVD/AVE-ablated worms could be mediated by the disinhibitory circuit. Indeed, in AVA/AVD/AVE-ablated worms, osmotic shock no longer stimulated RIM but instead inhibited RIM during reversals, which is similar to that observed in the other two behaviors (Figure 5E–F). This demonstrates that the disinhibitory circuit is functional in worms lacking the stimulatory circuit, suggesting that the disinhibitory circuit is responsible for the remaining avoidance response in these worms. This also suggests that the excitatory input to RIM was derived from AVA/AVD/AVE in osmotic avoidance behavior, consistent with the fact that these command interneurons form synaptic connections with RIM (White et al., 1986). Finally and importantly, simultaneous ablation of both the disinhibitory and stimulatory circuits rendered worms virtually incapable of initiating reversals in response to osmotic shock (Figure 5H). Thus, in osmotic avoidance behavior, worms employ the stimulatory circuit as the primary pathway and the disinhibitory circuit as the salvage pathway to trigger reversals; in addition, worms recruit neurons in the disinhibitory circuit to suppress head oscillations to facilitate efficient escape from high osmolarity environment. This illustrates an example in which the two circuits cooperate to promote avoidance responses to noxious stimuli. This also shows that sensory cues (nose touch vs. osmotic shock) differentially regulate the activity patterns of these two circuits.

Electrophysiological recording of the activity of the disinhibitory and stimulatory circuits

Having identified the circuits that promote reversal initiation, we then set out to investigate the synaptic mechanisms by which the circuits process information. Though our CARIBN system can record the circuit activity in freely-behaving animals, this assay is indirect, since it measures the calcium level but not the membrane excitability of a neuron, and also lacks the capacity to resolve synaptic events in the circuitry. Thus, we decided to employ electrophysiological approaches to record the circuit activity by patch clamping. The small size of worm neurons (~2 µm in diameter), however, makes this type of recordings technically challenging (Goodman et al., 1998). We focused on the nose touch circuits due to the relative ease of delivering touch stimuli with precision in whole-cell recording. This was achieved by using a glass probe driven by a piezo actuator to press the nose tip (Figure 6A).

Figure 6. Electrophysiological characterization of the ASH-AVA and ASH-AIB synapses of the stimulatory and disinhibitory circuits in response to nose touch.

Figure 6

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(A) A schematic illustrating the setting of whole-cell recording (not drawn to scale).

(B) Nose touch depolarizes the sensory neuron ASH. The miniature upward spikes represent spontaneous activity of ASH. Clamping current: 0 pA.

(C–D) AVA is depolarized in response to nose touch in wild-type but not in eat-4(ky5) and glr-1(n2461) mutants. n≥7. Clamping current: 0 pA. **p<0.005 (t test).

(E–F) AIB is depolarized in response to nose touch, which requires eat-4 and glr-1. n≥5. Clamping current: 0 pA. **p<0.005 (t test).

(G) Nose touch behavior. n=10. *p<0.02; **p<0.005 (t tests used for two-group comparisons; ANOVA with the Dunnett test used for multi-group comparisons). All error bars: SEM.

See also Figure S2.

We recorded all of the four major neurons in the two circuits: the sensory neuron ASH and the interneurons AVA, AIB and RIM (Figure 7I). We focused on recording voltage signals through current clamp, due to the high input resistance of worm neurons (typically 2–5 GΩ)(Goodman et al., 1998; Liu et al., 2010). Nose touch evoked a depolarizing voltage response in ASH (Figure 6B). Similarly, a depolarizing voltage signal (i.e. EPSP) was detected in AVA and AIB upon nose touch (Figure 6C–F). By contrast, in the RIM neuron, nose touch triggered a hyperpolarizing voltage response (i.e. IPSP) (Figure 7A–B). Finally, we directly recorded the synaptic events between AIB and RIM by stimulating AIB with ChR2 (Figure S3A–B), and then recording postsynaptic responses in RIM. AIB stimulation by ChR2 led to a hyperpolarizing response (IPSP) in RIM (Figure 7C–D). These results are well consistent with our calcium imaging data from freely-behaving animals. Thus, activation of ASH by nose touch can turn on both the disinhibitory and stimulatory circuits, providing further evidence for our model. It is worthy to note that the resting potential of RIM was around −20 mV, much higher than that of AIB (~−50 mV), indicating a more depolarized state for RIM. This is consistent with our model that RIM remains in an active state to tonically inhibit the initiation of reversals during locomotion.

