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¹ Department of Anesthesiology, Department of Molecular Physiology and Biophysics, and Department of Pharmacology, Vanderbilt University Medical Center, Nashville, TN 37232
² Department of Medicine, Division of Nephrology, University of Rochester Medical Center, Rochester, NY 14642

Correspondence to Kevin Strange: kevin.strange{at}vanderbilt.edu

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
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.

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; I_CRAC, 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.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
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 I_CRAC, 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 I_CRAC (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

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
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 M. 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
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.

Microscopy
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 10x/0.3 N.A. and 40x/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 10x 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 x 2 binning. The 480/540 nm emission ratio after background subtraction and signal threshholding was used as an indicator of FRET efficiency.

Statistical Analysis
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.

	RESULTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
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).

View larger version (32K): [in this window] [in a new window]   Figure 1. Sequence alignment of human and C. elegans STIM1 homologues. Yellow and green shading indicates sequence identity and conserved amino acid substitutions, respectively. Conserved domains are shown in black boxes. Red boxes show location of N-linked glycosylation sites in human STIM1 (Williams et al., 2002). Light blue shading shows N-terminal cysteine residues conserved in human and Drosophila STIM homologues. Percent similarity and identity of signal peptide, EF-hand, SAM, transmembrane, and coiled-coil domains are 50.0% and 16.7%, 58.3% and 50.0%, 50.7% and 37.7%, 27.8% and 5.6%, and 57.4% and 21.3%, respectively. Alignment was performed using Vector NTI software (InforMax). Protein domains were identified by SMART (http://smart.embl-heidelberg.de/smart/change_mode.pl).

View larger version (32K): [in this window] [in a new window]   Figure 2. STIM-1::GFP expression pattern. (A) Low magnification confocal image of a whole worm. STIM-1::GFP expression is present throughout the intestine, but is more pronounced in the anterior (AI) and posterior intestine (PI). Expression is also present in the spermatheca (Sp) and gonadal sheath cells (SC). Bar, 100 µm. (B) Expression of STIM-1::GFP in head neurons. Arrows and arrowheads point to cell bodies and dendrites, respectively. Bar, 20 µm. (C) Series of confocal Z-sections through the anterior intestine. STIM-1::GFP appears to be localized to a membrane or submembrane region as well as punctate and presumably intracellular domains. Lu, intestinal lumen. Bar, 20 µm. (D) Series of confocal Z-sections through the posterior intestine. STIM-1::GFP appears to be localized to a membrane and/or submembrane region and a prominent intracellular reticular structure that resembles the intestinal endoplasmic reticulum (Rolls et al., 2002). Bar, 10 µm. (E) Series of confocal Z-sections through the spermatheca (Sp) and gonadal sheath cells (SC). Punctate localization pattern of STIM-1::GFP can be seen in sheath cells in the second and third panels. Bar, 10 µm.

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).

View larger version (10K): [in this window] [in a new window]   Figure 3. Effect of stim-1 RNAi on fertility. Brood size is defined as the total number of progeny produced over 4 d. rrf-1 encodes an RNA-directed RNA polymerase homologue required for RNAi in somatic but not germ cells. pk1417 is a predicted rrf-1 null allele (Sijen et al., 2001). Values are means ± SEM (n = 5–8).

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.

View larger version (61K): [in this window] [in a new window]   Figure 4. Sheath cell and spermatheca function in control (GFP RNAi) and stim-1(RNAi) worms. (A) Rates of sheath contraction during a single ovulatory cycle. Time 0 is defined as the time at which ovulation was completed in control worms. stim-1(RNAi) worms never ovulated (see Results). Therefore in these animals, time 0 is defined as the first time point after peak sheath contraction rate was observed. Values are means ± SEM (n = 9–10). (B) Sheath cell displacement under basal conditions and during ovulation. Basal and ovulatory displacement were measured between –15 and –5 min and between –2 and 0 min, respectively. Values are means ± SEM (n = 6–9). *, P < 0.0001 compared with control ovulatory displacement. (C) Differential interference contrast micrographs of control and stim-1 RNAi worms. Spermatheca fails to open during ovulation in stim-1(RNAi) worms and oocytes are trapped in proximal gonad where they undergo endomitosis. Em, embryo; Emo, endomitotic oocytes; Oo, oocytes; Sp, spermatheca; Ut, uterus. Note that the uterus in the stim-1(RNAi) worm is empty, whereas the uterus in the control animal contains three developing embryos.

