1,4,5-trisphosphate (IP3)-dependent Ca2+ signaling regulates gonad function, fertility, and rhythmic posterior body wall muscle contraction (pBoc) required for defecation in Caenorhabditis elegans. Store-operated Ca2+ entry (SOCE) is activated during endoplasmic reticulum (ER) Ca2+ store depletion and is believed to be an essential and ubiquitous component of Ca2+ signaling pathways. SOCE is thought to function to refill Ca2+ stores and modulate Ca2+ signals. Recently, stromal interaction molecule 1 (STIM1) was identified as a putative ER Ca2+ sensor that regulates SOCE. We cloned a full-length C. elegans stim-1 cDNA that encodes a 530–amino acid protein with ∼21% sequence identity to human STIM1. Green fluorescent protein (GFP)–tagged STIM-1 is expressed in the intestine, gonad sheath cells, and spermatheca. Knockdown of stim-1 expression by RNA interference (RNAi) causes sterility due to loss of sheath cell and spermatheca contractile activity required for ovulation. Transgenic worms expressing a STIM-1 EF-hand mutant that constitutively activates SOCE in Drosophila and mammalian cells are sterile and exhibit severe pBoc arrhythmia. stim-1 RNAi dramatically reduces STIM-1∷GFP expression, suppresses the EF-hand mutation–induced pBoc arrhythmia, and inhibits intestinal store-operated Ca2+ (SOC) channels. However, stim-1 RNAi surprisingly has no effect on pBoc rhythm, which is controlled by intestinal oscillatory Ca2+ signaling, in wild type and IP3 signaling mutant worms, and has no effect on intestinal Ca2+ oscillations and waves. Depletion of intestinal Ca2+ stores by RNAi knockdown of the ER Ca2+ pump triggers the ER unfolded protein response (UPR). In contrast, stim-1 RNAi fails to induce the UPR. Our studies provide the first detailed characterization of STIM-1 function in an intact animal and suggest that SOCE is not essential for certain oscillatory Ca2+ signaling processes and for maintenance of store Ca2+ levels in C. elegans. These findings raise interesting and important questions regarding the function of SOCE and SOC channels under normal and pathophysiological conditions.
Changes in cytoplasmic Ca2+ levels regulate a diverse array of physiological processes and function as an essential signaling mechanism in virtually all cells (Berridge et al., 2003). Cytoplasmic Ca2+ levels are altered by Ca2+ flux across the plasma membrane and by release of Ca2+ from intracellular stores. The ER is a major source of intracellular Ca2+ release, which is mediated by 1,4,5-trisphosphate (IP3) receptor (IP3R) Ca2+ channels (Berridge et al., 2003). Early studies on parotid acinar cells demonstrated that activation of intracellular Ca2+ release was associated with increased plasma membrane Ca2+ entry (for review see Parekh and Penner, 1997). Putney (1986) postulated that the Ca2+ content of the ER stores directly regulated Ca2+ influx across the plasma membrane and that Ca2+ entry was activated by store depletion. This process was initially termed capacitive Ca2+ entry, but is now known as store-operated Ca2+ entry (SOCE). SOCE is thought to be essential for refilling ER Ca2+ stores and for modulating the time course and amplitude of cytoplasmic Ca2+ signals (Parekh and Penner, 1997; Venkatachalam et al., 2002; Parekh and Putney, 2005).
Hoth and Penner (1992) identified the first store-operated Ca2+ (SOC) current in 1992 by patch clamp electrophysiology. This current, termed Ca2+ release–activated Ca2+ current or ICRAC, has been the most extensively characterized SOCE pathway (Parekh and Penner, 1997; Parekh and Putney, 2005). Despite intense study, the mechanism by which store depletion activates SOCE has remained unclear until recently. Using an RNAi screen of SOCE in Drosophila S2 cells, Roos et al. (2005) identified stromal interaction molecule 1 (STIM1) as an essential component of CRAC activation.
Human STIM1 was identified originally as a potential tumor suppressor gene (Parker et al., 1996; Sabbioni et al., 1997). Two STIM homologues, STIM1 and STIM2, are present in the human genome (Williams et al., 2001). siRNA knockdown of STIM2 has no effect on SOCE in HEK293 cells (Roos et al., 2005) but inhibits HeLa cell SOCE (Liou et al., 2005). Recent studies suggest that STIM2 interacts with STIM1 and may function normally to inhibit SOC channels (Soboloff et al., 2006a,b).
STIM1 and STIM2 each have a predicted single membrane-spanning domain. In the absence of store depletion, STIM1 colocalizes with ER markers (Liou et al., 2005; Zhang et al., 2005). An EF-hand Ca2+ binding domain is present on the N terminus of STIM1 and is localized to the ER lumen. Depletion of ER Ca2+ stores or disruption of Ca2+ binding by mutation of the EF-hand domain induces translocation of STIM1 toward the plasma membrane (Liou et al., 2005; Zhang et al., 2005) with subsequent activation of SOCE (Liou et al., 2005; Zhang et al., 2005) and ICRAC (Spassova et al., 2006).
The nematode C. elegans provides a number of experimental advantages for defining the genes and integrated genetic pathways involved in biological processes such as Ca2+ signaling (Barr, 2003; Strange, 2003). The worm has a short life cycle, is genetically tractable, and has a fully sequenced and well-annotated genome. It is also relatively easy and economical to manipulate and hence characterize gene function in this organism using transgenic and RNAi methods.
Both the gonad and intestine of C. elegans have proven to be useful models for characterizing IP3-dependent Ca2+ signaling pathways using genetic, molecular, and physiological approaches (Clandinin et al., 1998; Dal Santo et al., 1999; Bui and Sternberg, 2002; Estevez et al., 2003; Kariya et al., 2004; Yin et al., 2004; Espelt et al., 2005; Estevez and Strange, 2005; Teramoto and Iwasaki, 2006). We have shown previously that intestinal epithelial cells express SOC channels (Estevez et al., 2003) and have postulated that SOCE plays an important role in gonad function (Rutledge et al., 2001; Yin et al., 2004). The focus of the current study was to characterize the function of STIM homologues in IP3 and oscillatory Ca2+ signaling pathways in C. elegans. A single gene, stim-1, encoding a predicted STIM1 homologue is present in the worm genome. We demonstrate here that stim-1 is essential for the normal IP3- and Ca2+-dependent contractile activity of gonad sheath cells and the spermatheca. stim-1 RNAi dramatically reduces STIM-1 expression and inhibits SOC channel activity in intestinal epithelial cells but has no effect on intestine oscillatory Ca2+ signaling and associated behavioral rhythms or on the intestinal ER unfolded protein response. Our studies provide the first detailed characterization of STIM-1 function in an intact animal and are the first to link this protein to specific whole animal physiological processes. In addition, our results indicate that SOCE is not required for all oscillatory Ca2+ signaling processes and raise interesting and important questions regarding the function of SOC channels under normal and pathophysiological conditions.
