Identification of Store-independent and Store-operated Ca²⁺ Conductances in Caenorhabditis elegans Intestinal Epithelial Cells

Abstract

The nematode Caenorhabditis elegans offers significant experimental advantages for defining the genetic basis of diverse biological processes. Genetic and physiological analyses have demonstrated that inositol-1,4,5-trisphosphate (IP₃)–dependent Ca²⁺ oscillations in intestinal epithelial cells play a central role in regulating the nematode defecation cycle, an ultradian rhythm with a periodicity of 45–50 s. Patch clamp studies combined with behavioral assays and forward and reverse genetic screening would provide a powerful approach for defining the molecular details of oscillatory Ca²⁺ signaling. However, electrophysiological characterization of the intestinal epithelium has not been possible because of its relative inaccessibility. We developed primary intestinal epithelial cell cultures that circumvent this problem. Intestinal cells express two highly Ca²⁺-selective, voltage-independent conductances. One conductance, I_ORCa, is constitutively active, exhibits strong outward rectification, is 60–70-fold more selective for Ca²⁺ than Na⁺, is inhibited by intracellular Mg²⁺ with a K_1/2 of 692 μM, and is insensitive to Ca²⁺ store depletion. Inhibition of I_ORCa with high intracellular Mg²⁺ concentrations revealed the presence of a small amplitude conductance that was activated by passive depletion of intracellular Ca²⁺ stores. Active depletion of Ca²⁺ stores with IP₃ or ionomycin increased the rate of current activation ∼8- and ∼22-fold compared with passive store depletion. The store-operated conductance, I_SOC, exhibits strong inward rectification, and the channel is highly selective for Ca²⁺ over monovalent cations with a divalent cation selectivity sequence of Ca²⁺ > Ba²⁺ ≈ Sr²⁺. Reversal potentials for I_SOC could not be detected accurately between 0 and +80 mV, suggesting that P_Ca/P_Na of the channel may exceed 1,000:1. Lanthanum, SKF 96365, and 2-APB inhibit both I_ORCa and I_SOC reversibly. Our studies provide the first detailed electrophysiological characterization of voltage-independent Ca²⁺ conductances in C. elegans and form the foundation for ongoing genetic and molecular studies aimed at identifying the genes that encode the intestinal cell channels, for defining mechanisms of channel regulation and for defining their roles in oscillatory Ca²⁺ signaling.

INTRODUCTION

Fluctuating intracellular Ca²⁺ concentration is a ubiquitous signaling mechanism that controls numerous cellular processes, including gene expression, exocytosis and secretion, motility, cell proliferation, programmed cell death, and differentiation (Berridge et al., 2000). Elevation of cytoplasmic Ca²⁺ levels is brought about by Ca²⁺ release from intracellular stores and by influx across the plasma membrane. In excitable cells, Ca²⁺ influx is mediated to a large extent by voltage- and ligand-gated cation channels. Calcium influx into nonexcitable cells, such as blood cells and endothelial and epithelial cells, occurs primarily via second messenger– and store-operated Ca²⁺ channels (SMOCCs and SOCCs)* (Elliott, 2001; Zitt et al., 2002).

The ER is the principal Ca²⁺ store in nonexcitable cells. Agonist binding to plasma membrane tyrosine kinase– or G protein–coupled receptors activates phospholipase C, leading to the production of inositol 1,4,5-trisphosphate (IP₃). IP₃, in turn, activates IP₃ receptors in the ER membrane, inducing Ca²⁺ release that leads to either a sustained elevation of cytoplasmic Ca²⁺ concentration or Ca²⁺ oscillations (Shuttleworth, 1999; Berridge et al., 2000). Sustained Ca²⁺ elevation is often observed with high agonist concentrations and occurs in a biphasic manner. The first phase involves ER Ca²⁺ release. As the stores are depleted of Ca²⁺, SOCCs are activated, allowing Ca²⁺ to enter from the extracellular medium (Parekh and Penner, 1997; Taylor and Thorn, 2001). Cytoplasmic Ca²⁺ levels and Ca²⁺ influx remain elevated as long as the stimulus is maintained (Putney and McKay, 1999; Shuttleworth, 1999).

Lower concentrations of agonists typically trigger Ca²⁺ oscillations (Shuttleworth, 1999). The role of plasma membrane Ca²⁺ entry in generating and maintaining the oscillations is unclear (Shuttleworth, 1999). Calcium oscillations in some cell types continue for long periods in the absence of extracellular Ca²⁺ (Lechleiter and Clapham, 1992). In contrast, oscillatory Ca²⁺ signals in other cell types are strictly dependent on Ca²⁺ influx (Torihashi et al., 2002; Wu et al., 2002).

The molecular identity of both SOCCs and SMOCCs, the mechanisms by which they are regulated, and their precise functional roles in local and global Ca²⁺ signaling are unclear. Genetic model organisms provide a number of powerful experimental advantages for defining the genes and genetic pathways involved in biological processes such as Ca²⁺ signaling. The nematode Caenorhabditis elegans is a particularly attractive model system for such studies (Barr, 2003; Strange, 2003). C. elegans 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.

C. elegans exhibits a number of relatively simple stereotyped behaviors that have formed the bases for powerful forward genetic screens. The defecation cycle is one such behavior. Defecation is an ultradian rhythm 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). Loss-of-function mutations in the IP₃ receptor gene itr-1 slow or eliminate the defecation cycle, whereas overexpression of the gene increases the rate of defecation (Dal Santo et al., 1999). Oscillatory changes in intestinal epithelial cell Ca²⁺ levels track the defecation cycle, with Ca²⁺ levels peaking just before the initiation of posterior body wall muscle contraction. Calcium oscillations are slowed or absent in animals with loss-of-function mutations in itr-1 (Dal Santo et al., 1999). Dal Santo et al. (1999) have suggested that IP₃-dependent Ca²⁺ signals may control the secretion of a factor from the intestinal epithelium that regulates contraction of surrounding body wall muscles.

The ability to combine physiological tools, such as patch clamp analysis and Ca²⁺ imaging, with behavioral assays and forward and reverse genetic screening would provide a powerful approach for defining the molecular details of intestinal cell IP₃-dependent Ca²⁺ signaling. However, electrophysiological characterization of somatic cells in C. elegans is difficult due to the small size of the animal and the presence of a tough, pressurized cuticle that limits access. To circumvent this problem, we recently developed methods that allow the primary culture and terminal differentiation of nematode embryo cells (Christensen et al., 2002).

We report here the electrophysiological characterization of cultured C. elegans intestinal epithelial cells. Intestinal cells express two highly Ca²⁺-selective whole-cell cation conductances. One conductance is insensitive to store depletion, shows strong outward rectification, and is inhibited by intracellular Mg²⁺. Intracellular Ca²⁺ store depletion activates an inwardly rectifying SOCC current. These studies provide the first detailed electrophysiological characterization of voltage-independent Ca²⁺-selective cation conductances in C. elegans and form the foundation for ongoing genetic and molecular studies aimed at identifying the genes that encode the channels, for defining mechanisms of channel regulation and for defining their roles in oscillatory Ca²⁺ signaling.

