The C.elegans ric‐3 gene is required for maturation of nicotinic acetylcholine receptors

Sarah Halevi, Jim McKay, Mark Palfreyman, Lina Yassin, Margalit Eshel, Erik Jorgensen, Millet Treinin

Author Affiliations

  1. Sarah Halevi1,
  2. Jim McKay2,
  3. Mark Palfreyman3,
  4. Lina Yassin1,
  5. Margalit Eshel1,
  6. Erik Jorgensen3 and
  7. Millet Treinin*,1
  1. 1 Department of Physiology, Hebrew University–Hadassah Medical School, Jerusalem, 91120, Israel
  2. 2 Department of Molecular Biology, University of Texas Southwestern Medical Center, 5323 Harry Hines Blvd, Dallas, TX, 75235‐9148, USA
  3. 3 Department of Biology, University of Utah, 257 South 1400 East, Salt Lake City, UT, 84112‐0840, USA
  1. *Corresponding author. E‐mail: millet_t{at}
View Full Text


Mutations in ric‐3 (resistant to inhibitors of cholinesterase) suppress the neuronal degenerations caused by a gain of function mutation in the Caenorhabditis elegans DEG‐3 acetylcholine receptor. RIC‐3 is a novel protein with two transmembrane domains and extensive coiled‐coil domains. It is expressed in both muscles and neurons, and the protein is concentrated within the cell bodies. We demonstrate that RIC‐3 is required for the function of at least four nicotinic acetylcholine receptors. However, GABA and glutamate receptors expressed in the same cells are unaffected. In ric‐3 mutants, the DEG‐3 receptor accumulates in the cell body instead of in the cell processes. Moreover, co‐expression of ric‐3 in Xenopus laevis oocytes enhances the activity of the C.elegans DEG‐3/DES‐2 and of the rat α‐7 acetylcholine receptors. Together, these data suggest that RIC‐3 is specifically required for the maturation of acetylcholine receptors.


Ionotropic acetylcholine receptors are members of the ligand‐gated ion channel superfamily. These acetylcholine receptors are pentamers, usually heteromers, composed of specific α and non‐α subunits. These subunits have a large N‐terminal extracellular domain, four transmembrane domains, and a large intracellular loop between transmembrane domains III and IV (Numa et al., 1983). The receptor subunits fold and assemble into pentamers in the endoplasmic reticulum (ER), and are then exported through the Golgi apparatus to the plasma membrane. A strict requirement for folding and pentamer assembly prior to exit from the ER has been suggested (Smith et al., 1987; Gu et al., 1991; Kreienkamp et al., 1995; Keller et al., 2001). This maturation process is time consuming, requiring 2 h (Merlie and Lindstrom, 1983), and depends upon poorly understood cellular machinery. The low surface expression of several acetylcholine receptors when expressed in heterologous cells indicates that cell‐specific assembly and trafficking proteins may be required (Cooper and Millar, 1997; Dineley and Patrick, 2000; Sweileh et al., 2000). However, except for the broad‐specificity ER chaperones, BiP, calnexin and 14‐3‐3η, little is known about proteins that mediate folding, assembly or targeting of acetylcholine receptors (Blount and Merlie, 1991; Chang et al., 1997; Jeanclos et al., 2001).

To identify genes needed for acetylcholine receptor activity in Caenorhabditis elegans, we screened for suppressors of a dominant mutation in the acetylcholine receptor subunit DEG‐3 (Treinin and Chalfie, 1995; Treinin et al., 1998). The DEG‐3(u662) mutation is a substitution of an amino acid in the pore‐forming transmembrane domain (TMDII) of this subunit. The mutant subunit assembles with DES‐2 to form a non‐desensitizing channel that ultimately causes degeneration and necrotic death of neurons that express this gene (Treinin and Chalfie, 1995; Treinin et al., 1998). This screen led to the identification of mutations in des‐2, a subunit of the DEG‐3 receptor, and also in ric‐3 (resistant to inhibitors of cholinesterase). ric‐3 mutations were also identified in screens for mutants defective for cholinergic transmission at the neuromuscular junction (Nguyen et al., 1995; Miller et al., 1996). These data suggested that ric‐3 activity is required for the function of both receptors: the neuronal acetylcholine receptor and the muscle acetylcholine receptor. Here we demonstrate that RIC‐3 is needed for the activity of multiple acetylcholine gated channels and is not needed for the activity of other ligand gated channels. Our characterization of ric‐3 function suggests that in the absence of this gene, acetylcholine receptors are not transported to the cell surface and that heterologous expression of this protein greatly improves surface expression of these receptors. Thus, RIC‐3 is the first identified component of the biosynthetic machinery specifically required for the functional maturation of acetylcholine receptors.