Materials And Methods
C. elegans Strains
Nematodes were cultured using standard methods (Brenner, 1974). Wild-type worms were the Bristol N2 strain. The following alleles were used: itr-1(sa73, sy290, sy327), ipp-5(sy605), lfe-2(sy326), rrf-1(pk1417), elt-2∷gfp(wIs84), rde-1(ne219), and hsp-4∷gfp(zcIs4). itr-1(sa73) is a temperature-sensitive allele. Worms harboring this mutation were grown at the permissive temperature of 16°C. For RNAi experiments, eggs from itr-1(sa73) worms were placed on RNAi feeding plates and grown at 25°C. All other strains were grown at 16–25°C. Unless stated otherwise, all experimental results presented were from studies conducted on young adult hermaphrodites.
Analysis of Sheath Cell Contraction and Ovulation
Synchronized L1 larvae were grown for 32–36 h at 25°C and then anesthetized in M9 solution containing 0.1% tricaine and 0.01% tetramisole for 20–40 min. Anesthesized worms were mounted onto 2% agarose pads (McCarter et al., 1999) and were imaged at room temperature (22–23°C) by differential interference contrast (DIC) microscopy using a Nikon Eclipse TE2000 inverted microscope and a Superfluor 40X/1.3 N.A. oil immersion objective lens. Images were recorded at 30 frames/s on videotape using a DAGE-MTI CCD100 camera and analyzed offline. Sheath contractions were counted in 1-min intervals. The intensity or “force” of individual contractions was estimated by measuring lateral displacement of the sheath as described previously (Miller et al., 2001).
Measurement of Brood Size
Brood size was quantified at 25°C by transferring single L4 larvae to new growth plates daily for 4 d. The number of progeny on each plate was counted 24–36 h after eggs hatched.
Characterization of pBoc Cycle
Posterior body wall muscle contraction (pBoc) was monitored at room temperature (21–22°C) in young adult worms. A minimum of 10 pBoc cycles was measured in each animal. Worms were imaged using a Carl Zeiss MicroImaging Inc. Stemi SV11 M2BIO stereo dissecting microscope (Kramer Scientific Corp.) equipped with a DAGE-MTI DC2000 CCD camera. pBoc rhythmicity in individual worms was assessed by calculating coefficient of variance (CV), which is the standard deviation expressed as a percent of the mean.
Dissection and Fluorescence Imaging of Intestines
Calcium oscillations and waves were measured in isolated intestines as described previously (Espelt et al., 2005). In brief, worms were placed in control saline (137 mM NaCl, 5 mM KCl, 1 mM MgCl2, 1 mM MgSO4, 0.5 mM CaCl2, 10 mM HEPES, 5 mM glucose, 2 mM l-asparagine, 0.5 mM l-cysteine, 2 mM l-glutamine, 0.5 mM l-methionine, 1.6 mM l-tyrosine, 28 mM sucrose, pH 7.3, 340 mOsm) and cut behind the pharynx using a 26-gauge needle. The hydrostatic pressure in the worm spontaneously extruded the intestine, which remained attached to the rectum and the posterior end of the animal. Isolated intestines were incubated for 15 min in bath saline containing 5 μM fluo-4 AM and 1% BSA. Imaging was performed at room temperature (21–22°C) using a Nikon TE2000 inverted microscope, a Superfluor 40X/1.3 N.A. oil objective lens, a Photometrics Cascade 512B cooled CCD camera (Roper Industries), and MetaFluor software (Universal Imaging Corporation). Fluo-4 was excited using a 490-500BP filter, and a 523-547BP filter was used to detect fluorescence emission. Changes in fluo-4 intensity were quantified using region-of-interest selection and MetaFluor software (Universal Imaging Corporation).
Calcium oscillation period, rise time (RT), and fall time (FT) were quantified as described previously (Prakash et al., 1997; Espelt et al., 2005). Fluorescence images were typically acquired at 0.2 Hz to avoid photobleaching and damage to the intestinal epithelium. However, when Ca2+ wave velocity and Ca2+ spike rise and fall times were quantified, images were acquired at 1 Hz.
C. elegans Embryonic Cell Culture and Patch Clamp Electrophysiology
Embryonic cells were cultured on 12-mm-diameter acid-washed glass coverslips using methods described previously (Christensen et al., 2002; Estevez et al., 2003). Intestinal cells were identified in culture by expression of the intestine-specific reporter elt-2∷GFP (Fukushige et al., 1998; Estevez et al., 2003).
Coverslips with cultured embryo cells were placed in the bottom of a bath chamber (model R-26G; Warner Instrument Corp.) that was mounted onto the stage of a Nikon TE2000 inverted microscope. Cells were visualized by fluorescence and video-enhanced DIC microscopy. Patch electrodes were pulled from soft glass capillary tubes (PG10165-4; World Precision Instruments) that had been silanized with dimethyl-dichloro silane. Pipette resistance was 4–7 MΩ. Bath and pipette solutions contained 145 mM NaCl, 20 mM CaCl2, 10 mM HEPES, 20 mM glucose, pH 7.2 (adjusted with NaOH), 345–350 mOsm, and 147 mM sodium gluconate (NaGluconate), 0.6 mM CaCl2, 6 mM MgCl2, 10 mM BAPTA, 10 mM HEPES, 10 μM IP3, pH 7.2 (adjusted with CsOH), 330 mOsm, respectively.
Whole-cell currents were recorded using an Axopatch 200B (Axon Instruments) patch clamp amplifier. Command voltage generation, data digitization, and data analysis were performed on a 1.6 GHz Pentium computer (Dimension 4400; Dell Computer Corp.) using a Digidata 1322A AD/DA interface with pClamp 8.2 and Clampfit 8.2 software (Axon Instruments). Electrical connections to the amplifier were made using Ag/AgCl wires and 3 M KCl/agar bridges. Leak current was defined as the current observed immediately after obtaining whole-cell access and was subtracted from all subsequent current records obtained from the cell.
RNA interference was induced by feeding worms bacteria producing dsRNA (e.g., Kamath et al., 2000; Rual et al., 2004). stim-1 and sca-1 RNAi bacterial strains were obtained from the ORF-RNAi feeding library (Open Biosystems) or from cDNAs generated by PCR. The ORF-RNAi stim-1 bacterial strain produced dsRNA that targeted the first eight exons of the gene, which encode amino acids 1–390 (see Fig. 1). For cameleon imaging studies, a stim-1 cDNA was generated by PCR and inserted into the RNAi feeding vector pPD129.36. dsRNA produced from this cDNA targeted amino acids 79–289 of the STIM-1 protein. BLAST searches of C. elegans genomic and EST databases failed to identify genes with homology to stim-1, indicating that off-target effects of RNAi are unlikely. GFP dsRNA–producing bacteria were engineered as described previously (Yin et al., 2004). Bacterial strains were streaked to single colonies on agar plates containing 50 μg/ml ampicillin and 12.5 μg/ml tetracycline. Single colonies were used to inoculate LB media containing 50 μg/ml ampicillin and cultures were grown at 37°C for 16–18 h with shaking. 300 μl of each bacterial culture was seeded onto 60-mm nematode growth medium agar plates containing 50 μg/ml ampicillin and 1 mM IPTG to induce dsRNA synthesis. After seeding, plates were left at room temperature overnight.