The ASH-AVA and ASH-AIB synapses are glutamatergic and require an AMPA/kainate-type glutamate receptor

We first characterized the pre-synaptic mechanisms of the nose touch circuits. Initially, we focused on the ASH-AVA and ASH-AIB synapses. ASH is known to be glutamatergic, and worms deficient in glutamatergic transmission are severely defective in nose touch behavior (Mellem et al., 2002). We thus performed recordings on eat-4 mutant worms where glutamatergic transmission is deficient. eat-4 encodes a vesicular glutamate transporter (Lee et al., 1999). Nose touch-evoked EPSPs in AVA and AIB were severely defective in eat-4 mutant worms (Figure 6C–F). Furthermore, expression of wild-type eat-4 gene in ASH restored nose touch-evoked EPSPs in AVA and AIB (Figure 6D, 6F, S2A, and S2C), as well as nose touch behavioral response in eat-4 mutant worms (Figure 6G). These results support the view that the ASH-AVA and ASHAIB synapses are glutamatergic.

We then turned our attention to the post-synaptic receptors, asking which glutamate receptors are required for the EPSP responses in AVA and AIB. GLR-1 is the closest C. elegans homolog of AMPA/kainate-type glutamate receptors and has been reported as the primary excitatory glutamate receptor in AVA and AIB (Chalasani et al., 2007; Hart et al., 1995; Maricq et al., 1995; Mellem et al., 2002). Consequently, worms lacking GLR-1 are severely defective in nose touch avoidance behavior (Hart et al., 1995; Maricq et al., 1995). We recorded the activity of AVA and AVB in response to nose touch in glr-1 mutant worms. No EPSP signals could be evoked by nose touch in AVA or AIB of mutant worms (Figure 6C–F), indicating that GLR-1 is required for EPSPs in these two interneurons. Furthermore, expression of wild-type glr-1 gene in AVA or AIB restored nose touch-evoked EPSP responses in AVA or AIB of glr-1 mutant worms, respectively (Figure 6D, 6F, S2B, and S2D), as well as nose touch behavioral responses (Figure 6G). Thus, GLR-1 is an essential subunit of the postsynaptic receptors mediating EPSPs in AVA and AIB.

The AIB-RIM synapses are also glutamatergic and but require a glutamate-gated Cl− channel

Lastly, we characterized the AIB-RIM synapses. Notably, AIB also appears to be glutamatergic, as it expresses eat-4 (Ohnishi et al., 2011). As expected, nose touch can no longer trigger IPSPs in RIM of eat-4 mutant worms (Figure 6A–B). However, this can also be explained by a defect in the sensory neuron ASH as eat-4 is expressed in ASH as well. We therefore knocked down eat-4 specifically in AIB by expressing an eat-4 RNAi as a transgene specifically in AIB. RNAi of eat-4 in AIB led to a strong deficit in nose touch-evoked IPSP in RIM (Figure 7B and S3C). This RNAi treatment also resulted in a significant defect in nose touch behavior to an extent similar to that caused by AIB ablation (Figure 6G and 4G). These data suggest the AIB-RIM synapses are glutamatergic. To provide further evidence, we directly interrogated the AIB-RIM synapses by recording the activity of RIM in response to AIB stimulation by ChR2 in eat-4 mutant worms. No IPSP was detected in RIM following stimulation of AIB by ChR2 in mutant worms (Figure 7C–D), further suggesting that the AIB-RIM synapses are glutamatergic.

The question arises as to how glutamate, a well-known excitatory neurotransmitter, triggers an inhibitory response (IPSP) in RIM. In addition to glutamate-gated cation channels such as GLR-1, the C. elegans genome encodes at least half a dozen glutamate-gated Cl− channels (Yates et al., 2003). Notably, the IPSP response in RIM reversed its sign around -50 mV, close to the equilibrium potential of Cl−, suggesting that it is mediated by a Cl− channel (Figure S3D). Moreover, using a high Cl− pipette solution, we detected an EPSP rather than IPSP response in RIM (Figure S3E), further suggesting that it is carried by a Cl− channel. To provide additional evidence, we directly perfused glutamate towards RIM. Glutamate evoked a hyperpolarizing current in RIM with a reversal potential around −50 mV (Figure 7E–G). Increasing the Cl− concentration in the pipette solution shifted the reversal potential close to 0 mV (Figure 7G). These data together suggest that the IPSP response in RIM is mediated by a glutamate-gated Cl− channel.