MATERIALS AND METHODS

C. elegans Strains

All strains were derived from the wild-type N2 line and maintained at 20–25°C using standard methods (Brenner, 1974). The elt-2–GFP strains used in these studies were JR1838 (wIs84) and JM63 (caIs13). GFP strains contain integrated transgenes.

C. elegans Embryonic Cell Culture

Embryonic cells were prepared by treating synchronized adult nematodes with an alkaline hypochlorite solution (0.5 M NaOH and 1% NaOCl) for 5 min (Lewis and Fleming, 1995). Eggs released by this treatment were pelleted by centrifugation and then washed three times with egg buffer containing 118 mM NaCl, 48 mM KCl, 2 mM CaCl₂, 2 mM MgCl₂, and 25 mM HEPES (pH 7.3, 345 mOsm) (Edgar, 1995). Adult carcasses were separated from washed eggs by density centrifugation in 30% sucrose. The egg layer was removed by pipette and washed one time with egg buffer and then pelleted. Eggshells were removed by resuspending pelleted eggs in egg buffer containing 1–2.5 U/ml of chitinase for 45–90 min at room temperature. After digestion of the eggshell, the suspension was gently pipetted up and down several times to dissociate the cells. Cells were washed two times with L-15 cell culture medium (Life Technologies) containing 10% FBS (Hyclone), 50 U/ml penicillin, and 50 μg/ml streptomycin and adjusted to 345 mOsm with sucrose.

Dissociated embryo cells were filtered through a sterile 5-μm Durapore syringe filter (Millipore) to remove undissociated embryos and newly hatched larvae. Filtered cells were plated on 12-mm-diameter glass coverslips coated with 0.5 mg/ml peanut lectin agglutinin. Cultures were maintained at 24°C in a humidified incubator in L-15 cell culture medium.

Patch Clamp Recordings

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 TE300 inverted microscope. Cells were visualized by fluorescence and video-enhanced differential interference contrast (DIC) microscopy.

Patch electrodes were pulled from soft glass capillary tubes (PG10165-4; World Precision Instruments) that had been silanized with dimethyl-dichloro silane. The standard pipette solution for whole-cell recording from intestinal cells contained (mM) 147 sodium gluconate (NaGluconate), 0.6 CaCl₂, 1 MgCl₂, 10 EGTA or 10 BAPTA, 10 HEPES, 2 Na₂ATP, 0.5 Na₂GTP, pH 7.2 (adjusted with CsOH), 325 mOsm. The standard bath solution contained (mM) 145 NaCl, 1 CaCl₂, 5 MgCl₂, 10 HEPES, 20 Glucose, pH 7.2 (adjusted with NaOH). Osmolality was adjusted to 340–345 mOsm with sucrose. Free Ca²⁺ and free Mg²⁺ levels in the various solutions used were calculated using MaxChelator software WINMAXC v.2.1 (www.stanford.edu/~cpatton/maxc.html).

Whole-cell currents were recorded using an Axopatch 200B (Axon Instruments, Inc.) 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, Inc.). Currents were filtered at 5 kHz and digitized at 20–40 kHz. Electrical connections to the amplifier were made using Ag/AgCl wires and 3-M KCl/agar bridges.

Ion substitution studies were performed by replacement of bath Na⁺ with various test cations. For all ion substitution experiments, changes in liquid junction potentials were measured directly using a free-flowing 3-M KCl electrode. Reversal potentials were corrected for these changes. Relative permeabilities were calculated using the following equations derived from the Goldman-Hodgkin-Katz equation (Lewis, 1979; Hille, 2001):

(1)

(2)

where [X⁺]_o is the extracellular concentration of the monovalent substitute cation; [Na⁺]_o is the extracellular Na⁺ concentration; [Ca²⁺]_o is the extracellular Ca²⁺ concentration; ΔE_rev is the change in reversal potential; and F, R, and T have their usual meanings. In studies of the intestinal cell store-operated current, we defined leak current as the current observed immediately after obtaining whole-cell access. This leak current was subtracted from all subsequent current records obtained in the cell.

Chemicals

Thapsigargin, ionomycin, and BAPTA were purchased from Molecular Probes. IP₃ and 2-APB were purchased from Calbiochem. All other chemicals were obtained from Sigma-Aldrich.

Statistical Analyses

Data are presented as means ± SEM. Statistical significance was determined using a two-tailed t test or ANOVA followed by a Bonferroni multiple comparisons test. P values of <0.05 were taken to indicate statistical significance.

RESULTS

Identification of C. elegans Intestinal Epithelial Cells in Primary Culture

As described previously, isolated C. elegans embryo cells undergo terminal differentiation when cultured in vitro (Christensen et al., 2002). Culturing embryo cells from worm strains expressing cell-specific GFP reporters allows identification of differentiated cell types. ELT-2 is a GATA transcription factor expressed exclusively in intestinal cells and is required for intestinal morphogenesis beginning at the 44–46-cell stage of embryonic development (Fukushige et al., 1999). Primary cell cultures were prepared from worm strains expressing an elt-2–GFP transcriptional fusion construct (worm strains provided by J. Rothman [University of California, Santa Barbara, CA] and J. McGhee [University of Calgary, Calgary, Alberta, Canada]). Fig. 1 shows combined DIC and fluorescence micrographs of a transgenic worm expressing elt-2–GFP and an intestinal epithelial cell cultured from elt-2–GFP-expressing worms.

Figure 1.

Expression of elt-2–GFP reporter in C. elegans intestinal cells. Images are overlays of DIC and fluorescence micrographs of a transgenic worm (left) and cultured intestinal cell (right) expressing elt-2–GFP in the cell nucleus. GFP fluorescence is shown in green. Bars: (left) 10 μm; (right) 2.5 μm. Em, developing embryo in uterus; Oo, oocyte in proximal gonad; IC, intestinal cell; N, intestinal cell nucleus. Arrowheads denote refractile granules that are most likely intracellular storage granules.

Our previous studies have shown that various muscle and neuronal cell types in primary culture are present at a frequency remarkably close to that observed in a newly hatched L1 larva (Christensen et al., 2002). An L1 larva is comprised of 558 cells of which 20, or 3.6%, are intestinal cells. Intestinal cells represented 4.1 ± 0.1% (n = 3) of the total cells in culture. elt-2–GFP expression was concentrated in the nuclei of cultured intestinal cells (Fig. 1, right), similar to that observed in vivo (Fig. 1, left). In addition, the cytoplasm of the cultured cells contained numerous refractile granules (Fig. 1, right, arrowheads) that were also highly autofluorescent (unpublished data). These most likely represent storage granules, which are a prominent characteristic of intestinal cells in the intact worm (Kostich et al., 2000; Fig. 1, left).

Intestinal Cells Express an Outwardly Rectifying Cation Current

Cultured intestinal cells were patch clamped readily in the whole-cell mode. An outwardly rectifying whole-cell current (Fig. 2, A and B) was observed when cells were bathed and dialyzed with control bath (145 mM NaCl) and pipette (147 mM NaGluconate) solutions. Whole-cell current amplitude typically increased approximately two- to threefold and then stabilized within 1–2 min after obtaining the whole-cell configuration (Fig. 2 C).