For cell culture studies, a stim-1 DNA template was generated by PCR from the ORF-RNAi clone using T7 primers. dsRNA was synthesized from the DNA template by T7 polymerase reactions (MEGAscript kit; Ambion, Inc.).
Isolated embryo cells were seeded onto glass coverslips in individual wells of four-well culture plates (Nalge Nunc International). The cells were incubated initially with 100 μl of modified (see Christensen et al., 2002) L-15 cell culture medium (Life Technologies) containing 15 μg/ml stim-1 dsRNA. After 2 h, the culture medium volume was increased to 300 μl and the final dsRNA concentration diluted to 5 μg/ml. An additional 100 μl of modified L-15 containing 5 μg/ml stim-1 dsRNA was added on the second and third day of culture. Cells were patched clamped 2–3 d after seeding.
Analysis of the Unfolded Protein Response
Synchronized L1 stage hsp-4∷GFP worms were transferred to RNAi feeding plates seeded with stim-1 or sca-1 dsRNA-producing bacteria or bacteria containing the empty feeding vector pPD129.36 as a control. Worms were maintained at 25°C for 30 h. 24 h after feeding was initiated, a group of control worms was transferred for 6 h to another control feeding plate containing 10 μg/ml tunicamycin. Changes in hsp-4∷GFP fluorescence were determined using a COPAS Biosort (Union Biometrica).
Construction of Transgenes and Transgenic Worms
A full-length stim-1 cDNA was cloned from C. elegans N2 total RNA by RT-PCR. Primers were designed based on the predicted WormBase sequence of Y55B1BM.1. Aspartate residues at positions 55 and 57 were mutated to alanine (i.e., D55A;D57A) using a Quikchange site-directed mutagenesis kit (Stratagene) and confirmed by DNA sequencing. Translational GFP reporters for wild type and D55A;D57A stim-1 were generated by insertion into vector pPD95.77. Expression of these reporters was driven by 1.9 kb of promoter sequence upstream of the stim-1 start codon. This sequence was amplified by PCR from C. elegans N2 genomic DNA and linked to the stim-1 cDNA by a PCR fusion-based method (Hobert, 2002). Transgenic worms were generated by DNA microinjection as described by Mello et al. (1991) using rol-6 as a transformation marker.
An integrated line of worms expressing STIM-1(D55A;D57A)∷GFP was generated by exposing 50 P0 Pstim-1∷STIM-1(D55A;D57A)∷GFP;rol-6(su1006) transgenic L4 animals to a dose of 30,000 μJ/cm2 of UV light that was generated with a UV cross-linker (Hoefer Scientific Instruments). We clonally isolated 500 F1 roller offspring and then isolated two F2 roller offspring from each F1 worm. A single integrated line was then isolated that segregated 100% GFP and rol-6 positive animals. This line was outcrossed four times to wild-type animals to generate KbIs15 (Pstim-1∷STIM-1(D55A;D57A)∷GFP; rol-6 (su1006)). As discussed in Results, STIM-1(D55A;D57A)∷GFP worms are sterile. To maintain the line, worms were grown on bacteria producing GFP dsRNA.
The yellow cameleon YC6.1 (Truong et al., 2001) was PCR amplified from pcDNA3.1YC6.1 and cloned into the Acc65I and EcoRI sites of the nematode expression vector pFH6.II (Nehrke and Melvin, 2002) to create pKT2. pKT2nhx-2 was created by cloning 4 kb of the nhx-2 promoter into NheI and SacII sites of pKT2. nhx-2 encodes an intestine-specific Na+/H+ exchanger (Nehrke and Melvin, 2002).
rde-1(ne219) worms were microinjected with a mixture of pKT2nhx-2 and pXXY2004.1rde-1 using rol-6 as a transformation marker. pXXY2004.1rde-1 encodes wild-type rde-1 driven by the 4-kb nhx-2 promoter (Espelt et al., 2005). Roller progeny from these microinjections expressed cameleon YC6.1 and wild-type rde-1 exclusively in the intestine. Wild-type worms were injected with pKT2nhx-2 and rol-6 only.
Fluorescence and DIC micrographs were obtained using a Carl Zeiss MicroImaging Inc. M2BIO stereo dissecting microscope and DAGE-MTI DC2000 CCD camera or a Nikon TE2000 inverted microscope and a Micromax CCD-1300 camera (Princeton Instruments). Confocal imaging was performed using an LSM510 confocal microscope equipped with 10×/0.3 N.A. and 40×/1.3 N.A. Plan-Neofluar lenses (Carl Zeiss MicroImaging Inc.). Pixel intensities were quantified using MetaMorph software (Universal Imaging Corporation).
FRET Imaging of Cameleon Protein
Fluorescence resonance energy transfer (FRET) imaging of intestinal cameleon was performed on L3 larvae freely moving on 60-mm agar plates using a Nikon 2000U inverted microscope equipped with a 10× objective lens, monochromatic light source (TILL Photonics), CCD camera detection system (Cooke), and Dual-View beamsplitter (Optical Insights). Cameleon was excited at 435 nm for 100 ms, and 480 nm and 540 nm emissions from the entire intestine were acquired simultaneously at 2 Hz using 2 × 2 binning. The 480/540 nm emission ratio after background subtraction and signal threshholding was used as an indicator of FRET efficiency.
Data are presented as means ± SEM. Statistical significance was determined using Student's two-tailed t test for unpaired means. When comparing three or more groups, statistical significance was determined by one-way analysis of variance. P values of ≤0.05 were taken to indicate statistical significance.
A Single STIM Homologue Is Present in the C. elegans Genome
BLAST searches of genomic and EST databases demonstrated that a single predicted STIM homologue (gene Y55B1BM.1; GenBank/EMBL/DDBJ accession no. AC024823) is present in the C. elegans genome (Williams et al., 2001). We cloned a full-length Y55B1BM.1 cDNA that encoded a 530–amino acid protein with a sequence identical to that of the WormBase-predicted transcript Y55B1BM.1a. The protein encoded by this cDNA has been termed STIM-1 (GenBank/EMBL/DDBJ accession no. DQ812088).