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 I_CRAC (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.

View larger version (77K): [in this window] [in a new window]   Figure 5. Fertility and morphology defects in transgenic worms expressing the STIM-1 EF-hand mutant D55A;D57A. (A) Brood size in wild type and STIM-1 transgenic worms. Brood size is defined as the total number of progeny produced over 4 d. Wild type;gfp(RNAi), wild-type worms fed GFP dsRNA–producing bacteria. STIM-1::GFP, transgenic worms expressing STIM-1 GFP fusion protein. D55A;D57A::GFP, transgenic worms expressing STIM-1 EF-hand mutant GFP fusion protein. D55A;D57A::GFP;gfp(RNAi), transgenic worms expressing STIM-1 EF-hand mutant GFP fusion protein fed GFP dsRNA–producing bacteria. Values are means ± SEM (n = 6–11). *, P < 0.001 compared with Wild type;gfp(RNAi), STIM-1::GFP and D55A;D57A::GFP;gfp(RNAi) worms. Brood size of D55A;D57A::GFP;gfp(RNAi) worms was not significantly (P > 0.05) different from Wild type:gfp(RNAi) animals. (B–E) Differential interference contrast micrographs of transgenic worms expressing STIM-1(D55A;D57A)::GFP. Gonads of STIM-1(D55A;D57A)::GFP worms appeared to be stunted and contained endomitotic oocytes (B). Fluid accumulation was observed in the pseudocoel (C, arrowheads) and large vacuoles were present in the uterus (D). Smaller vacuoles were observed next to the pharynx (E, arrow) in a location similar to that of STIM-1::GFP-expressing neurons (see Fig. 2 B). DG, distal gonad; Emo, endomitotic oocytes; Oo, oocytes; Phx, pharynx; Ut, uterus.

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).

View larger version (14K): [in this window] [in a new window]   Figure 6. Effect of stim-1 RNAi on pBoc period (A) and coefficient of variance (B), which is a measure of cycle rhythmicity in wild type and IP3 signaling mutant worms. itr-1(sa73) is a loss-of-function allele and itr-1(sy290) and itr-1(sy327) are gain-of-function alleles of the C. elegans IP3R. lfe-2(sy326) and ipp-5(sy605) are loss-of-function alleles of an IP3 kinase and IP3 phosphatase, respectively. Values are means ± SEM (n = 5–19).

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.

View larger version (22K): [in this window] [in a new window]   Figure 7. Examples of intestinal Ca2+ oscillations. (A and B) Calcium oscillation imaging in dissected intestines loaded with fluo-4. Intestines were isolated from wild-type worms fed either GFP (A) or stim-1 (B) dsRNA–producing bacteria. Fluorescence images were acquired during the last two oscillations shown in B at 1 Hz. All other images were acquired at 0.2 Hz. (C and D) In vivo Ca2+ oscillation imaging in intestines of freely moving L3 larvae expressing the FRET-based Ca2+ indicator protein cameleon. Worms were fed bacteria containing the empty RNAi feeding vector (C) or stim-1 dsRNA–producing bacteria (D) for three successive generations.

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.