Screen for suppressors of deg‐3‐induced cell death

deg‐3(u662) is a missense mutation in the pore‐forming domain of the acetylcholine receptor subunit DEG‐3. This mutation causes degeneration of the neurons expressing this subunit, and these cell deaths lead to uncoordinated movement and mechanosensory defects in mutant worms (Treinin and Chalfie, 1995; Treinin et al., 1998). We screened for mutations that suppressed the behavioral defects caused by the deg‐3(u662) mutation. Presumably, suppressor mutations would identify genes required for DEG‐3 receptor function. We identified 51 mutations that suppressed the behavioral defects as well as neuronal degeneration. These mutations fall into three groups: mutations in deg‐3 itself (36 mutations), mutations in des‐2 (a second subunit of the DEG‐3 receptor; 11 mutations; Treinin et al., 1998) and mutations in a third gene that we originally named des‐5 (four mutations: hm9, hm19, hm54 and hm65).

des‐5 mutations are strong suppressors of deg‐3(u662)‐induced cell degeneration. Cell swelling, the visible hallmark of deg‐3(u662)‐induced degenerations, is seen in 96% of deg‐3(u662) mutants 2 h after hatching. By contrast, only 4% of des‐5(hm65)deg‐3(u662) and 14% of des‐5(hm19)deg‐3(u662) animals exhibit degenerating cells (50 animals each). In wild‐type adults there are ∼11 cells (10.6 ± 1.9, n = 28 animals) which can be observed expressing a deg‐3lacZ construct (hmIs1). In deg3(u662), only three cells survive (3.36 ± 1.3, n = 50). However, seven cells survive in des‐5(hm9)deg‐3(u662) double mutants (7.1 ± 2.6, n = 50). Although des‐5 mutations suppress the uncoordinated phenotype caused by the deg‐3(u662) mutation, the double mutants do not exhibit wild‐type movement, and des‐5 animals are uncoordinated in the absence of the deg‐3(u662) mutation. The des‐5 animals coil when moving backwards. Moreover, they are slightly small and transparent, which is a phenotype of starved animals, a starved appearance that is probably caused by reduced pharyngeal pumping (see below). They also display a temperature‐sensitive growth phenotype: animals will either arrest growth or display reduced fertility upon transfer to 25°C.

We mapped des‐5 to an interval between unc‐8 and unc‐24 on chromosome IV (Figure 1A). ric‐3 is also a gene that falls in this region, and des‐5‐associated phenotypes are similar to phenotypes seen in ric‐3 mutants (Nguyen et al., 1995; Miller et al., 1996). A complementation test demonstrated that des‐5(hm9) is allelic to ric‐3(md158). In addition, ric‐3(md158), like the des‐5 alleles, suppresses deg‐3(u662)‐induced degeneration [only 4% of ric‐3(md158)deg‐3(u662) newly hatched larvae have degenerating cells; 50 animals]. We concluded that des‐5 and ric‐3 are alleles of the same gene, and the gene will be described henceforth by its previously published name of ric‐3.

Figure 1.

Cloning and molecular analysis of ric‐3. (A) Genetic map of chromosome IV showing the position of ric‐3 relative to nearby genes. (B) Physical map of cosmid T14A8, showing positions of predicted genes (T14A8.1, which encodes ric‐3, is in bold) and of restriction sites used in this study. (C) Structure of the ric‐3 gene. Exon structure was determined from the analysis of four cDNAs. Blocks, exons; bold line, non‐translated regions; thin line, 5′ upstream non‐transcribed region. The alternatively spliced exon is marked with an asterisk. Sites of mutation in ric‐3 are indicated: md1013, md1034, md1181, md1272, md1274, md1286 and md1442 are Tc1 insertions, hm9, hm19, hm54 and hm65 are nonsense mutations, and md146, md156, md158 and md226 are deletions (12.5 kb, 7 bp, 5 bp and 12 bp, respectively). (D) Sequence of the RIC‐3 protein derived from cDNAs (yk704a8 and yk722a6). Single underlines indicate transmembrane regions, double underlines indicate coiled‐coil regions, and the dashed line indicates amino acids that are absent in the alternatively spliced cDNA variants (yk719f3 and yk266d12), in which the last G in this segment is converted to an R.

Cloning of ric‐3

ric‐3 maps to a narrow region between unc‐8 and unc‐24 on chromosome IV (Figure 1A). This region is covered by 26 cosmids. These cosmids were injected into ric‐3 (md1181) animals and assayed for their ability to rescue ric‐3‐associated phenotypes. The cosmid T14A8 rescued ric‐3. In addition, we probed this interval using Southern blot analysis. Probing with a 9.7 kb BamHI fragment from the T14A8 cosmid revealed DNA polymorphisms in five ric‐3 mutants isolated from the mutator strain TR638 (Miller et al., 1996; md1181, md1274, md1286, md1013 and md1272; Figure 1B and C). Insertions consistent with the size of the C.elegans transposon Tc1 were identified in two adjacent EcoRI fragments of the single gene T14A8.1. ric‐3(md1181hm57) is a wild‐type revertant of ric‐3 (md1181). The insertion observed in md1181 was absent in the revertant, indicating that this putative transposon is the relevant mutation in this strain. A 6.5 kb XhoI–PstI subclone (Figure 1B) containing only this gene was sufficient for rescue of ric‐3‐associated phenotypes in transgenic animals (three of three transgenic lines), demonstrating that T14A8.1 is the coding region for ric‐3.