Sequence analysis indicated that STIM-1 is most similar to human STIM1 versus STIM2. Human STIM1 and Drosophila Stim possess several conserved domains, including an N-terminal signal peptide, an EF-hand Ca2+ binding motif, a SAM domain, a single predicted transmembrane domain, and a large C-terminal region predicted to encode a coiled-coil domain (Williams et al., 2001). These motifs are conserved in C. elegans STIM-1 (Fig. 1). The C terminus of human STIM1 contains a serine- and proline-rich domain and a lysine-rich domain. These domains are absent from Drosophila Stim (Williams et al., 2001). C. elegans STIM-1 also lacks the serine- and proline-rich domain, but a possible lysine-rich domain is present in its C terminus (Fig. 1). Williams et al. (2001) noted that human STIM1 and Drosophila Stim contain a pair of cysteine residues that are separated by six amino acids and that are located at similar positions in the N terminus (Fig. 1). These cysteine residues are not present in C. elegans STIM-1. N-linked glycosylation sites are present in human STIM1 at positions 131 and 171 (Williams et al., 2002; Fig. 1). These sites are conserved in Drosophila Stim (Zhang et al., 2005), whereas only the more C-terminal site is present in C. elegans STIM-1 (Fig. 1).
STIM-1 Is Expressed in Diverse Cell and Tissue Types
To identify cells in which STIM-1 is expressed, we generated transgenic worms expressing full-length STIM-1 fused to GFP. Expression was driven by 1.9 kb of the stim-1 promoter located immediately upstream of the start codon. Prominent expression of STIM-1∷GFP was detected in the spermatheca, gonad sheath cells, the intestine, and neurons in the head (Fig. 2). Expression was also detected in uterine epithelial cells (unpublished data). STIM-1∷GFP-expressing head neurons are likely amphid and/or inner labial (IL) neurons. Inner labial neurons may function in mechanosensation and chemosensation. Amphid neurons function to detect external osmotic, mechanical, and chemical stimuli (Bargmann and Mori, 1997). In vivo Ca2+ imaging studies have demonstrated a role for Ca2+ signaling in the response of ASH amphid neurons to noxious stimuli (Hilliard et al., 2005).
Expression of STIM-1∷GFP in the intestine was heterogeneous. The anterior and posterior intestine expressed the reporter very strongly while expression was weaker in the midsection (Fig. 2 A). Intestinal STIM-1∷GFP appeared to be localized to membrane and submembrane regions (Figs. 2, C and D). Confocal Z-sections also revealed a prominent punctate localization in the anterior intestine (Fig. 2 C) and sheath cells (Fig. 2 E). STIM-1∷GFP expression showed a striking localization to a reticular structure in the posterior intestine (Fig. 2 D). This reticular structure resembles the ER of C. elegans intestinal cells (Rolls et al., 2002). More detailed studies are required to identify the cellular domains in which STIM-1∷GFP is expressed.
It should be noted here that the absence of detectable STIM-1∷GFP expression in tissues other than those shown in Fig. 2 does not rule out a functional role for STIM-1 in other cell types. The 1.9-kb stim-1 promoter used in these studies may lack regulatory information required for cell-specific expression. In addition, STIM-1∷GFP expression levels may be below detection levels in other tissues. More definitive identification of STIM-1 expression sites awaits the development of suitable antibodies for immunolocalization.
STIM-1 Is Required for Fertility and Sheath Cell and Spermatheca Contractile Activity
To begin defining STIM-1 functions, we fed worms STIM-1 dsRNA-expressing bacteria beginning at the L1 larval stage. Young adult stim-1(RNAi) worms appeared healthy and exhibited no obvious defects in external morphology, movement, or feeding behavior. However, stim-1(RNAi) worms failed to lay eggs and were completely sterile (Fig. 3).
Somatic cells of rrf-1(pk1417) mutant worms are resistant to dsRNA, but their germline shows an apparently normal RNAi response (Sijen et al., 2001). To determine whether the sterility phenotype was due to disruption of somatic versus germline cell function, we fed rrf-1(pk1417) L1 larvae STIM-1 dsRNA-expressing bacteria and analyzed brood size. As shown in Fig. 3, stim-1 RNAi had no significant (P > 0.05) effect on brood size in rrf-1(pk1417) mutant worms. This indicates that the sterility induced by STIM-1 dsRNA is due largely to dysfunction of somatic cells.
Adult C. elegans hermaphrodites possess two U-shaped gonad arms connected via spermatheca to a common uterus. Oocytes form in the proximal gonad and accumulate in a single-file row of graded developmental stages. Developing oocytes remain in diakinesis of prophase I until they reach the most proximal position in the gonad arm where they undergo meiotic maturation and are then ovulated into the spermatheca for fertilization (for review see Hubbard and Greenstein, 2000).
Oocytes are surrounded by myoepithelial sheath cells (Hall et al., 1999). Prior to ovulation, sheath cells contract weakly at a basal rate of seven to eight contractions/min (McCarter et al., 1999). Release of the EGF-like protein LIN-3 from the maturing oocyte induces ovulation by increasing the rate and force of sheath cell contractions and by triggering opening of the gonad-spermatheca valve (Iwasaki et al., 1996; McCarter et al., 1999; Yin et al., 2004). The contractile activity of both the sheath cells (Yin et al., 2004) and spermatheca (Clandinin et al., 1998; Bui and Sternberg, 2002; Kariya et al., 2004) is regulated by IP3 and Ca2+ signaling.
To determine whether STIM-1 RNAi disrupted sheath cell and/or spermatheca function, we imaged anesthetized worms by DIC microscopy. The gonads of adult worms that had undergone several ovulation attempts were filled with endomitotic oocytes (e.g., Fig. 4 C) that were often stacked on top of one another, causing grossly distorted gonad morphology. We therefore isolated very young adult worms, which allowed us to image the first or second ovulation attempt in the absence of gonad morphology defects.
Fig. 4 shows sheath cell contractile activity in worms fed GFP dsRNA-producing bacteria as a control. The mean rate of basal sheath cell contraction measured at −5 min was six contractions/min (Fig. 4 A) with a mean displacement of 1.1 μm (Fig. 4 B). During ovulation, sheath contraction increased significantly (P < 0.0001) to a peak rate of 15 contractions/min (Fig. 4 A). The force of contractions also increased significantly (P < 0.0001), as indicated by an increase in mean displacement to 2.1 μm (Fig. 4 B).
Sheath cell contractility was greatly reduced in stim-1(RNAi) worms. The mean basal and peak ovulatory rates of sheath contraction were 2.5 contractions/min and 6 contractions/min (Fig. 4 A). Both rates were significantly (P < 0.0001) different from those observed in GFP RNAi control worms. During ovulation, the force of the sheath contraction increased slightly, but not significantly (P > 0.07); mean basal and ovulatory sheath displacement were 0.9 μm and 1.3 μm, respectively (Fig. 4 B). Taken together, these results indicate that STIM-1 plays an essential role in regulating sheath cell contractile activity. Loss of STIM-1 activity reduces the rate and force of sheath contraction.
We also noted that spermatheca function was defective in stim-1(RNAi) worms. During ovulation, the gonad- spermatheca valve opens, allowing the contracting sheath cells to pull the spermatheca over the maturing oocyte (Hubbard and Greenstein, 2000). In all 12 young adult stim-1(RNAi) worms examined, the gonad-spermatheca valve failed to open during ovulation attempts. Maturing oocytes were therefore trapped in the gonad where they underwent endomitosis (Fig. 4 C).