Finally, we sought to identify the glutamate-gated Cl− channel genes required for IPSPs in RIM. We focused on the alpha subunits of glutamate-gated Cl− channels, as they can form functional channels on their own (Yates et al., 2003). Five such genes are present in the C. elegans genome, including avr-14, avr-15, glc-1, glc-3 and glc-4 (Yates et al., 2003). While avr-15, glc-1, glc-3 and glc-4 mutant worms all expressed glutamate-gated Cl− currents in RIM (Figure S3F), mutations in avr-14 abolished such currents (Figure 7E–F). As a result, nose touch can no longer evoke IPSPs in RIM of avr-14 mutant worms (Figure 7H). AVR-14 was expressed in RIM (Figure S3J), and expression of wild-type avr-14 gene in RIM rescued glutamate-gated Cl− currents (Figure 7F and S3G), as well as nose-touch evoked IPSP response in RIM (Figure S3H–I). Furthermore, AVR-14 can form a functional glutamate-gated Cl− channel in heterologous systems (Dent et al., 2000). These observations indicate that AVR-14 is an essential subunit of the postsynaptic receptor(s) mediating the glutamate-gated Cl− current underlying IPSPs in RIM.

Discussion

C. elegans has emerged as a genetic model to study motor control and sensorimotor integration (de Bono and Maricq, 2005). In this study, we interrogated the circuit and synaptic mechanisms underlying the initiation of reversals in spontaneous locomotion and some sensory behaviors by applying a multidisciplinary approach integrating calcium imaging, optogenetic interrogation, genetic manipulation, laser ablation, and electrophysiology. Performing calcium imaging and optogenetic assays on freely-behaving worms allowed us to reliably associate circuit activity with behavior. Genetic manipulation and laser ablation facilitated the interrogation of the role of individual genes and neurons in the circuitry. The use of electrophysiology enabled us to validate the circuitry and also to dissect the synaptic mechanisms by which the circuitry processes information. A combination of these approaches permits a rigorous dissection of the neural and genetic basis of behavior. To our knowledge, such a comprehensive approach has not been applied to map neural circuits underlying behavior in other organisms.

We found that our current model of C. elegans locomotion circuitry needs to be significantly revised. In particular, we showed that the command interneurons AVA/D/E, which were long believed to be essential for the initiation of reversals, are in fact not required for this motor program. Genetic ablation of these neurons and others also suggested a similar conclusion (Zheng et al., 1999). More importantly, we identified an RIM inter/motor neuron-dependent disinhibitory circuit acting in concert with the command interneuron-mediated stimulatory circuit to promote the initiation of reversals (Figure 7I). RIM may control reversal initiation by regulating the activity of its downstream motor neurons and/or muscles, and possibly the command interneurons that control forward movement (e.g. AVB and PVC). The presence of two circuits may help ensure that this critical motor program be efficiently executed, and also provide flexibility for its modulation by sensory inputs and perhaps by experience.

These two circuits apparently do not act in isolation and are regulated by sensory cues. In addition to ASH, other sensory neurons may impinge on these circuits. Other interneurons may also modulate these circuits via AVA/D/E, RIM and AIB (Figure 7I). For example, AIZ and AIY form connections with RIM and may regulate RIM activity. Finally, the two circuits may regulate each other through cross-talk as shown in osmotic avoidance behavior. It should also be noted that our data do not exclude the possibility that additional circuits may function in parallel to regulate reversals. One interesting observation is that though connected by gap junctions, the activity patterns of RIM and AVA are not synchronized in spontaneous locomotion or nose touch behavior, suggesting that these electrical synapses are dynamically regulated under different physiological contexts. Similar observations have been observed in vertebrate retinal circuits (Bloomfield and Volgyi, 2009). This presents an example in which distinct sensory inputs (nose touch vs. osmotic shock) differentially regulate the dynamics of motor circuits. Future studies will elucidate whether and how other sensory cues, sensory neurons and interneurons regulate these two circuits, how they regulate each other through cross-talk, and whether and how they are modulated by experience.

Interestingly, the disinhibitory circuit identified in this study is functionally analogous to those found in the mammalian basal ganglia that facilitate the initiation of motor programs. These circuits allow the brain to suppress competing or non-synergistic motor programs that would otherwise interfere with sensory and goal-directed behaviors (Purves et al., 2008). In the case of C. elegans, as its pharynx cannot efficiently take up surrounding bacteria (i.e. worm food) during backward locomotion, such a circuit would provide a potential mechanism for the animal to suppress reversals; in doing so, the animal would be able to spend most of its time moving forward or dwelling to facilitate feeding and only initiate reversals stochastically (spontaneous reversals) or in response to sensory cues.