Four cDNA clones (gift of Y.Kohara) from this locus were obtained and sequenced. Two of these cDNAs were SL1 spliced and thus indicate that the entire open reading frame (ORF) has been identified (Krause and Hirsch, 1987). Two of these four cDNAs lacked a small second intron and thus indicate that the ric‐3 transcript is alternatively spliced. These cDNAs encode proteins of 363 or 378 amino acids (Figure 1D). The first ATG found in these ORFs is preceded by CAAA, a close match to the C.elegans translation initiation consensus (Blumenthal and Steward, 1997). Three Tc1 insertions are located within a 0.5 kb fragment in the promoter of this gene; these Tc1 insertions are likely to affect transcription of ric‐3. All other ric‐3 mutations, including Tc1 insertions, deletions and point mutations, affect the first half of these ORFs. All point mutations and deletions except md226 lead to severe truncation of the protein (Figure 1C and D). md158 is a 5 bp deletion and hm9 is a nonsense mutation, and are used as null mutations in these studies.

The RIC‐3 protein

Sequence analysis of ric‐3 shows that it encodes a novel protein (Altschul et al., 1990) showing no homology to characterized proteins in the databases. RIC‐3 is a highly charged protein, comprised of 8% aspartic acid, 13% glutamic acid, 10% lysine and 8% arginine (Figure 1D). Structure predictions suggest that RIC‐3 lacks a signal peptide but contains two transmembrane domains followed by three coiled‐coil regions (Berger et al., 1995; Sonnhammer et al., 1998). According to these predictions, RIC‐3 is located on membranes with its N‐terminal domain and C‐terminal coiled‐coil domains in the cytoplasm (Figure 2B; Hartmann et al., 1989). Coiled‐coil motifs have previously been implicated in protein–protein interactions where they directly associate with other coiled‐coil domain‐containing proteins (Stefancsik et al., 1998).

Figure 2.

A RIC‐3 homolog. (A) Homology with Drosophila CG9349 protein in the region spanning the transmembrane domains. Underlines indicate putative transmembrane domains. (B) Structure and topology predictions for RIC‐3 and CG9349. Barrels indicate positions of coiled‐coil regions. CG9349 has only one predicted coiled‐coil domain.

Although RIC‐3 is not homologous to previously studied proteins, it is weakly homologous to a protein, CG9349, identified by the Drosophila genome project (The Drosophila Sequencing Consortium, 2000). This protein is 35% identical to RIC‐3 in the two transmembrane domains and in a small region between these domains (Figure 2A). In addition, both genes contain coiled‐coil regions C‐terminal to the transmembrane domains. Structure predictions also suggest that, like RIC‐3, the Drosophila protein CG9349 has no signal peptide and has its coiled‐coil domain in the cytoplasm (Hartmann et al., 1989).

To determine in which cells the ric‐3 gene is expressed, we inserted the coding region for green fluorescent protein (GFP) after the first ATG and in‐frame with the rest of the ric‐3 gene in the rescuing genomic fragment. Expression of the GFP‐tagged protein in ric‐3(md1181) fully rescued all ric‐3‐associated phenotypes. GFP was found in the pharyngeal muscles, the body wall muscles and in many neurons (Figure 3). Using the number and position of RIC‐3‐expressing cell bodies as our criteria, we could account for all neurons in the body (apart from the major head and tail ganglia). As the density of cell bodies in these ganglia precludes accurate counting, we specifically focused on one group of sensory neurons, the amphid and phasmid neurons, most of which stain with the lipophilic dye DiD. All DiD‐staining neurons also expressed ric‐3gfp (data not shown). Thus, RIC‐3 is expressed in most if not all C.elegans neurons, including all motor neurons of the ventral cord, many sensory neurons (touch receptor and chemosensory neurons) and interneurons.

Figure 3.

Localization of RIC‐3. RIC‐3 localization was visualized using a RIC‐3::GFP functional fusion. (A) Head region, showing the pharyngeal muscles (the double‐lobed organ in the middle) and head ganglia neurons seen between the two lobes of the pharynx. Confocal section, scale bar: 10 μM. (B) Body muscles and neurons. The row of neurons in the middle marked by asterisks are motor neurons, on both sides of which are seen the body muscles. In the left corner, marked by an arrow, are the posterior–lateral ganglion neurons showing strong fluorescence in cell bodies and weak fluorescence of processes. Confocal section, scale bar: 50 μM. (C) RIC‐3 localization in the body muscles as visualized using a myo‐3ric‐3gfp fusion. Scale bar: 20 μM.

In addition to the identification of cells expressing RIC‐3, we also looked at the intracellular localization of this functional GFP fusion. In neurons, fluorescence was absent in the nucleus and was strong in the soma, and in the body muscles fluorescence was often seen in cytoplasmic membranous structures (Figure 3B). Because RIC‐3 was found in the cell soma and is a membrane protein, it is possible that the protein is localized to the ER. Localization to the ER would suggest a role for RIC‐3 in the assembly of acetylcholine receptors. However, weak GFP fluorescence is also seen in neuronal processes including the ventral cord and nerve ring; thus, other roles such as in transport cannot be excluded.

ric‐3 is specifically required for cholinergic transmission

ric‐3 mutants are resistant to the acetylcholinesterase inhibitor aldicarb (Nguyen et al., 1995), which indicates a defect in cholinergic transmission at neuromuscular junctions of the body wall muscle. This neurotransmission defect appears to be postsynaptic, since ric‐3 animals are also resistant to the anthelmintic levamisole (Miller et al., 1996). Levamisole is an agonist of the body wall muscle acetylcholine receptor (Lewis et al., 1980). Given that RIC‐3 is also required for the neuronal acetylcholine receptor DEG‐3/DES‐2, these data suggested that RIC‐3 may play a general role in acetylcholine receptor function.