To further examine the role of STIM-1 in gonad function, we mutated aspartate residues at positions 55 and 57 to alanine (i.e., D55A and D57A). These residues are located in the predicted Ca2+ binding EF-hand domain (Fig. 1). Mutation of the analogous amino acids in Drosophila and human STIM1 homologues constitutively activates SOCE (Liou et al., 2005; Zhang et al., 2005) and ICRAC (Spassova et al., 2006). STIM-1(D55A;D57A) was fused to GFP and expression was driven by 1.9 kb of the stim-1 promoter.
Worms expressing STIM-1∷GFP exhibited normal fertility (Fig. 5 A). However, STIM-1(D55A;D57A)∷GFP animals were sterile. This sterility was suppressed by feeding the worms GFP dsRNA-producing bacteria (Fig. 5 A). We did not detect developing embryos or unfertilized oocytes in the uteri of STIM-1(D55A;D57A)∷GFP worms, indicating that the fertility defect is due at least in part to defects in ovulation.
In addition to sterility, STIM-1(D55A;D57A)∷GFP worms also exhibited a number of other defects. The worms grew more slowly than wild-type animals and their gonads appeared to be stunted and contained endomitotic oocytes (Fig. 5 B). Fluid accumulation was observed in the pseudocoel (Fig. 5 C), suggesting that the EF-hand mutation disrupts whole animal osmoregulation. Large vacuoles were present in the uterus (Fig. 5 D). Smaller vacuoles were observed next to the pharynx (Fig. 5 E) in a location similar to that of STIM-1∷GFP-expressing neurons (Fig. 2 B). The vacuoles may be indicators of cell death (e.g., Xu et al., 2001; Bianchi et al., 2004) induced by constitutive activation of STIM-1 and presumably SOCE, which in turn likely disrupts cellular Ca2+ homeostasis.
STIM-1 Regulates SOC Channel Activity in Intestinal Epithelial Cells but Is Not Required for IP3-dependent Oscillatory Ca2+ Signaling
The C. elegans digestive tract consists of a pharynx, intestine, and rectum. Food is pumped into the pharynx, ground up, and then moved into the intestine for further digestion and nutrient absorption. The intestine is comprised of 20 epithelial cells with extensive apical microvilli. Intestinal epithelial cells secrete digestive enzymes, absorb nutrients, and store lipids, proteins and carbohydrates (White, 1988; Leung et al., 1999; Ashrafi et al., 2003).
Defecation in C. elegans is a highly rhythmic process that occurs once every 45–50 s when nematodes are feeding and is mediated by sequential contraction of the posterior body wall muscles, anterior body wall muscles, and enteric muscles (Iwasaki and Thomas, 1997). Posterior body wall muscle contraction (pBoc) is controlled by IP3-dependent Ca2+ oscillations in intestinal epithelial cells (Dal Santo et al., 1999; Espelt et al., 2005; Teramoto and Iwasaki, 2006).
Oscillatory Ca2+ signaling is thought to be critically dependent on SOCE (Parekh and Penner, 1997; Venkatachalam et al., 2002; Parekh and Putney, 2005). We therefore examined the effect of stim-1 knockdown on the defecation cycle. Surprisingly, stim-1(RNAi) worms had normal defecation rhythms. Mean pBoc periods in control and stim-1(RNAi) worms were 51 s and 52 s, respectively, and were not significantly (P > 0.3) different (Fig. 6 A). The mean coefficient of variance (CV), which is a measure of pBoc cycle rhythmicity (Espelt et al., 2005), was 4.5% in control worms (Fig. 6 B). The low CV indicates that the pBoc cycle is highly rhythmic. Mean pBoc CV was 5.6% in stim-1(RNAi) worms and was not significantly (P > 0.1) different from that of control animals (Fig. 6 B).
Calcium levels in the ER lumen regulate Ca2+ flux through the IP3R (for review see Burdakov et al., 2005), which in turn modulates the characteristics of intracellular Ca2+ oscillations and waves. Mutations in the IP3R that alter channel activity and regulation could conceivably sensitize IP3R Ca2+ flux to reductions in ER Ca2+ levels brought about by a loss of SOCE. We therefore examined the effect of stim-1 RNAi on worm strains harboring IP3R mutations.
A single gene, itr-1, encodes the IP3 receptor in C. elegans (Baylis et al., 1999; Dal Santo et al., 1999). sa73 is a loss-of-function itr-1 allele and sy290 and sy327 are gain-of-function alleles. The sa73 mutation is located in the modulatory region of ITR-1 close to a putative Ca2+ binding domain (Dal Santo et al., 1999). sy290 and sy327 were isolated as dominant mutations that suppress or “rescue” the phenotype induced by loss-of-function mutations in upstream IP3 signaling components (Clandinin et al., 1998). sy290 is an arginine to cysteine substitution at residue 511, which is located in the IP3 binding domain (Clandinin et al., 1998). This mutation increases in vitro binding affinity for IP3 approximately twofold (Walker et al., 2002). sy327 substitutes leucine 899 with phenylalanine in a putative Ca2+ binding domain (unpublished data). As shown in Fig. 6, stim-1 RNAi had no effect on pBoc period or rhythmicity in any of these mutants.
lfe-2 and ipp-5 encode an IP3 kinase and phosphatase, respectively (Clandinin et al., 1998; Bui and Sternberg, 2002). Loss-of-function mutations in these genes elevate intracellular IP3 levels by inhibiting conversion of IP3 into IP4 or IP2 (e.g., Clandinin et al., 1998; Kariya et al., 2004; Espelt et al., 2005). Increased IP3 levels in turn should increase IP3R activity and store Ca2+ release. In the absence of refilling mechanisms, enhanced Ca2+ release may lead to store depletion with subsequent disruption of intracellular Ca2+ signals. The effect of stim-1 RNAi should therefore be enhanced in lfe-2 and ipp-5 mutants if SOCE is required for refilling intestinal ER Ca2+ stores. However, as shown in Fig. 6, STIM-1 knockdown had no effect on pBoc period or rhythmicity in lfe-2 and ipp-5 mutants.
The absence of an effect of stim-1 RNAi on pBoc in wild type and IP3 signaling mutants suggests that STIM-1/SOCE does not play a role in generating intestinal Ca2+ oscillations. To more directly examine the role of stim-1 in Ca2+ signaling, we used fluo-4 to monitor Ca2+ oscillations and waves in isolated intestines. Fig. 7 shows examples of Ca2+ oscillations in a control intestine and an intestine isolated from a stim-1(RNAi) worm. The characteristics of intestinal Ca2+ oscillations and waves are shown in Table I. stim-1 RNAi had no significant (P > 0.1) effect on oscillation period, oscillation rise and fall time, and Ca2+ wave velocity.