Figure 2.

Characteristics of outwardly rectifying intestinal cell whole-cell current. (A) Whole-cell currents recorded from a cultured intestinal cell using standard bath (145 mM NaCl) and pipette (147 mM NaGluconate) solutions. Currents were elicited by stepping membrane voltage from −100 to +100 mV in 20-mV steps from a holding potential of 0 mV. Voltage steps were 400 ms long. Inset shows inactivation behavior observed at strongly hyperpolarized voltages. (B) Steady-state I–V relationship for the whole-cell currents shown in A. (C) Whole-cell current activation observed after membrane rupture. Current recordings from three intestinal cells are shown. Values shown are relative to those observed at time 0, which is defined as the time at which whole-cell access was obtained. Whole-cell current typically increased two- to threefold within 1–3 min after membrane rupture and then stabilized.

The whole-cell current was voltage and time dependent (Fig. 2, A and B). Strong depolarization and hyperpolarization activated and inactivated the current, respectively. At +100 mV, current activation was well fit by a double exponential describing mean ± SEM fast (τ_f) and slow (τ_s) time constants of 19 ± 2 and 154 ± 29 ms (n = 43), respectively. Current inactivation was also well fit by a double exponential. Mean ± SEM τ_f and τ_s at −100 mV were 14 ± 2 and 195 ± 36 ms (n = 43), respectively.

The ionic nature of the outwardly rectifying whole-cell current was determined by ion substitution studies. Replacement of 147 mM Na⁺ in the pipette solution with NMDG⁺ dramatically inhibited outward current (Fig. 3 A) and increased E_rev significantly (P < 0.005) from a mean ± SEM value of 21 ± 3 (n = 5) to 37 ± 3 mV (n = 5).1 The shift in E_rev and reduction in outward current are consistent with a cation current.

Figure 3.

Ionic dependence of outwardly rectifying whole-cell current. (A) Replacement of 147 mM Na⁺ (Na_i, □) in the pipette solution with NMDG⁺ (▪) increased reversal potential (E_rev) by +16 mV (P < 0.005) and dramatically reduced outward current.1 Values are means ± SEM (n = 5). (B) Reduction of bath Cl⁻ concentration (Cl_o) from 157 (□) to 10 mM (▪) (gluconate substitution) caused a small increase in outward current and decrease in E_rev of −3 ± 2 mV. The changes in current amplitude and E_rev were not statistically significant (P > 0.1). Values are means ± SEM (n = 3). Changes in E_rev and whole-cell current amplitude indicate that the current is carried primarily by cations. Voltage clamp protocol was the same as described in Fig. 2.

Reduction of bath Cl⁻ from 157 to 10 mM (gluconate replacement2) induced a small increase in outward current and shift in mean ± SEM E_rev from 29 ± 3 to 24 ± 1 mV (n = 3; Fig. 3 B). The small changes in current amplitude and E_rev induced by reduction of bath Cl⁻ were not statistically significant (P > 0.1) and are opposite to those expected for anion-selective channels. Taken together, the data in Fig. 3 demonstrate that the whole-cell current is carried predominantly by cations.

The Outwardly Rectifying Cation Conductance Has a High Selectivity for Ca²⁺ over Monovalent Cations

The relative permeability to various cations of the channel responsible for the outwardly rectifying whole-cell current was determined using the Goldman-Hodgkin-Katz equation after complete substitution of bath Na⁺ with various test cations. Removal of bath Ca²⁺ and Mg²⁺ and addition of 1 mM EGTA (nominally divalent-free medium) caused an immediate and substantial increase in whole-cell current (Fig. 4) . The mean ± SEM increase observed at +80 mV was 286 ± 58 pA (n = 27; P < 0.0001). Removal of Ca²⁺ and Mg²⁺ also caused a significant (P < 0.0001) decrease in E_rev, from 24 ± 1 (n = 48) to 11 ± 0.5 mV (n = 27), and altered current voltage and time dependence (Fig. 4 A). At potentials of +80 mV and above, currents observed in nominally divalent-free medium typically showed partial inactivation (Fig. 4 A). Current inactivation was fit by a single exponential describing a mean ± SEM time constant (τ) at +100 mV of 194 ± 42 ms (n = 17).

Figure 4.

Effect of extracellular Ca²⁺ and Mg²⁺ removal on amplitude of the outwardly rectifying cation current. (A) Whole-cell currents observed under control (1 mM Ca²⁺ and 5 mM Mg²⁺) and nominally divalent-free (0 mM Ca²⁺, 0 mM Mg²⁺, and 1 mM EGTA) conditions. Voltage clamp protocol was the same as described in Fig. 2. (B) Steady-state I–V relationships of outwardly rectifying currents observed under control conditions (•), in nominally divalent-free medium (buffered with 1 mM EGTA; ○) and in divalent-free medium (buffered with 1 mM EDTA; □). Values are means ± SEM (n = 6–33).

Micromolar concentrations of extracellular Mg²⁺ block channels such as the NMDA receptor (Mayer et al., 1984; Nowak et al., 1984) and the recently described Mg²⁺-inhibited cation (MIC) channel (Hermosura et al., 2002; Kozak et al., 2002). To determine whether a similar block occurs in the channel responsible for the outwardly rectifying cation current, we exposed cells to divalent-free medium containing 1 mM EDTA in order to fully chelate extracellular Mg²⁺. As shown in Fig. 4 B, the current-to-voltage relationship of the outwardly rectifying current was largely unaffected by this maneuver.

Replacement of bath Na⁺ with K⁺, Cs⁺, or NMDG⁺ shifted E_rev to more negative values (Table I) . The calculated relative permeabilities (i.e., P_cation/P_Na; Table I) determined from the changes in E_rev yielded a monovalent cation selectivity sequence of Na⁺ > K⁺ > Cs⁺ ≫ NMDG⁺.

TABLE I.

Relative Cation Permeability of the Outwardly Rectifying Conductance

Cation	ΔErev	Pcation/PNa
	mV
K+	−10 ± 1	0.67 ± 0.02 (7)
Cs+	−34 ± 2	0.27 ± 0.02 (7)
NMDG+	−90 ± 5	0.03 ± 0.01 (7)
Ca2+	29 ± 1	64 ± 2 (6)a

Whole-cell currents were elicited by stepping membrane voltage from −100 to +100 mV in 20-mV steps from a holding potential of 0 mV. Voltage steps were 400 ms long. Steady-state current-to-voltage relationships were plotted for determination of E_rev in the presence of Na⁺ and various test cations. Relative permeabilities were calculated using equations derived from the Goldman-Hodgkin-Katz equation (see materials and methods). Values are means ± SEM (number of cells). All changes in E_rev are statistically significant (P < 0.0001).

^aSee footnote 3.