There are four well characterized acetylcholine receptors in C.elegans: the neuronal receptor DEG‐3/DES‐2, the pharyngeal muscle receptor, the body muscle levamisole‐sensitive receptor and the body muscle nicotine‐sensitive receptor. In vivo electrophysiological recordings of the last three receptors are currently feasible. To assay directly the effects of ric‐3 mutations on the activity of acetylcholine receptors, we examined acetylcholine currents from these receptors in ric‐3 mutants.

Pharyngeal contraction is regulated by two neurotransmitters: acetylcholine initiates muscle contraction and glutamate terminates it. The effects of these transmitters can be assayed using extracellular recordings called electropharyngeograms (EPGs; Raizen and Avery, 1994). In accordance with the starved appearance of ric‐3 mutants, pharyngeal pumping is greatly reduced in ric‐3(hm9) adults (45 ± 5 pumps/min) compared with the wild type (>200 pumps/min; 20 adult animals each). Interestingly, the reduced pumping rates are observed in adults but not in larvae. Contraction of the pharyngeal muscle is initiated by the cholinergic MC neuron (Raizen et al., 1995). Synaptic currents from MC can be observed as a small inward current in the electropharyngeogram of the wild type (Figure 4). In ric‐3(hm9) mutants, normal MC spikes are not seen; however, 70% of pumps are still preceded by a small depolarizing peak (40 pumps), possibly sub‐threshold MC spikes (Figure 4). Thus, it is likely that the activity of the pharyngeal muscle acetylcholine receptor is also affected by ric‐3 mutations. Pharyngeal contractions are terminated by inhibitory glutamate transmission from the M3 motor neuron. In contrast to cholinergic transmission, these glutamate‐mediated outward currents are unaffected in ric‐3 mutants (Figure 4). Thus, in the pharyngeal muscle, RIC‐3 is specifically required for cholinergic transmission and does not cause a general defect in synaptic transmission.

Figure 4.

Defective cholinergic transmission in the C.elegans pharyngeal muscle. Representative traces for the electrical activity of the pharynx of wild‐type (N2) and ric‐3(hm9) mutant animals. MC EPSP is the excitatory cholinergic activity, seen as a small spike preceding muscle depolarization (large upward spike). M3 IPSP is the inhibitory glutamatergic activity, seen as a series of small spikes preceding muscle repolarization (large downward spike). In the ric‐3(hm9) mutant, the small depolarizatory current seen in the beginning of the trace may represent I phase activity, i.e. a sub‐threshold MC spike.

The body wall muscle expresses two distinct acetylcholine receptors and one GABA receptor (Richmond and Jorgensen, 1999). Using whole‐cell, voltage‐clamp recordings of individual body wall muscles, it is possible to distinguish the two acetylcholine receptors because one is sensitive to levamisole while the other is sensitive to nicotine (Richmond and Jorgensen, 1999). Application of acetylcholine in wild‐type animals activates both receptors. In ric‐3(md158) animals, the response to acetylcholine was greatly diminished compared with the wild type (12 ± 2 pA, n = 6 and 1560 ± 255 pA, n = 8, respectively), suggesting that both receptors are inactive in ric‐3 mutants (Figure 5). Responses to nicotine were absent in ric‐3(md158) animals but were robust in the wild type (0.1 ± 0.1 pA, n = 9 and 646 ± 124 pA, n = 8, respectively). Response to levamisole was also eliminated in ric‐3(md158) compared with the wild type (0.3 ± 0.3 pA, n = 6 and 203 ± 30 pA, n = 8, respectively). However, responses were observed at very high agonist concentrations; 1 mM acetylcholine was able to elicit very low amplitude responses (97.1 ± 11 pA, n = 4), suggesting that in ric‐3 mutants small numbers of functional acetylcholine receptors are still present on the cell surface. In contrast, application of GABA elicits the same response in both ric‐3 (2860 ± 636 pA, n = 10) and wild‐type animals (2840 ± 714 pA, n = 6; Figure 5). Thus, ric‐3 is specifically required for cholinergic transmission in both pharyngeal muscles and body muscles.

Figure 5.