Stimulatory circuits have also been widely employed by mammals to control motor initiation (Purves et al., 2008). For example, in response to painful sensory stimuli, nociceptive DRG neurons can bypass the basal ganglia and the upper motor nervous system to trigger a limb withdrawal response by directly activating the local circuitry in the spinal cord (Purves et al., 2008). This would ensure that animals can rapidly escape from painful stimuli (Purves et al., 2008). In the case of C. elegans, the disinhibitory circuit functions in spontaneous locomotion and nose touch behavior. Interestingly, when encountering more noxious stimuli (e.g. osmotic shock), worms also bypass the disinhibitory circuit and primarily depend on the stimulatory circuit to trigger reversals. Our results suggest that despite the great diversity of their anatomy, the nervous systems from distantly related organisms may adopt similar strategies to control motor output.

Concluding remarks

As the only organism with a structural map of the entire nervous system available, C. elegans has emerged as a model to dissect how genes and neural circuits generate behavior (de Bono and Maricq, 2005). Nevertheless, much of the information regarding motor circuits was inferred from the structural map and thus has not been extensively tested at the experimental level. It has become increasingly clear that a structural map of the nervous system, though highly informative, cannot be directly transcribed into a functional map (de Bono and Maricq, 2005). Apparently, an understanding of the functional map requires rigorous interrogation of the functional roles of individual neurons in the circuitry in the context of behavior and of how genes, environment and experience regulate circuit dynamics and hence behavioral output. Our study illustrates an example how a multidisciplinary approach can be employed to study these questions in a genetic model organism.

Experimental procedures

The CARIBN system and calcium imaging

As diagramed in Figure 1B, the automated CARIBN system consists of an upright microscope (Zeiss M2Bio), EMCCD camera (Andor), dual-view beamsplitter (Optical Insights), Xenon light source (Sutter), motorized stage, and computer (Dell). A C-mount (0.63×) is used to couple the camera to the beamsplitter. A dual band excitation filter (Chroma) simultaneously excites G-CaMP and DsRed at 488 nm and 560 nm, respectively. This system can be readily adapted to monitor fluorescent signals from Cameleon that has also been extensively used for imaging calcium transients in C. elegans neurons and muscles (Clark et al., 2006; Faumont and Lockery, 2006; Kerr et al., 2000). In this case, a different set of filters are needed. We used a 20× objective in conjunction with a 1.6× zoom lens to acquire images. A home-developed software package controls the system and follows fluorescent objects (neurons of the worm) in dark field by their size and brightness. Specifically, a feedback loop system is introduced to track the object (neurons of the worm) by instructing the stage to move the object to the center of the camera field (re-centering) every half second (2 Hz). Under this setting, we very rarely (<1%) lose track of the worm over a 10 min window. Images were acquired with 5–25 ms exposure time (depending on fluorescence intensity of the transgene) at up to 22 Hz without binning. To facilitate identification of neurons for ratio computation, a mask image was generated for each frame by applying the following digital filters: a spatial filter to sharpen the image by correcting the motion blur; an intensity filter and size filter to single out the neuron of interest from other neurons and the nerve ring. None of these digital filters would alter the ratio of G-CaMP/DsRed fluorescence, as the ratio computation was solely based on the raw images. Nevertheless, there are always a few frames particularly those captured during stage movement that are of poor image quality; these frames are thus not processed and marked with dotted lines in the traces. A series of digital spatial filters and morphological filters were used to selectively enhance the autofluorescence emitted from the worm body, such that the outline of the worm body (head and a portion of the anterior body) can be identified to derive behavioral parameters such as backward/forward movement, speed, and trajectory. To compute the ratio change during a reversal event, we first determined the precise starting and ending frame numbers of the reversal. The image data ~2 sec before the starting frame were used as the basal line, and the mean ratio value of this basal line was used to compute the ratio change. The first peak or trough within the reversal period was identified to calculate the ratio change.