The increase in whole-cell current upon removal of extracellular divalent cations (Fig. 4) suggested strongly that the channel is permeable to Ca²⁺ and/or Mg²⁺ and that permeation by divalents blocks monovalent cation flux. To examine this possibility directly, we measured whole-cell currents in the presence of extracellular solutions containing varying proportions of Ca²⁺ and Na⁺. Addition of 1 mM Ca²⁺ to a solution containing 74 mM Na⁺ caused a dramatic reduction in whole-cell current (Fig. 5 A). As the proportion of Ca²⁺ was elevated, the current passed through a minimum and then increased (Fig. 5 A). This anomalous mole-fraction behavior is characteristic of multi-ion channels where two or more ions simultaneously occupy and move through the channel pore (Hille, 2001). Both the number and type of ions in the pore determine the overall permeability properties of multi-ion channels (Hille, 2001). Many highly Ca²⁺-selective cation channels exhibit anomalous mole-fraction behavior (Hoth, 1995; Vennekens et al., 2001). High affinity Ca²⁺ binding in the channel pore appears to be responsible for their high selectivity for Ca²⁺ versus monovalent cations (Almers and McCleskey, 1984; Lepple-Wienhues and Cahalan, 1996).

Figure 5.

Calcium selectivity of the outwardly rectifying cation conductance. (A) Relationship between relative whole-cell current amplitude and fractional concentration of extracellular Ca²⁺ ([Ca²⁺]_o/([Na⁺]_o + [Ca²⁺]_o). As the proportion of Ca²⁺ is elevated, the current passes through a minimum and then increases. This anomalous mole-fraction behavior is characteristic of highly Ca²⁺-selective cation channels (Hoth, 1995; Vennekens et al., 2001). All solutions were nominally Mg²⁺ free. The Ca²⁺-free solution was buffered with 1 mM EGTA. Values are means ± SEM (n = 6–11). (B) I–V relationships of whole-cell current in the presence of 150 mM Na⁺ in the bath (□) and when Na⁺ was replaced with 130 mM NMDG⁺ and 10 mM Ca²⁺ (▪). Elevation of extracellular Ca²⁺ increased E_rev by 29 ± 1 mV (P < 0.0001). Calculated relative Ca²⁺ permeability (P_Ca/P_Na) is 64:1 (Table I). Voltage clamp protocol was the same as described in Fig. 2.

Given the results in Fig. 4 and Fig. 5 A, we measured relative Ca²⁺ permeability by replacing bath Na⁺ with 130 mM NMDG⁺ and 10 mM Ca²⁺. Elevation of bath Ca²⁺ increased E_rev significantly (P < 0.0001), by 29 ± 1 mV (n = 6) (Fig. 5 B; Table I). The calculated relative permeability of Ca²⁺ to Na⁺ is 64:1 (Table I).3 Together, the data in Figs. 4 and 5 demonstrate that a highly Ca²⁺-selective cation channel carries the outwardly rectifying whole-cell current. We hereafter refer to the outwardly rectifying Ca²⁺ current as I_ORCa.

Pharmacological Characteristics of I_ORCa

Lanthanum is a trivalent cation that inhibits voltage-gated and store-operated Ca²⁺ channels as well as some members of the transient receptor potential (TRP) cation channel superfamily (Aussel et al., 1996; Halaszovich et al., 2000; Beedle et al., 2002). Exposure of cultured intestinal cells to 100 μM La³⁺ in the bath solution virtually abolished the outwardly rectifying current (P < 0.05; Fig. 6 A). Current inhibition by 100 μM La³⁺ was fully reversible (Fig. 6 A). The concentration of La³⁺ required for 50% inhibition of I_ORCa is 3.4 μM (Fig. 6 B).

Figure 6.

Pharmacology of the outwardly rectifying cation conductance. (A) Addition of 100 μM La³⁺, 100 μM SKF 96365, or 100 μM 2-APB (black bar) to the bath inhibited whole-cell current by 50–90%. *, P < 0.05; **, P < 0.01 (compared with control, white bar). Inhibitory effects of La³⁺, SKF 96365, and 2-APB were fully reversible. Current levels observed after drug washout (gray bar) were not significantly (P > 0.05) different from those observed before drug addition (white bar). Values are means ± SEM (n = 4–5). (B) Dose–response relationship for the inhibitory effect of La³⁺. Data were fit using the equation I = 1/1 + ([Mg²⁺]/K_1/2)ⁿ. K_1/2 and n are 3.4 μM and 1.1, respectively. Values are means ± SEM (n = 5–9). Voltage clamp protocol was the same as described in Fig. 2.

SKF 96365 inhibits receptor-activated and voltage-gated calcium influx (Merritt et al., 1990) as well as the Ca²⁺ release–activated channel (CRAC) (Kozak et al., 2002; Prakriya and Lewis, 2002) and TRP channel activity (Halaszovich et al., 2000). 100 μM SKF 96365 inhibited I_ORCa ∼60% (P < 0.01) in a reversible manner (Fig. 6 A). 2-APB inhibits IP₃ receptor Ca²⁺ channel activity (Bilmen and Michelangeli, 2002) and plasma membrane Ca²⁺ entry channels, including CRAC and TRP channels (Prakriya and Lewis, 2001; Bootman et al., 2002; Hermosura et al., 2002; Prakriya and Lewis, 2002). 100 μM 2-APB reversibly inhibited I_ORCa by ∼50% (P < 0.01; Fig. 6 A). Lanthanum, SKF 96365, and 2-APB all inhibited I_ORCa rapidly; current inhibition was maximal within 30–60 s after adding the drugs to the bath (unpublished data). Washout of the drugs and recovery of I_ORCa occurred with a similar time course (unpublished data).

Depletion of Intracellular Ca²⁺ Stores Does Not Regulate I_ORCa

As shown in Fig. 2 C, I_ORCa activated two- to threefold during the first 1–2 min after obtaining whole-cell access. Two observations suggested that current activation might be due to depletion of intracellular Ca²⁺ stores. First, the biophysical characteristics of the current resemble those of members of the TRP cation channel superfamily. Evidence suggests that some TRPs may be regulated by store depletion (Clapham et al., 2001; Elliott, 2001; Montell, 2001; Zitt et al., 2002). Second, the pipette solutions used in the studies shown in Fig. 2 C contained 10 mM EGTA and low Ca²⁺, which could lead to passive store depletion (Hoth and Penner, 1993).

We performed three experiments to directly examine the role of store depletion in regulation of I_ORCa. First, cells were patch clamped with a Ca²⁺-free pipette solution to passively deplete stores or a solution containing 175 nM free Ca²⁺. Calcium levels >90 nM are expected to maintain store Ca²⁺ levels (Hermosura et al., 2002; Kozak et al., 2002). As shown in Fig. 7 A, there was no significant (P > 0.2) difference in either the rate of current activation or the peak current amplitude observed in these two experiments.

Figure 7.