View larger version (34K): [in this window] [in a new window]   Figure 8. Effect of stim-1 RNAi on STIM-1::GFP expression. (A) Fluorescence micrographs of anterior and posterior intestines from wild type and STIM-1::GFP worms and STIM-1::GFP worms fed stim-1 dsRNA–producing bacteria. Images were obtained using a CCD camera and inverted microscope (see Materials and methods). Camera settings were the same for all three groups of worms. Weak fluorescence in wild-type worms is autofluorescence from intestinal granules. (B) Maximal mean pixel intensity in anterior and posterior intestines. Mean pixel intensity was measured in a region 28 µm wide by 18 µm high. The measuring region was positioned over the anterior or posterior intestine at a point where mean pixel intensity was maximal. Values are means ± SEM (n = 7–9). *, P < 0.002 compared with STIM-1::GFP worms.

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 I_CRAC (Estevez et al., 2003). Fig. 9 A shows typical activation of I_SOC 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 I_SOC in 7 of 10 patch-clamped cells (Figs. 9, B and C). Mean ± SEM peak I_SOC 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.

View larger version (10K): [in this window] [in a new window]   Figure 9. Effect of stim-1 RNAi on cultured intestinal cell SOC current. Examples of whole cell currents induced by store depletion in a control cell (A) and a cell treated with stim-1 dsRNA (B). Store depletion was induced using a pipette solution containing 10 µM IP3, 10 mM BAPTA, and 18 nM free Ca2+. Currents were elicited by ramping membrane voltage from –120 mV to +80 mV at 200 mV/s every 5 s. (C) Effect of stim-1 dsRNA on peak ISOC measured 5 min after obtaining whole cell access. Solid lines are the mean currents for the cells shown.

View larger version (14K): [in this window] [in a new window]   Figure 10. Effect of expression of stim-1 EF-hand mutation on pBoc period and rhythmicity. (A) pBoc cycles of individual STIM-1 (D55A;D57A)::GFP worms fed bacteria containing the empty vector (top) or fed stim-1 dsRNA-producing bacteria (bottom). (B) Mean pBoc period and coefficients of variance for wild-type worms and worms shown in A. Wild type;gfp(RNAi), wild-type worms fed GFP dsRNA–producing bacteria. D55A;D57A::GFP, transgenic worms expressing STIM-1 EF-hand mutant GFP fusion protein fed bacteria containing the empty feeding vector. D55A;D57A::GFP;stim-1(RNAi), transgenic worms expressing STIM-1 EF-hand mutant GFP fusion protein fed stim-1 dsRNA–producing bacteria. Values are means ± SEM (n = 8–9). Wild-type data are reproduced from Fig. 6. *, P < 0.001 compared with wild-type worms. **, P < 0.001 compared with D55A;D57A::GFP worms. pBoc period and coefficient of variance in D55A;D57A::GFP;stim-1(RNAi) worms were not significantly (P > 0.05) different from wild type;gfp(RNAi) worms.

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).

View larger version (40K): [in this window] [in a new window]   Figure 11. Induction of the intestinal unfolded protein response. (A) Fluorescent micrographs of control hsp-4::GFP worms fed bacteria containing the empty feeding vector, control hsp-4::GFP worms treated with tunicamycin, and hsp-4::GFP worms fed either sca-1 or stim-1 dsRNA–producing bacteria. Worms were imaged using a stereo dissecting microscope and CCD camera (see Materials and methods). Images were obtained with identical camera settings. (B) Changes in whole worm fluorescence induced by tunicamycin (Tun), sca-1 RNAi, or stim-1 RNAi. Values are means ± SEM (n = 275–732) and are plotted relative to fluorescence in control animals. *, P < 0.001 compared with control worms. (C) Effect of combined sca-1 and stim-1 RNAi on whole worm fluorescence. Values are means ± SEM (n = 242–756) and are plotted relative to fluorescence in control animals. *, P < 0.0001 compared with sca-1(RNAi) worms. Fluorescence levels in B and C were quantified using a COPAS BioSort and normalized to time-of-flight (i.e., GFP fluorescence/time-of-flight), which is a measure of worm size. Raw GFP fluorescence levels showed an identical pattern of change and statistical significance.

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, 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.

	DISCUSSION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
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).

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