Defective response to acetylcholine receptor agonists in the body muscles. Electrical responses of wild‐type and mutant ric‐3(md158) body muscles to puffs of ACh, nicotine, levamisole or GABA (100 μM each). On the left are representative traces of the responses; arrows indicate the time of agonist application. On the right is shown the average of the responses.

ric‐3 mutants do not respond to acetylcholine receptor agonists, evidence for a defect in receptor biogenesis or activity. To determine whether RIC‐3 is required within the same cells that express these receptors, we examined whether expression of RIC‐3 within the body muscles is sufficient for normal activity of the body muscle levamisole receptor. Indeed, expression of RIC‐3 from the myo‐3 promoter (Fire and Waterson, 1989) is sufficient for rescue of the levamisole insensitivity of ric‐3(md1181) mutants; transgenic ric‐3(md1181) animals expressing RIC‐3 in the body muscle alone arrest movement on 1 mM levamisole (82%), unlike the original ric‐3 (md1181) animals, which show no arrest of movement (0%, 50 animals each). In order to rule out the possibility that RIC‐3, like agrin, an extracellular matrix protein needed for clustering of nAChRs, can be supplied by both muscles and neurons (Reist et al., 1992), we examined whether expression of RIC‐3 in neurons could also rescue the insensitivity of ric‐3(md1181) to levamisole. For this purpose we fused the ric‐3 ORF to the aex‐3 promoter, a general neuronal promoter (Iwasaki et al., 1997). This construct, unlike the myo‐3::ric‐3 construct, did not rescue levamisole insensitivity in two lines of ric‐3(md1181) transgenic animals (0% arrested animals, 50 animals each). However, this analysis and our RIC‐3 localization studies (above) could not rule out the possibility that RIC‐3 functions in the synapses for trafficking or function of the levamisole‐sensitive receptor. Thus we examined whether rescue of this receptor's activity depends on synaptic localization of RIC‐3. For this purpose we looked at the localization of a myo‐3::ric‐3gfp fusion that rescues the levamisole insensitivity of ric‐3(md1181) in two lines examined (68 and 82% arrested animals, n = 50 each). This fusion is seen in muscles only, and is localized to membranous structures within these cells (Figure 3C). No GFP staining was detected in the ventral cord or dorsal cord, sites of C.elegans neuromuscular junctions (50 animals). Neither could we detect synaptic RIC‐3 in confocal thin sections (1 μm sections, n = 10). Thus, it is likely that the effect of RIC‐3 on the levamisole‐sensitive receptor is carried out within the muscle, and does not require co‐localization in the synapse.

RIC‐3 is needed for receptor assembly or trafficking

ric‐3 mutations could affect acetylcholine receptors by altering expression, localization or function. To determine whether these mutations affect the expression of an acetylcholine receptor subunit, we examined DEG‐3 immunostaining in ric‐3 mutants. In ric‐3 mutants, strong DEG‐3 staining is seen in the cell bodies, demonstrating that RIC‐3 is not required for DEG‐3 expression. However, there was a change in distribution of DEG‐3 immunoreactivity in ric‐3 mutants. For example, in wild‐type animals, both cell bodies and axons of the PVD neurons stain with similar intensities. However, in ric‐3 animals, DEG‐3 immunoreactivity in axons is always much weaker than immunoreactivity of cell bodies (50 animals; Figure 6). These data suggest that the absence of cholinergic transmission is not caused by a functional defect in properly transported receptors, but is caused by a defect in receptor folding, assembly or trafficking.

Figure 6.

DEG‐3 accumulates in cell bodies of ric‐3 mutants. Immunohistochemical analysis using DEG‐3 antibodies on young adults. Shown is the PVD neuron and processes in (A) wild type and (B) ric‐3(md158). Note the size difference between wild type and ric‐3, both of which are young adults. Confocal section, scale bar: 20 μM.

To directly test a role for RIC‐3 in acetylcholine receptor maturation we examined RIC‐3 function in Xenopus oocytes. We previously demonstrated that the DEG‐3/DES‐2 receptor can be expressed in oocytes (Treinin et al., 1998). Choline is the preferred ligand of the DEG‐3/DES‐2 receptor (Yassin et al., 2001). Reliable but small currents (83 ± 8 μA, n = 38 oocytes; 11 experiments) can be detected when non‐saturating concentrations of cRNA for both DES‐2 and DEG‐3 are co‐injected (Figure 7). However, when RIC‐3 cRNA is injected with DES‐2 and DEG‐3 cRNAs, peak currents are increased nearly 5‐fold (397 ± 36 μA, n = 38 oocytes; 11 experiments). This enhancement is seen on the second and third day following injection, and disappears on the fourth day. Expression of RIC‐3 alone does not produce detectable choline‐activated currents, nor was there any increase in the amplitude of glutamate‐activated currents when GluR3 was co‐expressed with RIC‐3 (Figure 7). Thus, RIC‐3 maintains specificity for acetylcholine receptors in Xenopus oocytes. Although RIC‐3 is not absolutely necessary for the maturation of acetylcholine receptors in Xenopus oocytes, it does confer increased efficiency on the maturation of acetylcholine receptors.

Figure 7.

RIC‐3 co‐expression enhances acetylcholine receptor activity in Xenopus oocytes. Amplitude of choline‐ (3.2 mM) or glutamate‐ (1 mM) dependent currents measured in oocytes together with (striped) or without (filled) co‐expression of RIC‐3. The DEG‐3/DES‐2 experiments represent n = 38 (oocytes), N = 6 (frogs) each. α‐7, n = 23, N = 4; GluR3, n = 7, N = 2; and RIC‐3 alone, n = 10, N = 3. The effects of co‐expressing RIC‐3 on DEG‐3/DES‐2‐ and α‐7‐dependent current amplitudes are significant using a paired t‐test at >99%.