The outwardly rectifying cation current is not regulated by depletion of Ca²⁺ stores. (A) Rates of whole-cell current activation (black bar) and peak current amplitudes (white bar) in cells dialyzed with a Ca²⁺-free pipette solution to passively deplete intracellular Ca²⁺ stores and a pipette solution containing 175 nM free Ca²⁺. Removal of intracellular Ca²⁺ had no significant (P > 0.2) effect on the rate of current activation or the peak current amplitude. Intracellular Ca²⁺ was buffered with 1 mM EGTA. Currents were measured using the standard bath medium containing both Ca²⁺ and Mg²⁺. Whole-cell currents were elicited by ramping membrane potential from −80 to +80 mV at 160 mV/s every 5 s. (B) Rates of whole-cell current activation (black bar) and peak current amplitudes (white bar) in control cells and cells treated with 10 μM IP₃ and 1 μM thapsigargin. Depletion of intracellular stores with IP₃ and thapsigargin had no significant (P > 0.1) effect on the rate of current activation or the peak current amplitude. Intracellular Ca²⁺ was buffered to 11 nM with 10 mM EGTA. Cells were exposed to nominally divalent-free bathing medium immediately after obtaining whole-cell access. Voltage clamp protocol was the same as described in Fig. 2. (C) Effect of depletion of Ca²⁺ stores in intact cells on rate of current activation (black bar) and initial and peak current amplitude (white bar). Experimental cells were preincubated for 9–17 min in nominally divalent-free bathing medium containing 1 μM thapsigargin and then patch clamped with a pipette solution containing 10 μM IP₃. Control cells were exposed to nominally divalent-free bathing medium immediately after obtaining whole-cell access and were not treated with IP₃ or thapsigargin. Store depletion had no significant (P > 0.1) effect on initial whole-cell current or subsequent current activation. Intracellular Ca²⁺ was buffered to 11 nM with 10 mM EGTA. Values are means ± SEM (n = 3–6). Voltage clamp protocol was the same as described in A. Initial rates of current activation were quantified by performing linear regression analysis on whole-cell currents measured during the first 60–180 s after obtaining whole-cell access.

Second, we patch clamped cells with a pipette solution containing 10 μM IP₃. Immediately after obtaining whole-cell access, IP₃-treated cells were exposed to a nominally divalent-free bath solution containing 1 μM thapsigargin to inhibit store Ca²⁺ uptake. Neither the rate of current activation nor the peak current amplitude were significantly (P > 0.1) altered by IP₃ and thapsigargin.

Finally, we attempted to activate I_ORCa in intact cells by depleting intracellular Ca²⁺ stores before patch clamping. Cells were incubated in nominally divalent-free bath solution containing 1 μM thapsigargin for 9–17 min and then were patch clamped with a pipette solution containing 10 μM IP₃. Control cells were patch clamped with a standard pipette solution without IP₃ and were exposed to nominally divalent-free bath immediately after obtaining whole-cell access. If store depletion activates I_ORCa, initial current levels should be higher in cells pretreated with Ca²⁺- and Mg²⁺-free bath containing thapsigargin. However, as shown in Fig. 7 C, initial whole-cell current amplitudes were not significantly (P > 0.1) different in control and experimental cells. The outward currents in both groups of cells showed gradual activation, but the rates of current activation and peak current amplitudes were not significantly (P > 0.5) different (Fig. 7 C). We conclude from the data shown in Fig. 7 that I_ORCa is not activated by depletion of intracellular Ca²⁺ stores.

I_ORCa Is Inhibited by Intracellular Mg²⁺

We examined the effect of intracellular Mg²⁺ concentration on the activity of I_ORCa. Cells were patch clamped with an ATP- and GTP-free pipette solution in which EGTA was replaced by 10 mM BAPTA. The concentration of MgCl₂ added to the pipette solution varied between 0 and 6 mM. As shown in Fig. 8 (A and B), increasing free Mg²⁺ concentrations inhibited I_ORCa in a saturable manner. The estimated K_1/2 value for free Mg²⁺ is 692 μM (Fig. 8 B).

Figure 8.

Inhibition of the outwardly rectifying cation current by intracellular Mg²⁺. (A) I–V relationships for cells patch clamped with ATP- and GTP-free pipette solutions containing 0–6 mM MgCl₂. Chloride concentrations in the solutions were maintained constant by addition of 0–12 mM NMDGCl. EGTA was replaced with 10 mM BAPTA to buffer intracellular Ca²⁺ at 14 nM. Note that the Mg²⁺ concentrations indicated on the figure are those added to the pipette solution. Therefore, 0 Mg²⁺ should be considered nominally Mg²⁺ free. (B) Dose–response relationship for inhibition of the outwardly rectifying current by intracellular free Mg²⁺. Data were fit using the equation I = 1/1 + ([Mg²⁺]/K_1/2)ⁿ. K_1/2 and n are 692 μM and 0.8, respectively. Open circle is inhibition observed when free Mg²⁺ concentration is elevated by addition of 6 mM MgATP. (C) Effect of Mg²⁺ nucleotides on the outwardly rectifying cation current. Whole-cell current is inhibited ∼60% by 6 mM MgATP. Calculated concentration of free Mg²⁺ in the pipette solution containing 6 mM MgATP is 700 μM. The degree of inhibition is similar to that observed when free Mg²⁺ is elevated by addition of MgCl₂ (see B). *, P < 0.05 (compared with 6 mM TrisATP). Values are means ± SEM (n = 5–9). Currents were measured in standard bath medium containing both Ca²⁺ and Mg²⁺ 3–4 min after obtaining whole-cell access when activation was complete. Voltage clamp protocol was the same as described in Fig. 2.

To determine if I_ORCa is also inhibited by Mg-nucleotides, 6 mM MgATP or 6 mM TrisATP was added to the pipette solution. As shown in Fig. 8 C, 6 mM MgATP inhibited whole-cell current ∼60% at +80 mV (P < 0.05). The calculated free Mg²⁺ concentration in a solution containing 6 mM MgATP is 700 μM. When plotted as a function of calculated free Mg²⁺ concentration (Fig. 8 B, open circle), the degree of inhibition observed with 6 mM MgATP is very similar to that observed with Mg²⁺ alone. These results indicate that I_ORCa is inhibited by free Mg²⁺ but is insensitive to MgATP.

Intracellular Ca²⁺ Store Depletion Activates an Inwardly Rectifying Current

SOCCs play important roles in IP₃-dependent intracellular Ca²⁺ signaling pathways (Putney and McKay, 1999; Lewis, 2001; Taylor and Thorn, 2001). Given the dependence of the C. elegans defecation cycle on IP₃ and oscillatory Ca²⁺ signaling in the intestine (Dal Santo et al., 1999), we performed patch clamp studies to determine if intestinal cells expressed store-operated channels.

I_ORCa, which dominates whole-cell recordings, was inhibited by inclusion of 5 mM free Mg²⁺ in the pipette solution (Kozak et al., 2002; Prakriya and Lewis, 2002). To prevent Ca²⁺ store depletion, we patch clamped cells with a pipette solution containing ATP, GTP, and 200 nM free Ca²⁺ buffered with 10 mM BAPTA. The bath solution contained 145 mM Na⁺ and 20 mM Ca²⁺. Using these solutions, we observed that whole-cell current remained stable for at least 5–7 min after obtaining whole-cell access in three out of three cells (Fig. 9 C). The mean ± SEM current observed just before loss of the whole-cell seal was −5.5 ± 2.7 pA/pF at −120 mV (n = 3).

Figure 9.