Calcium imaging was performed on day 1 adult worms under the standard laboratory condition where worms were allowed to freely move on the surface of an NGM plate covered with a thin layer of bacteria (OP50) without any physical restraint. Nose touch stimulus was delivered as described (Kaplan and Horvitz, 1993). A small drop of 2M glycerol was placed on the path of a forward-moving worm to induce osmotic avoidance response as described (Mellem et al., 2002). OP50 was not included in the osmotic assay. A positive response was scored if the worm stopped forward movement and also initiated a reversal lasting at least half of a head swing. We only scored the reversals initiated within the first 3 sec after the animal encountered the drop. Each worm was tested five times with a ~5 min interval between each test, and a percentage score was tabulated for each worm. To image the activity of RIM in response to ChR2 stimulation by ChR2, worms were first tracked under the DsRed channel excited with yellow light and then switched to the G-CaMP/DsRed channels excited with both blue and yellow light. To control intrinsic phototaxis responses (Ward et al., 2008), imaging was performed on lite-1(xu7) worms insensitive to blue light (Liu et al., 2010).

Optogenetics

Worms grown on NGM plates supplied with 5 µM all-trans retinal were tested on retinal-free NGM plates spread with a thin layer of OP50. ChR2 experiments were carried out in lite-1(xu7) worms lacking intrinsic phototaxis responses (Liu et al., 2010). Unless otherwise indicated, a 5 sec pulse of blue (470±20 nm; 0.1–0.2 mW/mm2) or yellow light (575±25 nm; 25 mW/mm2) was delivered from an Arc lamp (EXFO) by a 10× objective (Zeiss M2Bio) to the head of a forward-moving worm to turn on ChR2 or NpHR, respectively. A positive response was scored if the worm stopped forward movement and also initiated a reversal lasting more than half of a head swing. Only the reversals initiated during the 5 sec light stimulus were scored. Each worm was tested five times with a ~5 min interval between each test, and a percentage score was tabulated for each worm. Because worms exhibit spontaneous reversals, a basal level of reversals was observed in controls. This number shows some variation, which may be contributed by temperature, humility and quality of NGM plates. As worms reared on retinal-containing plates show a slightly higher frequency of spontaneous reversals under our conditions, transgene-free siblings (rather than worms grown on retinal-free plates) were used as controls in behavioral tests.

Electrophysiology

Patch-clamp recordings were performed under an Olympus microscope (BX51WI) using an EPC-10 amplifier and the Pulse software (HEKA) as previously described (Kang et al., 2010). Briefly, we glued worms to a sylgard-coated coverglass covered with bath solution and then carefully cut a small piece of cuticle in the head to expose head neurons while keeping the nose tip intact. The animal was kept alive during recording. To preserve synaptic functions, it is important to avoid displacing neurons from their original position during dissection; otherwise, chemical synapses may get disrupted/depressed and their activity may also quickly run down (though electric synapses tend to be preserved). Blue light pulses (0.2 mW/mm2; 470±20 nm; 0.5–1 sec) were delivered from an Arc lamp (EXFO Xcite) coupled to a mechanical shutter (Sutter) triggered by the amplifier. A glass probe driven by a piezo actuator (PI) mounted on a micromanipulator was used to deliver nose touch stimuli (10 µm) towards the nose tip. The normal bath solution contains (in mM): 145 NaCl, 5 KCl, 1 CaCl2, 5 MgCl2, 11 dextrose, and 5 HEPES (330 mOsm; pH adjusted to 7.3). The pipette solution contains 115 K-gluconate, 15 KCl, 5 MgCl2, 10 HEPES, 0.25 CaCl2, 20 sucrose, 5 EGTA, 5 Na2ATP and 0.5 NaGTP. When recording nose touch- and ChR2-evoked responses, supernatant from freshly-grown OP50 culture was diluted (1:10) into the bath solution to mimic the conditions of behavioural assays and also to help prevent the run down of synaptic functions. In the high Cl− pipette solution, 115 mM K-gluconate was replaced with KCl. Cells were mostly recorded by current-clamp, and currents were clamped at 0 pA unless otherwise indicated.

Molecular genetics and laser ablation

Standard methods were used to generate plasmids and transgenes driven by cell-specific promoters. Laser ablation was also conducted using standard protocols. See supplemental information for details.

Supplementary Material

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Acknowledgments

We thank J. Gao, W. Li, and A. Ward for technical assistance; L. Looger for the G-CaMP3.0 plasmid; A. Gottschalk for ChR2 plasmid; K. Deisseroth for NpHR plasmid; J. Dent and L. Avery for avr-14 strains and plasmids; P. Hu, A. Kumar and B. Ye for comments on the manuscript. Some strains were obtained from the CGC and Knockout Consortiums in the U.S.A. and Japan. This work was supported by grants from the NIGMS and Pew scholar program (X.Z.S.X).

Footnotes

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Associated Data

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Supplementary Materials

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03
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