To examine whether RIC‐3 could enhance the maturation of non‐C.elegans acetylcholine receptors, we co‐expressed RIC‐3 with the rat α‐7 homomeric receptor (Seguela et al., 1993). Although the sequence of rat α‐7 is only 30% identical to those of DEG‐3 or DES‐2 (Treinin et al., 1998), co‐expression of RIC‐3 with rat α‐7 leads to a 4.2 ± 0.36‐fold increase in peak current amplitude (Figure 7; n = 23 oocytes; six experiments). Thus, it appears that the ability of RIC‐3 to increase acetylcholine receptor activity is mediated by conserved domains within these subunits.


Mutations in ric‐3 were isolated because they were able to suppress necrotic cell death caused by the deregulated activity of a mutant acetylcholine receptor subunit DEG‐3. We cloned ric‐3 and found that it encodes a novel protein expressed in muscles and neurons (Figures 1 and 3). Detailed examination of ric‐3 mutants shows a reduction in cholinergic transmission mediated by four different ionotropic acetylcholine receptors: the DEG‐3/DES‐2 receptor, the pharyngeal muscle receptor, and the levamisole‐ and nicotine‐sensitive body muscle receptors (Figures 4 and 5). While ric‐3 mutations reduce synaptic transmission in a large number of cells, the gene appears to be specific for acetylcholine‐sensitive ligand‐gated ion channels. Neither GABA transmission mediated by the UNC‐49 receptor (Richmond and Jorgensen, 1999) nor glutamate transmission mediated by the pharyngeal receptor AVR‐15 (Dent et al., 1997) is affected in ric‐3 mutants (Figures 4 and 5). This specificity of the effects of RIC‐3 appears to conflict with its wide expression. However, the C.elegans genome codes for an unusually large number of nAChRs (∼40 predicted nAChR subunits; Bargmann, 1998; Mongan et al., 1998). Thus, the wide expression of RIC‐3 suggests that it is needed for the activity of many of these nAChRs. For example, B motor neurons express RIC‐3 and also ACR‐5, a nAChR α subunit (Winnier et al., 1999); thus, it is possible that RIC‐3 is also needed for the activity of ACR‐5.

Co‐expression with RIC‐3 is required for acetylcholine receptor activity in C.elegans body muscles and is sufficient for enhanced acetylcholine receptor activity in Xenopus oocytes. RIC‐3 appears to be a protein required for acetylcholine receptor biogenesis. First, the RIC‐3 protein appears to be localized to a membranous organelle in the cell body, consistent with localization to the ER (Figure 3). Secondly, the DEG‐3 receptor subunit accumulates in neuronal cell bodies in ric‐3 mutants (Figure 6). It is known that acetylcholine receptors are assembled in the ER and that improperly folded or assembled receptors accumulate in the ER (Smith et al., 1987). Thus, RIC‐3 may function during folding, assembly or transport from the ER into the Golgi apparatus. Consistent with this role, co‐expression of the C.elegans DEG‐3/DES‐2 or the rat α‐7 acetylcholine receptors with RIC‐3 was sufficient for enhanced acetylcholine receptor activity, likely to be a result of increased surface expression of these receptors. RIC‐3 is the first example of a protein specifically required for the maturation of acetylcholine receptors.

Biogenesis of membrane proteins is a complex process requiring 2–3 h (reviewed in Green, 1999; Keller and Taylor, 1999). Specifically for acetylcholine receptors, this process includes folding, generation of the appropriate membrane topology, assembly with specific subunits to form a pentamer, glycosylation, disulfide bond formation and, finally, targeting to specific sites on the membrane. The only proteins previously implicated in acetylcholine receptor biogenesis were BiP, calnexin and 14‐3‐3η (Blount and Merlie, 1991; Chang et al., 1997; Jeanclos et al., 2001). However, these are proteins required for the folding and maturation of many proteins. Our work on ric‐3 provides evidence for an acetylcholine receptor‐specific processing pathway. However, it seems unlikely that this processing pathway is unique to C.elegans. Evidence for specific requirements for acetylcholine receptor biogenesis in mammals is provided by studies showing low cell‐surface expression of acetylcholine receptors following heterologous expression. Specific ally, cell‐surface expression of the α‐7 acetylcholine receptor was shown to be cell type dependent, suggesting that a specific biogenesis machinery is required for α‐7 cell‐surface expression (Cooper and Millar, 1997; Sweileh et al., 2000). Proteins similar to RIC‐3 may mediate the interaction between acetylcholine receptors and a general protein‐processing machinery, thus conferring on this machinery acetylcholine receptor specificity. A requirement for a RIC‐3‐like protein in vertebrate receptors is consistent with enhanced α‐7 activity observed with co‐expression of RIC‐3 in Xenopus oocytes.