Activation of an inwardly rectifying store-operated current by depletion of intracellular Ca²⁺ stores. (A) An inwardly rectifying current activates when store depletion is induced by addition of 10 μM IP₃ to a pipette solution containing 18 nM free Ca²⁺. Currents were elicited by ramping membrane voltage from −120 to +80 mV at 200 mV/s every 5 s. Leak current was subtracted from the traces shown. (B) Inwardly rectifying whole-cell currents elicited by stepping membrane voltage from −120 to +80 mV from a holding potential of 0 mV. Steps were 400 ms in duration. Currents were measured after activation induced by IP₃ was complete. (C) Changes in whole-cell current observed in the absence of store depletion (○), during passive store depletion (•), and during active store depletion induced by IP₃ (▪). Leak current was subtracted from the currents induced by active and passive store depletion. All experiments shown were performed in the presence of 5 mM free Mg²⁺ in the pipette solution to inhibit I_ORCa and 145 mM Na⁺/20 mM Ca²⁺ in the bath.

Active depletion of Ca²⁺ stores was induced by dialyzing cells with a nucleotide-free pipette solution containing 18 nM free Ca²⁺ and 10 μM IP₃. A strongly inwardly rectifying, voltage-independent cation current was activated under these conditions (Fig. 9, A–C). The mean time to the start of current activation after membrane rupture was 84 s, and the current activated to a peak value of −31 pA/pF at −120 mV at an initial rate of −26 pA/pF/min (Table II) .

TABLE II.

Store Depletion–induced Activation Characteristics of the Inwardly Rectifying Current

Store depletion protocol	Time to start of current activationa	Rate of current activationb	Time to peak current	Peak current
	s	pA/pF/min	s	pA/pF
Passive	163 ± 26 (8)	−3.3 ± 0.5 (8)	462 ± 52 (6)	−16 ± 3 (6)
Active
10 μm IP3	84 ± 12c (81)	−26 ± 3e (81)	230 ± 17d (77)	−31 ± 2d (77)
Active
2 μm ionomycin		−72 ± 17e (8)		−38 ± 14 (8)

Currents were elicited by ramping membrane voltage from −120 to +80 mV at 200 mV/s every 5 s. Peak current and rate of current activation were measured at −120 mV.

^aTime to start of current activation is the time after obtaining whole-cell access that current activation began.

^bRate of current activation was quantified by performing linear regression analysis on whole-cell currents measured during the first 60–120 s after initiation of current activation. The time course of current activation in many cells was sigmoidal in nature (e.g., Fig. 9 C), displaying an initial very slowly rising phase followed by a much steeper and rapid linear increase in current amplitude. To avoid ambiguity in characterizing current activation, we measured the rate of activation during the steepest initial part of this sigmoidal curve. The first point to fall within this region was defined as the start of current activation. Values are means ± SEM (number of cells).

^cP < 0.05, compared with passive store depletion.

^dP < 0.01, compared with passive store depletion.

^eP < 0.001, compared with passive store depletion.

It is conceivable that the inwardly rectifying current is activated by IP₃ rather than store depletion per se. To test for this possibility, we passively depleted intracellular Ca²⁺ stores by dialyzing cells with a pipette solution containing 18 nM free Ca²⁺ alone. Passive store depletion also activated an inwardly rectifying current (Fig. 9 C), albeit at a rate ∼13% of that observed when stores were depleted actively. The mean time to the start of current activation after membrane rupture was 163 s (Table II). This delay in current activation was significantly (P < 0.05) longer than that observed in the presence of IP₃ (Table II). Mean initial rate of current activation and peak current at −120 mV were −3.3 pA/pF/min and −16 pA/pF, respectively (Table II). The rate of current activation and peak current were significantly (P < 0.01) decreased compared with that observed with active store depletion induced by IP₃ (Table II). These results are consistent with store depletion being the mechanism responsible for activation of the inwardly rectifying current.

As a final test for the involvement of store depletion in current activation, we dialyzed cells with a pipette solution containing 18 nM free Ca²⁺ and then exposed them 30–150 s after obtaining whole-cell access to 2 μM ionomycin for 30–60 s. Ionomycin is a Ca²⁺ ionophore that is expected to induce efflux of Ca²⁺ from intracellular stores and activation of store-dependent cation channels (Hoth and Penner, 1993; Parekh, 1998; Voets et al., 2001). In the presence of ionomycin, the inwardly rectifying current activated rapidly. The initial rate of current activation and peak current in the presence of ionomycin were −72 pA/pF/min and −38 pA/pF, respectively, at −120 mV. The rate of current activation was ∼22-fold faster (P < 0.001) than that observed with passive store depletion (Table II). Taken together, the results shown in Fig. 9 and Table II demonstrate clearly that depletion of IP₃-dependent intracellular Ca²⁺ stores activates an inwardly rectifying store-operated current. Currents activated by the three store depletion protocols had the same I–V relationships (unpublished data), indicating that they were likely carried by the same channel.

Selectivity of the Store-operated Channel

In the presence of 145 mM Na⁺ and 20 mM Ca²⁺ in the bath, the store-operated current showed very strong inward rectification (Fig. 9, A and B) and voltage-independent gating (Fig. 9 B). For the majority of cells examined, the slope of the I–V plot at positive voltages was extremely shallow, and a clear reversal of current direction was not detectable (Fig. 9 A).

Replacement of bath Na⁺ and Ca²⁺ with NMDG⁺ completely blocked inward current (Fig. 10 A), demonstrating that the channel is highly cation selective. In the presence of 130 mM NMDG⁺ and 10 mM Ca²⁺, a strongly inwardly rectifying current was observed (Fig. 10 A). Mean ± SEM current density at −120 mV was −31 ± 5 pA/pF (n = 4), which is similar to that observed in the presence of Na⁺ and Ca²⁺ (Table II). These results demonstrate that the store-operated channel has a high selectivity for Ca²⁺ or Na⁺. The inability to accurately measure E_rev at positive voltages precludes accurate calculation of relative Ca²⁺ to Na⁺ permeability. However, if E_rev is >+80 mV, the estimated P_Ca/P_Na is >1,000:1. Based on these results, we hereafter refer to the inwardly rectifying store-operated Ca²⁺ channel as SOCC and the channel current as I_SOC.

Figure 10.

Cation selectivity of the inwardly rectifying current. (A) Whole-cell currents observed in the presence of various extracellular cations. Exposure of cells to divalent-free (buffered with 1 mM EDTA) 150 mM Na⁺ bath increases inward and outward monovalent current (□; compare with Fig. 9 A). Replacement of bath Na⁺ with 150 mM NMDG⁺ (•) eliminates inward current. In the presence of 10 mM Ca²⁺ and 130 mM NMDG⁺ (○), inward current is restored. (B) Relative organic cation currents (i.e., I_cation/I_Na). Whole-cell currents were measured at −120 mV in divalent-free (buffered with 1 mM EDTA) 150 mM Na⁺-containing bath and after complete replacement of Na⁺ with various organic cations. **, P < 0.01 (compared with current observed with Na⁺). (C) Relative divalent cation currents (i.e., I_divalent/I_Na). Whole-cell currents were measured at −120 mV in a divalent-free (buffered with 1 mM EDTA) 150 mM Na⁺-containing bath and in a bath containing 130 mM NMDG⁺ and 10 mM Ca²⁺, Ba²⁺, or Sr²⁺. **, P < 0.01; ***, P < 0.001 (compared with current observed with Ca²⁺). Values are means ± SEM (n = 4–13). Whole-cell currents were elicited by stepping membrane voltage from −120 to +80 mV from a holding potential of 0 mV. Steps were 400 ms in duration. Whole-cell current was activated by inclusion of 10 μM IP₃ in the patch pipette solution. Leak currents were subtracted from all current records. Experiments were performed in the presence of 5 mM free Mg²⁺ in the pipette solution to inhibit I_ORCa.