As a final test for the role of stim-1 in intestinal Ca2+ signaling, we measured Ca2+ oscillations in intact, freely moving worms using the FRET-based Ca2+ indicator protein cameleon (Truong et al., 2001). To maximize stim-1 knockdown, we selectively rescued rde-1 in the intestines of rde-1(ne219) loss-of-function mutant worms (see Espelt et al., 2005). rde-1 (RNAi defective) encodes a protein involved in translation initiation (Tabara et al., 1999; Fagard et al., 2000) and rde-1 loss-of-function mutants are strongly resistant to RNAi induced by dsRNA injection, feeding, or expression (Tabara et al., 1999). Selective rescue of rde-1 in the intestine allows normal fertility to be maintained in worms that are fed stim-1 dsRNA-producing bacteria for multiple successive generations. Fig. 7 (C and D) shows examples of Ca2+ oscillations in a worm fed stim-1 dsRNA-producing bacteria for three generations and a control worm fed bacteria containing empty feeding vector. The mean ± SEM oscillation periods in control and stim-1(RNAi) worms were 56 ± 4 s (n = 3) and 57 ± 2 s (n = 3), respectively, and were not significantly (P > 0.8) different.
The absence of an effect of stim-1 RNAi on intestinal Ca2+ signaling could be due to ineffective knockdown of STIM-1 expression. To address this issue, we examined STIM-1 expression levels in the anterior and posterior intestines of STIM-1∷GFP transgenic worms fed stim-1 dsRNA-producing bacteria. Fig. 8 A shows fluorescence micrographs of the anterior and posterior intestines of wild-type worms, STIM-1∷GFP transgenic worms, and transgenic worms fed stim-1 dsRNA–producing bacteria for 36 h. stim-1 RNAi dramatically reduced GFP fluorescence in both intestinal regions.
To quantify the effect of stim-1 RNAi, we measured maximal mean pixel intensity in a 28-μm-wide by 18-μm-high region. stim-1 RNAi significantly (P < 0.002) reduced anterior and posterior intestine pixel intensity by ∼15- and ∼19-fold, respectively (Fig. 8 B). Maximal mean pixel intensity in stim-1(RNAi) worms was approximately twofold higher than autofluorescence levels detected in wild-type worms. Given that transgenes are typically overexpressed compared with endogenous genes, these results demonstrate that stim-1 RNAi is highly effective in reducing STIM-1 protein expression.
We also monitored the effect of stim-1 RNAi on intestinal cell SOC channel activity. Cultured intestinal cells express a SOC channel current with many of the same biophysical characteristics as ICRAC (Estevez et al., 2003). Fig. 9 A shows typical activation of ISOC in a control intestinal cell patch clamped with a pipette solution containing 10 μM IP3, 10 mM BAPTA, and a free Ca2+ concentration of ∼18 nM. Treatment of cultured intestinal cells with stim-1 dsRNA for 2–3 d eliminated ISOC in 7 of 10 patch-clamped cells (Figs. 9, B and C). Mean ± SEM peak ISOC was reduced nearly 18-fold (P < 0.02) from −19.4 ± 6.2 pA/pF (n = 11) in control cells to −1.1 ± 0.7 pA/pF (n = 10) in cells exposed to stim-1 dsRNA (Fig. 9 C). These results indicate that, as in mammalian and Drosophila cells (Liou et al., 2005; Roos et al., 2005; Zhang et al., 2005; Spassova et al., 2006), STIM-1 is a modulator of SOC channels.
Transgenic worms expressing STIM-1(D55A;D57A)∷GFP had an increased pBoc cycle time and exhibited striking pBoc arrhythmia (Fig. 10 A, top, and Fig. 10 B). Feeding STIM-1(D55A;D57A)∷GFP worms for 52–55 h with bacteria producing stim-1 dsRNA dramatically suppressed the effect of the EF-hand mutant on pBoc period and rhythmicity (Fig. 10 A, bottom, and Fig. 10 B). Mean pBoc period and CV in STIM-1(D55A;D57A)∷GFP;stim-1(RNAi) animals were 46 s and 7.3%, respectively, and were not significantly (P > 0.05) different from wild-type worms (see Fig. 6). Taken together, the electrophysiological, STIM-1∷GFP expression, and pBoc data (Figs. 8–10,9) demonstrate clearly that stim-1 RNAi is highly effective in disrupting both STIM-1 expression and SOC activity in the worm intestine.
STIM-1 Is Not Required for Intestinal Cell ER Ca2+ Homeostasis under Normal Physiological Conditions
ER Ca2+ homeostasis is not only important for intracellular Ca2+ signaling, but also for proper protein synthesis and folding. Disruption of ER Ca2+ homeostasis triggers the UPR. The UPR is an intracellular signaling and transcriptional/translational program activated by the accumulation of unfolded proteins in the ER lumen (for reviews see Rao et al., 2004; Schroder and Kaufman, 2005). In mammalian cells, inhibition of the sarcoplasmic/ER Ca2+ ATPase (SERCA) with thapsigargin depletes ER Ca2+ stores and triggers the UPR (e.g., Hong et al., 2004; Durose et al., 2006). We reasoned that if STIM-1 and SOCE were essential for refilling ER Ca2+ stores during oscillatory Ca2+ signaling, then stim-1 RNAi should induce the UPR in the worm intestine.
To monitor the intestinal UPR, we used a transgenic worm strain expressing an hsp-4 transcriptional GFP reporter. hsp-4 encodes a C. elegans homologue of the ER chaperone protein GRP78/BiP and is induced by ER stress (Calfon et al., 2002). As shown in Fig. 11 and as described previously (Calfon et al., 2002), hsp-4∷GFP is expressed at low levels in the intestine under control conditions. Exposure of worms for 6 h to agar containing 10 μg/ml tunicamycin, which induces ER stress, caused significant (P < 0.001) and striking up-regulation of intestinal hsp-4∷GFP expression (Fig. 11; see also Calfon et al., 2002).
To determine whether store Ca2+ depletion activates the UPR in C. elegans, we inhibited SERCA by feeding worms sca-1 dsRNA–producing bacteria for 30 h. sca-1 encodes the C. elegans SERCA homologue (Zwaal et al., 2001). As shown in Fig. 11 B, sca-1 RNAi caused a nearly threefold increase (P < 0.001) in hsp-4∷GFP expression. However, hsp-4∷GFP expression was not significantly (P > 0.05) altered by stim-1 RNAi (Fig. 11 B).
In a separate series of experiments, we examined the effect of combined sca-1 and stim-1 RNAi on the UPR. As shown in Fig. 11 C, knockdown of both proteins together significantly (P < 0.0001) increased whole worm fluorescence ∼33% compared with sca-1 alone. Similar results were observed in a second independent experiment (mean ± SEM relative fluorescence values in sca-1(RNAi) and sca-1(RNAi);stim-1(RNAi) worms were 3.2 ± 0.3, n = 21 and 4.2 ± 0.2, n = 121, respectively; P < 0.008). Based on the results shown in Figs. 6–11,7891011, we conclude that STIM-1 and SOCE do not play significant roles in intestinal cell oscillatory Ca2+ signaling or ER Ca2+ homeostasis under normal physiological conditions. However, the effects of combined sca-1 and stim-1 RNAi on the UPR suggest that SOCE may contribute to the regulation of store Ca2+ levels during experimental manipulations that induce extreme store Ca2+ depletion.