RIC‐3 is one of a limited number of molecules that have a protein‐specific role in the biogenesis of membrane proteins (reviewed in Herrmann et al., 1999). The reasons for such specificity are unknown. One possibility is that specificity is only required in special cases. For example, Drosophila rhodopsin molecules depend on interactions with the photoreceptor specific chaperone NinaA for their surface expression (Schneuwly et al., 1989; Colley et al., 1995). It has been suggested that the requirement for NinaA is related to the unusually large numbers (1 × 108 functional receptors per cell) of rhodopsin molecules required in Drosophila photoreceptors, numbers that cannot be processed by the generalized processing machinery (Baker et al., 1994). The formation of a functional heteromeric pentamer having the proper order and stoichiometry, as is needed for the formation of acetylcholine receptors, may also be such a special case. However, an interesting alternative is that the ability to recognize and separately process different receptors provides an additional mechanism for regulating one class of receptors while leaving other membrane proteins unaffected. The inefficient maturation of acetylcholine receptors (only 30% of newly synthesized subunits assemble to a functional receptor; Merlie and Lindstrom, 1983) opens the way for regulating acetylcholine receptor activity through changes in the efficiency of receptor maturation. It is interesting in this regard to note that RIC‐3 is expressed widely and affects at least four different acetylcholine receptors. Thus, RIC‐3 may mediate global signals that regulate cholinergic transmission. An example of such a control mechanism, where a global signal regulates surface expression, is provided by the effect of insulin on the GABAA receptor (Wan et al., 1997).

Materials and methods

Strain maintenance and genetics

The wild type was N2 Bristol and all strains were grown as previously described (Wood et al., 1988). Screens for suppressors of deg‐3(u662) were described previously (Treinin and Chalfie, 1995). In short, >30 000 EMS mutagenized haploid genomes were screened for suppression of the Unc and Mec phenotypes associated with deg‐3(u662).

Complementation tests were conducted by crossing ric‐3(md158)/+ males with dpy‐13(e184)des‐5(hm9) hermaphrodites. Half of the non‐Dpy progeny of this cross were coilers and resistant to levamisole. Mapping was carried out as follows: among 256 Dpy non‐Ric progeny of dpy‐20(e1282)ric‐3(md146)/unc‐24(e138), 251 had progeny with the unc‐24(e138) phenotype; among 74 Dpy non‐Ric progeny of dpy‐13(e184)ric‐3(md146)/unc‐8(e49), 69 had progeny with the unc‐8(e49) phenotype; and among 74 Dpy non‐Ric progeny of dpy‐13(e184)ric‐3(md146)/unc‐24(e138), all 74 had progeny with the unc‐24(e138) phenotype. This analysis shows that ric‐3 is within the 0.26 m.u. interval between unc‐8 and unc‐24, approximately at position 3.52 on the genetic map.


Levamisole resistance was examined on NGM plates (Wood et al., 1983) containing 1 mM levamisole (Sigma). Sensitive L4‐adult animals contract, arrest movement within minutes, and eventually die. Aldicarb resistance was examined on NGM plates with 1 mM aldicarb (Chemservice). Two L4 animals were placed on each plate and then examined for the generation of fertile progeny 7 and 14 days later.

Electrophysiology of C.elegans muscles

Extracellular recordings from the pharynx were made as in Raizen and Avery (1994). For the pumping rate data, L4s were transferred to fresh agar plates 1 day prior to the pumping rate assay and kept at 20°C. Immediately before the assay, animals were transferred to room temperature in order to count pumps. Electrophysiology of the body muscles was as in Richmond and Jorgensen (1999).

Antibody, lacZ and DiD staining

DEG‐3 localization was visualized as described in Yassin et al. (2001). Strains carrying the following alleles of ric‐3 were examined: md146, md158, hm9, md226 and md1181. We saw the same defect in all strains. Survival of DEG‐3 expressing neurons and effects of RIC‐3 on DEG‐3 transcription were assayed using a deg‐3‐lacZ construct (Treinin et al., 1998). DiD C18 (Molecular Probes D‐7757) was dissolved in DMF at 10 mg/ml. ric‐3gfp transgenics were incubated for 1 h at room temperature with rocking in 1:1000 DiD diluted in M9 buffer, followed by 2 h destaining on seeded NGM plates. Visualization was performed on 2% agar pads + 4% paraformaldehyde.

Identification of RIC‐3 expressing neurons

RIC‐3::GFP stained all 57 motor neurons in the ventral cord in five of 10 animals examined, all six cells in the left posterior lateral ganglion in five of eight animals examined, all four cells of the right lateral ganglion in two of seven animals examined, and all six other cells between the vulva and head in nine of 10 animals examined. In the tail ganglia, we never counted the full complement (33 cells), rather we counted 26 ± 2 cells (n = 10). Among the cells in the tail ganglia, we identified PVC L, DVA, DVB and DVC by position, and PHA L/R and PHB L/R using DiD staining. In the head ganglia, no attempt was made to count cells. However, the amphid cells ASH L/R, ASJ L/R, ASK L/R and ADL L/R were identified using DiD staining.