In the presence of divalent-free (buffered with 1 mM EDTA) 150 mM Na⁺-containing bath, significant inward and outward current was detected (Fig. 10 A). Mean ± SEM E_rev of the Na⁺ current was 13 ± 1 mV (n = 28). Replacement of bath Na⁺ with Cs⁺ or Rb⁺ shifted E_rev to more negative values (Table III) . Substitution of Na⁺ with Li⁺ had no significant (P > 0.2) effect on E_rev (Table III). The calculated relative cation permeabilities (i.e., P_cation/P_Na; Table III) determined from the changes in E_rev yielded a monovalent inorganic cation selectivity sequence of Na⁺ ≈ Li⁺ > Rb⁺ ≈ Cs⁺.

TABLE III.

Relative Inorganic Monovalent Cation Permeability of the Inwardly Rectifying Conductance

Cation	ΔErev	Pcation/PNa
	mV
Li+	1.4 ± 0.9	1.06 ± 0.04 (5)
Cs+	−16 ± 5a	0.60 ± 0.10 (7)
Rb+	−16 ± 4b	0.57 ± 0.08 (6)

Whole-cell currents were elicited by stepping membrane voltage from −120 to +80 mV in 20-mV steps from a holding potential of 0 mV. Voltage steps were 400 ms long. Steady-state current-to-voltage relationships were plotted for determination of E_rev in the presence of Na⁺ and various test cations. Relative permeabilities were calculated using equations derived from the Goldman-Hodgkin-Katz equation (see materials and methods). Values are means ± SEM (number of cells).

^aP < 0.03.

^bP < 0.02.

We attempted to measure relative organic cation permeability of SOCC by replacing bath Na⁺ with TMA⁺, TRIS⁺, or NMDG⁺. Reversal potentials could not be accurately estimated for NMDG⁺ and TRIS⁺. We therefore quantified organic cation current relative to Na⁺ current (i.e., I_cation/I_Na) at −120 mV. TMA⁺ and TRIS⁺ currents were ∼80 and 5%, respectively, of those observed with Na⁺; NMDG⁺ was effectively impermeant (Fig. 10, A and B). The diameters of TMA⁺, TRIS⁺, and NMDG⁺ are 0.55 nm, 0.64 nm, and 0.68 nm, respectively (Hille, 2001). This suggests that the pore diameter of SOCC is ∼0.6–0.7 nm.

To determine whether the observed organic solute permeabilities are reflective of SOCC rather than nonspecific leak, we measured TMA⁺ and TRIS⁺ currents relative to Na⁺ in the absence of store depletion and I_SOC activation. Mean ± SEM I_cation/I_Na for TMA⁺ and TRIS⁺ were 1.0 ± 0.2 (n = 5) and 1.6 ± 0.9 (n = 6), respectively. These values were not significantly different (P > 0.6), indicating that the leak does not discriminate between the two cations. In contrast, SOCC discriminates strongly between TMA⁺ and TRIS⁺ (Fig. 10 B).

We also attempted to measure relative divalent cation permeability of the channel. Cells were bathed initially with a divalent-free solution containing 150 mM Na⁺. Sodium was then replaced by a solution containing 130 mM NMDG⁺ and 10 mM Ca²⁺, Ba²⁺, or Sr²⁺. Relative divalent cation currents (i.e., I_divalent/I_Na) at −120 mV are shown in Fig. 10 C. Both Ba²⁺ and Sr²⁺ were significantly (P < 0.01) less permeable than Ca²⁺. The divalent cation selectivity sequence of SOCC is Ca²⁺ > Ba²⁺ ≈ Sr²⁺.

Pharmacological Characteristics of I_SOC

We compared the pharmacological properties of I_SOC to I_ORCa. Exposure of intestinal cells to 100 μM La³⁺ inhibited I_SOC ∼90% (P < 0.01; Fig. 11, A and B) . This inhibitory effect was partially reversible (Fig. 11 A). The concentration of La³⁺ required to inhibit I_SOC 50% was 9 μM (Fig. 11 B), a value twofold higher than that observed for I_ORCa (Fig. 6 A).

Figure 11.

Pharmacology of the inwardly rectifying current. (A) Addition of 100 μM La³⁺, 100 μM SKF 96365, or 100 μM 2-APB (black bar) to the bath inhibited whole-cell current by 65–90%. *, P < 0.05; **, P < 0.01 (compared with control, white bar). 5 μM 2-APB had no effect on whole-cell current (black bar). Inhibitory effects of SKF 96365 and 2-APB were fully reversible (gray bar). Current levels observed after washout of SKF 96365 and 100 μM 2-APB (gray bar) were not significantly (P > 0.05) different from those observed before drug addition (white bar). Washout of 5 μM 2-APB caused a small but significant (*, P < 0.05) stimulation of whole-cell current. Values are means ± SEM (n = 4–6). (B) Dose–response relationship for the inhibitory effect of La³⁺. Data were fit using the equation I = 1/1 + ([Mg²⁺]/K_1/2)ⁿ. K_1/2 and n are 9 μM and 1.2, respectively. Values are means ± SEM (n = 4–8). Voltage clamp protocol was the same as described in Fig. 9 B. Whole-cell current was activated by inclusion of 10 μM IP₃ in the patch pipette solution. Leak currents were subtracted from all current records. Experiments were performed in the presence of 5 mM free Mg²⁺ in the pipette solution to inhibit I_ORCa and 145 mM Na⁺/20 mM Ca²⁺ in the bath.

100 μM SKF 96365 inhibited I_SOC ∼65% (P < 0.01) in a reversible manner (Fig. 11 A). Concentrations of 2-APB >10 μM irreversibly inhibit CRAC (Prakriya and Lewis, 2001, 2002; Voets et al., 2001; Hermosura et al., 2002). However, at lower concentrations (5 μM), the drug stimulates CRAC activity (Prakriya and Lewis, 2001, 2002). The intestinal cell SOCC has some characteristics that resemble those of CRAC (see discussion). We therefore tested the effects of low and high concentrations of 2-APB on the current. Exposure to 5 μM 2-APB had no significant (P > 0.05) effect on I_SOC (Fig. 11 A). Washout of the drug induced a small, but statistically significant (P < 0.05), increase in current. Exposure to 100 μM 2-APB inhibited I_SOC by ∼90% (Fig. 11 A). Inhibition was completely reversed by drug washout (Fig. 11 A).