C. elegans STIM-1 overall shares ∼21% amino acid identity with human STIM1 (Fig. 1). The strongest conservation of primary structure occurs in the EF-hand and SAM domains. SAM domains play important roles in mediating protein–protein interactions that regulate the activity, localization, and assembly of numerous proteins (Qiao and Bowie, 2005). EF-hands play well-established roles as Ca2+ binding sites (Strynadka and James, 1989). Mutagenesis studies indicate that the EF-hand of STIM homologues is likely responsible for monitoring ER lumen Ca2+ levels (Liou et al., 2005; Zhang et al., 2005; Spassova et al., 2006). Expression of the STIM-1 EF-hand D55A;D57A mutant under the control of the stim-1 promoter causes sterility (Fig. 5 A) and pBoc arrhythmia (Fig. 10) as well as morphological abnormalities (Figs. 5, B–E) that are consistent with disruption of Ca2+ signaling and possible Ca2+-induced cellular injury.
Interestingly, when we expressed STIM-1(D55A;D57A)∷GFP under the control of the promoter for let-858, a gene that functions in most C. elegans cell types (Kelly et al., 1997), worms showed numerous serious defects and survived very poorly (unpublished data). This suggests that STIM-1 and SOC channels function in more cell types than suggested by GFP reporter studies (Fig. 2) and/or that the EF-hand mutant has cellular effects in addition to SOCE activation. The absence of defects induced by stim-1 RNAi other than gonad dysfunction does not preclude additional physiological roles for the gene. For example, stim-1 RNAi–induced sterility (Fig. 3) could obscure a role for STIM-1 in embryonic development. A role for stim-1 in the C. elegans nervous system may have gone undetected in our studies given the relative insensitivity of neurons to RNAi (Simmer et al., 2002). Finally, stim-1 RNAi could give rise to subtle phenotypes not detected in our assays. It will be valuable in future studies to isolate stim-1 deletion mutants and perform more extensive physiological and behavioral analyses.
STIM-1 plays an essential role in regulating IP3-dependent sheath cell and spermatheca contractile activity required for fertility (Figs. 3 and 4). However, despite striking knockdown of STIM-1 expression (Figs. 8 and 10) and intestinal SOC channel activity (Fig. 9), stim-1 RNAi surprisingly has no effect on oscillatory Ca2+ signaling in the intestine (Figs. 6 and 7 and Table I) or on intestinal ER Ca2+ homeostasis (Fig. 11). These results can be explained if only a very nominal amount of STIM-1/SOC channel activity, which may remain after RNAi treatment, is required to maintain ER Ca2+ levels. Other explanations though are also possible and worth considering. It is widely assumed that SOCE is an essential component of intracellular Ca2+ signaling events and is required for maintaining ER Ca2+ levels (Parekh and Penner, 1997; Venkatachalam et al., 2002; Parekh and Putney, 2005). With the exception of immune cells (for review see Lewis, 2001), SOCE and CRAC activation in most cell types has not been observed under physiologically relevant conditions, but only under conditions of extreme store depletion experimentally induced by SERCA inhibition, supraphysiological IP3R activation, exposure to high concentrations of ionomycin, and/or increases in cytoplasmic Ca2+ buffering (e.g., Parekh et al., 1997; Golovina et al., 2001; Machaca, 2003). Furthermore, direct demonstration of store Ca2+ depletion during intracellular Ca2+ oscillations induced by physiologically relevant stimuli is lacking. In the one detailed study conducted to date, little or no change in store Ca2+ levels was detected during acetylcholine-induced Ca2+ oscillations in pancreatic acinar cells. Store depletion was only observed upon supraphysiological acetylcholine stimulation (Park et al., 2000). These results indicate that in acinar cells, SOCE is likely not activated during normal oscillatory Ca2+ signaling events. One caveat of these studies is that store depletion may not have been observed if it occurred in ER microdomains that were not resolved by the imaging methods used. However, photobleaching and Ca2+ uncaging experiments suggested that the acinar cell ER is a continuous compartment and that Ca2+ released from microdomains is rapidly replenished by Ca2+ in the bulk ER (Park et al., 2000).
In the worm intestine, it is unclear why SOC channels would be needed to refill stores during oscillatory Ca2+ signaling. Patch clamp studies on cultured intestinal cells have demonstrated the presence of a highly Ca2+ selective channel, ORCa, that is constitutively active and has biophysical properties resembling those of certain TRPM channels (Estevez et al., 2003; Estevez and Strange, 2005). This channel is likely encoded by the TRPM6/7 homologues gon-2 and gtl-1 (Teramoto et al., 2005; unpublished data), both of which are required for maintaining normal intestinal Ca2+ oscillations (unpublished data) and pBoc rhythm (Teramoto et al., 2005; unpublished data). Calcium influx through the ORCa channel raises global intracellular Ca2+ levels (Estevez and Strange, 2005), and this Ca2+ in turn should be available for store refilling. Indeed, in any cell type, non-SOC channel Ca2+ entry pathways that are active during Ca2+ signaling events could function to refill stores as well as modulate the characteristics of Ca2+ signals. A case-in-point is the arachidonic acid–regulated Ca2+ channel (ARC) (Mignen and Shuttleworth, 2000). Shuttleworth has argued that ARC is the predominant mode of Ca2+ entry during stimulation of certain nonexcitable cell types with physiologically relevant concentrations of agonists and that SOCE is operational only with supraphysiological stimulation (e.g., Shuttleworth, 1999; Shuttleworth and Mignen, 2003).
The widespread existence of SOCE in numerous cell types supports the notion that it must be essential for Ca2+ signaling. However, our findings in the C. elegans intestine and the arguments outlined above raise the question of whether SOCE is ubiquitously indispensable for the generation and maintenance of Ca2+ signals. If it is not indispensable, why are SOC channels expressed in the intestine (Fig. 9; Estevez et al., 2003) and why has SOCE been so widely observed? One possibility is that the primary function of SOCE is to provide cells with a failsafe mechanism for protecting store Ca2+ levels during pathophysiological insults and stress conditions. For example, certain viral proteins (Tian et al., 1995; van Kuppeveld et al., 1997) and bacterial toxins (Bryant et al., 2003; Saha et al., 2005) as well as cellular stressors including ischemia (Lehotsky et al., 2003) and oxidants (Henschke and Elliott, 1995; Pariente et al., 2001) induce store Ca2+ loss and depletion. Failure to maintain store Ca2+ levels under pathophysiological and stress conditions can exacerbate injury by disrupting ER protein synthesis and processing and lead ultimately to cell death (Rao et al., 2004; Schroder and Kaufman, 2005). Consistent with this idea, we observed that stim-1 RNAi had no effect on the intestinal UPR under normal physiological conditions (Fig. 11). However, knockdown of STIM-1 expression enhanced the UPR in worms where Ca2+ stores were depleted by sca-1 (i.e., SERCA) RNAi (Fig. 11 C and Results).