Molecular biology

General molecular biology methods followed the protocols in Sambrook et al. (1989). Southern blots were probed with random primed cosmids or DNA fragments. Specifically, using a 9.7 kb BamHI fragment from cosmid T14A8, we could visualize 1.6 kb insertions in DNA from the ric‐3 mutant strains. Specifically, in md1013 and md1272, the 4.5 kb EcoRI fragment was shifted, and in md1181, md1274 and md1286, the adjacent 2.2 kb EcoRI fragment shifted. Both of these bands were absent in md146. For rescue experiments, cosmids or subclones were co‐injected with the rol‐6 DNA (pRF4) (Mello and Fire, 1995) and roller progeny of stable transgenic lines were examined for pumping and levamisole resistance. The smallest rescuing subclone is a 6.5 kb XhoI–PstI fragment from cosmid T14A8 ligated into pBluescript SK (Stratagene).

To tag RIC‐3 with GFP, an EcoRV site was inserted immediately downstream of the first ATG of RIC‐3 using PCR‐mediated mutagenesis. An EcoRV fragment containing GFP from the plasmid pPD102.33 (Fire et al., 1990) was ligated to this EcoRV site. This reporter construct was injected at 5 ng/μl and supplemented to 50 ng/μl with pBluescript. Integration of the extrachromosomal array was obtained using UV irradiation (Mitani, 1995).

ric‐3 alleles were sequenced directly from PCR‐amplified DNA. Sites of Tc1 insertions were estimated from the size of the PCR product generated by Tc1‐specific primers combined with RIC‐3‐specific primers. The structure of the RIC‐3 gene was obtained from the sequences of four cDNAs, yk266d12, yk704a8, yk719f3 and yk722a6, which were kindly provided by Y.Kohara. Three of these, yk704a8, yk719f3 and yk722a6, were obtained from libraries enriched for full‐length cDNAs (Suzuki et al., 1997). yk266d12, yk704a8 and yk722a6 all terminate 12 bp upstream of the first ATG in the RIC‐3 ORF. In both yk704a8 and yk722a6, this site is trans‐spliced to an SL‐1 leader sequence (Krause and Hirsch, 1987); only yk719f3 terminates downstream of this site with its 5′ end 77 bp downstream, and may represent a truncated cDNA. The second intron encoding 15 amino acids is not spliced out of yk704a8 and yk722a6.

Expression from the myo‐3 promoter was achieved by inserting a 2.5 kb SalI–AscI fragment containing the myo‐3 promoter (Fire and Waterson, 1989) in place of a 2.5 kb XhoI–AscI fragment containing the RIC‐3 promoter. For this purpose, an AscI site was inserted immediately upstream of the RIC‐3 ATG using PCR‐mediated mutagenesis. A SalI GFP fragment from pPD103.87 was inserted into an internal SalI site in the same construct to generate the myo‐3ric‐3gfp construct. In this construct, GFP is fused to the full‐length RIC‐3 downstream of the first coiled‐coil domain. The aex‐3ric‐3 construct was generated by inserting an ApaI–PstI aex‐3 promoter fragment from pMH50 into appropriate sites generated using linkers upstream of the RIC‐3 ATG. These sites were inserted into the myo3ric‐3 construct in place of the myo‐3 promoter.

Sequence analysis

Structure prediction analysis was carried out using programs on and The Flybase accession number for CG9349 is FBgn0034575. Alignment was generated using Clustal_W analysis (McVector).

Heterologous expression and electrophysiology

An EcoRI fragment containing RIC‐3 ORF was cut out of the yk266d12 cDNA clone and cloned into a pCDNA3 variant that includes 5′ and 3′ UTRs from the Xenopus β‐globin gene (Invitrogene). This clone contains 12 bp of the ric‐3 5′ untranslated sequences and no 3′ untranslated sequences. In vitro transcribed cRNAs were injected at a final concentration of 0.1–0.5 ng. In each experiment, final concentrations of DEG‐3, DES‐2, α‐7 or GluR3 were kept constant between injections with or without RIC‐3. Injections and recordings were carried out as described in Treinin et al. (1998). Choline, which is the preferred agonist of the DEG‐3/DES‐2 channel (Yassin et al., 2001), was used at saturating concentrations of 3.2 mM and glutamate was used at 1 mM. For the glutamate receptor experiments we used a mutant GluR3(L507Y), which enables better visualization of glutamate‐induced currents in Xenopus oocytes, as the L507Y point mutation eliminates AMPA receptor desensitization without significantly affecting other parameters (Stern‐Bach et al., 1998). In each experiment, 3–5 oocytes with RIC‐3 and 3–5 oocytes without RIC‐3 were examined. Comparisons of current amplitude were made for experiments where the average current amplitude for oocytes not injected with RIC‐3 was between 50 and 150 nA, reducing the variability inherent to the oocyte expression system. Errors are standard errors of means, and significance was examined using the paired t‐test.


We thank Y.Stern‐Bach for many helpful suggestions, Jim Rand for ric‐3 alleles, A.Fire for vectors, Y.Kohara for cDNA clones, Marc Hammarlund for pMH50, the C.elegans genetics stock center for strains and the Sanger center for cosmids. This work was supported by a US–Israel Binational Science Foundation Grant 1999‐074‐01 and by The Israel Science Foundation founded by the Israel Academy of Sciences and Humanities–DOROT Science Fellowship Foundation.


View Abstract