Recent identification of the gene Orai1 (Feske et al., 2006; see also Vig et al., 2006; Zhang et al., 2006) supports the hypothesis that SOCE may not be an essential component of all Ca2+ signaling events. Severe combined immunodeficiency (SCID) is a disease caused by defects in immune cell function (Huang and Manton, 2005). A subset of SCID patients has been shown to have defective T cell, B cell, and fibroblast SOCE and T cell ICRAC activation (Partiseti et al., 1994; Le Deist et al., 1995; Feske et al., 2001, 2005). As discussed below, SOCE plays an essential role in T cell activation by antigens (Lewis, 2001). In an elegant study, Feske et al. (2006) used linkage analysis and a Drosophila S2 cell genome-wide RNAi screen to identify Orai1 as an essential component of the CRAC channel and as the gene mutated in SCID patient immune cells lacking SOCE. Accumulating evidence suggests that Orai1 likely encodes the CRAC channel itself (Mercer et al., 2006; Peinelt et al., 2006; Soboloff et al., 2006b; Zhang et al., 2006).
Interestingly, in the SCID patients studied by Feske et al. (2000), the only nonimmunological defects observed were nonprogressive muscle hypotonia and mild psychomotor and mental retardation. This finding is unexpected if SOCE plays a ubiquitous and essential role in Ca2+ signaling. It is possible that the human Orai1 homologues, Orai2 and Orai3, function as SOC channels in cell types other than T cells, B cells, and fibroblasts. However, only a single Orai homologue is present in the C. elegans genome. Our ongoing studies have demonstrated that RNAi knockdown of this gene has no effect on pBoc rhythm but phenocopies the sterility defect and sheath cell and spermatheca dysfunction seen with stim-1 RNAi (Figs. 3 and 4) (unpublished data). Sterility induced by knockdown of the C. elegans Orai1 homologue has also been observed in genome-wide RNAi screens (Maeda et al., 2001; Rual et al., 2004).
While it is unclear whether SOCE plays an essential role in all Ca2+ signaling pathways, it is certainly critical for some physiological processes. A role for SOCE in immune cell activation is well established (for review see Lewis, 2001). Stimulation of antigen receptors with antigenic peptides triggers cell replication and a gene transcription program required for the immune response. Antigens induce intracellular Ca2+ oscillations that are strictly dependent on extracellular Ca2+ influx via CRAC. Evidence suggests that CRAC does more than simply refill stores and may function directly in the generation of Ca2+ oscillations (Dolmetsch and Lewis, 1994). The characteristics of Ca2+ oscillations in turn increase the efficiency and specificity of gene expression and enhance the detection of low levels of antigens (Dolmetsch et al., 1997, 1998).
In C. elegans, STIM-1 and presumably SOCE are essential for regulating the contractile activity of gonad sheath cells and the spermatheca (Fig. 4). The precise functions of SOC channels in these tissues are uncertain. However, it is well established that IP3 signaling regulates their contractile activity (Clandinin et al., 1998; Bui and Sternberg, 2002; Kariya et al., 2004; Yin et al., 2004). Sheath cell and spermatheca contractile activity may require relatively high and prolonged Ca2+ elevation that is dependent on concomitant activation of both store Ca2+ release and influx across the plasma membrane. Activation of SOCE in these tissues may be facilitated by limited Ca2+ stores that become rapidly depleted during IP3-regulated Ca2+ release. It is also possible that the functional properties of SOC channels are uniquely suited to Ca2+ signaling events in the gonad. For example, we have shown previously that a ClC-type anion channel, CLH-3b, is activated in worm oocytes undergoing meiotic maturation and that the channel functions to regulate contraction of surrounding gap junction–coupled sheath cells (Rutledge et al., 2001; Denton et al., 2004). Loss of CLH-3b activity by RNAi (Rutledge et al., 2001) or deletion mutagenesis (unpublished data) causes premature initiation of ovulatory sheath contractions. We have suggested that CLH-3b–induced depolarization of the oocyte and surrounding sheath cells may function to inhibit contraction by inhibiting Ca2+ entry through a store-operated CRAC-like channel (Rutledge et al., 2001; Yin et al., 2004). The very high Ca2+ selectivity and the lack of voltage-dependent gating of CRAC channels (Lewis, 2001; Parekh and Putney, 2005) facilitate modulation of CRAC-mediated Ca2+ influx by membrane voltage changes regulated by other ion channels. This in turn provides mechanisms for increasing the complexity and thus information content of Ca2+ signals.
In summary, we have identified a C. elegans STIM1 homologue, have demonstrated that it is required for SOC channel activity in intestinal cells, and have shown that it plays essential roles in some but not all IP3-regulated physiological processes. The exact role of SOCE and SOC channels in the maintenance of store Ca2+ levels and in the regulation of Ca2+ signaling events in most cell types, with the notable exception of immune cells, is unclear. The breakthrough discoveries of STIM-1 (Roos et al., 2005) and Orai1 (Feske et al., 2006; Vig et al., 2006; Zhang et al., 2006) along with development of appropriate animal and cell models makes it possible to address key questions in the field, including the specific functional roles of SOCE under physiological and pathophysiological conditions and the molecular mechanisms of SOC channel regulation. Combining powerful forward and reverse genetic analyses with direct physiological measurements of Ca2+ signaling and ion channel activity in C. elegans should be particularly useful in this regard. A detailed molecular and integrative physiological understanding of SOCE is essential for understanding the role of SOC channels in disease and for the potential development of pharmacological strategies to modulate their activity.
We thank Dr. Joel Rothman (University of California at Santa Barbara, Santa Barbara, CA) for providing the elt-2∷GFP-expressing worm strain. Other strains used in this work were provided by the Caenorhabditis Genetics Center (University of Minnesota, Minneapolis, MN). Confocal microscopy was performed in the Vanderbilt University Medical Center Cell Imaging Shared Resource, which is supported by National Institutes of Health (NIH) grants CA68485, DK20593, DK58404, HD15052, DK59637, and EY08126.
This work was supported by NIH grants GM74229 and DK51610 to K. Strange, and HL08010 to K. Nehrke. T. Lamitina was supported by a postdoctoral fellowship from the National Kidney Foundation.
Olaf S. Andersen served as editor.
A.Y. Estevez's present address is Biology Department, St. Lawrence University, Canton, NY 13617.
T. Lamitina's present address is Department of Physiology, University of Pennsylvania, Philadelphia, PA 19104.
Abbreviations used in this paper: CV, coefficient of variance; DIC, differential interference contrast; FRET, fluorescence resonance energy transfer; ICRAC, Ca2+ release–activated Ca2+ current; IP3, 1,4,5-trisphosphate; IP3R, IP3 receptor; SCID, severe combined immunodeficiency; SERCA, sarcoplasmic/ER Ca2+ ATPase; SOCE, store-operated Ca2+ entry; STIM1, stromal interaction molecule 1; UPR, unfolded protein response.