Inactivation of I_SOC

I_SOC activity was relatively stable in the presence of a bath solution containing 145 mM Na⁺ and 20 mM Ca²⁺ (Fig. 12 A). Mean ± SEM relative current observed 3 min after store depletion–induced activation was complete was 0.93 ± 0.02 (n = 18). In contrast, the SOCC-mediated Na⁺ current observed in divalent-free bath inactivated immediately after Ca²⁺ removal (Fig. 12 B). The mean ± SEM rate of Na⁺ current inactivation was −12 ± 2 pA/pF/min or −13 ± 2%/min (n = 9).

Figure 12.

Inactivation of the inwardly rectifying current. (A) Example of active store depletion–induced activation of whole cell in a bath solution containing 145 mM Na⁺ and 20 mM Ca²⁺. Pipette solution contained 10 μM IP₃ and 18 nM free Ca²⁺. Current remains stable after store depletion–induced activation is complete. (B) Effect of divalent-free (buffered with 1 mM EDTA) bath solution on whole-cell current. Current was activated by active store depletion with 10 μM IP₃. Sodium current begins to inactivate immediately after removal of divalent cations from the bath. Voltage clamp protocol was the same as described in Fig. 9 A. Leak currents were subtracted from all current records. Experiments were performed in the presence of 5 mM free Mg²⁺ in the pipette solution to inhibit I_ORCa.

DISCUSSION

Functional Properties of C. elegans Intestinal Epithelial Cell Ca²⁺ Conductances

The C. elegans intestine provides a unique model system in which to characterize the molecular details of IP₃-dependent oscillatory Ca²⁺ signaling. To begin defining the functional roles and regulation of cation channels involved in Ca²⁺ signaling events, we performed patch clamp analysis of intestinal cells cultured in vitro (Christensen et al., 2002). Intestinal epithelial cells develop and survive in culture (Fig. 1) and are present at a frequency similar to that observed in newly hatched L1 larvae (see results).

Our initial patch clamp studies on intestinal cells were performed using “physiological” bath (5 mM K⁺ and 145 mM Na⁺) and pipette (143 mM K⁺ and 4 mM Na⁺) solutions described originally by Lockery and coworkers (Goodman et al., 1998). Under these conditions, whole-cell current showed strong outward rectification (unpublished data). Ion substitution studies demonstrated that an outwardly rectifying cation channel carries this current (Figs. 2–4; Table I). The channel conducts both monovalent and divalent cations and has high selectivity for Ca²⁺ over Na⁺ (P_Ca/P_Na = 64:1; Table I). Lanthanum, 2-APB, and SKF 96365 reversibly inhibited the current (Fig. 6), and intracellular Mg²⁺ inhibited channel activity with a K_1/2 of 692 μM (Fig. 8).

The outwardly rectifying Ca²⁺ current (I_ORCa) was constitutively active in all cells examined. Currents typically increased two to threefold after whole-cell access was obtained. Activation is not mediated by depletion of intracellular Ca²⁺ stores (Fig. 7). Channel activation may be mediated by washout of intracellular Mg²⁺, changes in protein phosphorylation, and/or other as yet undefined mechanisms.

I_ORCa shares some characteristics with MIC, Mg²⁺-nucleotide–regulated metal ion (MagNuM), and TRPM7 currents. These shared characteristics include cation selectivity, permeability to Ca²⁺, strong outward rectification, gradual activation after obtaining whole-cell access, and insensitivity to store depletion (Nadler et al., 2001; Hermosura et al., 2002; Kozak et al., 2002; Prakriya and Lewis, 2002). Importantly, the degree of inhibition of ORCa by Mg²⁺ is similar to that of the recently described MIC/MagNuM currents in RBL and Jurkat T cells (Hermosura et al., 2002; Kozak et al., 2002; Prakriya and Lewis, 2002).

TRPM7 and/or other TRP genes have been proposed to encode MIC/MagNuM (Nadler et al., 2001; Clapham, 2002; Prakriya and Lewis, 2002). The biophysical similarities between MIC/MagNuM and ORCa suggest that the channels may have a common molecular origin. However, there are also a number of significant differences between the channel types. For example, removal of extracellular Mg²⁺ and Ca²⁺ causes the I–V relationship to become linear for MIC, MagNuM, and TRPM7 (Nadler et al., 2001; Hermosura et al., 2002; Kozak et al., 2002; Prakriya and Lewis, 2002), but does not alter I_ORCa rectification (Fig. 4). I_ORCa shows voltage- and time-dependent gating (Fig. 4), whereas gating of TRPM7 is largely voltage insensitive (Runnels et al., 2001). MIC, MagNuM, and TRPM7 discriminate poorly between Ca²⁺ and Na⁺ (Nadler et al., 2001; Runnels et al., 2001) and Cs⁺ and Na⁺ (P_Cs/P_Na ≈ 1) (Runnels et al., 2001; Kozak et al., 2002; Prakriya and Lewis, 2002). In contrast, I_ORCa is highly selective for Ca²⁺ over Na⁺ and P_Cs/P_Na is 0.27 (Table I). Finally, Na⁺ current through ORCa is half blocked by ∼1 mM Ca²⁺ (Fig. 5 A), whereas MIC is half blocked by <5 μM Ca²⁺ (Kerschbaum and Cahalan, 1998). Studies focused on identifying the gene or genes that encode ORCa are currently underway.

Given the role of IP₃ and Ca²⁺ signaling in regulating defecation rhythm in C. elegans (Dal Santo et al., 1999), we performed a series of studies to determine whether intestinal cells also express SOCCs. To observe SOCC activity, we inhibited I_ORCa by addition of millimolar concentrations of free Mg²⁺ (Hermosura et al., 2002; Kozak et al., 2002) to the patch pipette solution.

In the absence of store depletion, whole-cell current was stable (Fig. 9 C). Active or passive depletion of Ca²⁺ stores activated a strongly inwardly rectifying current (Fig. 9, A and B). When stores were depleted actively by inclusion of 10 μM IP₃ in the patch pipette solution or by addition of 2 μM ionomycin to the bath, the rates of current activation were increased ∼8- and ∼22-fold compared with passive store depletion (Fig. 9 C; Table II). These results demonstrate clearly that Ca²⁺ store depletion activates an inwardly rectifying store-operated channel.

The inwardly rectifying channel is highly cation selective. Inward current was undetectable when bath Na⁺ and Ca²⁺ were replaced by NMDG⁺ (Fig. 10 A). Addition of Ca²⁺ to an NMDGCl bath solution induced inward current with an amplitude similar to that observed with Na⁺ and Ca²⁺ (Fig. 9 A; Fig. 10 A; Table II). In a divalent-free Na⁺-containing bath, E_rev was 13 mV (Fig. 10 A). Addition of 20 mM Ca²⁺ shifted E_rev to more positive potentials. However, it was not possible to measure E_rev accurately under these conditions because the slope of the I–V plot from 0 to +80 mV was extremely shallow (Fig. 9 A). Nevertheless, these results demonstrate that the store-operated channel is highly selective for Ca²⁺ over monovalent cations. If E_rev > +80 mV, the channel would have a Ca²⁺ to Na⁺ selectivity of at least 1,000:1.

CRAC is the most extensively characterized SOCC and is probably expressed ubiquitously in vertebrate cells (Parekh and Penner, 1997). The intestinal cell SOCC shares a number of characteristics with CRAC, including activation by passive and active store depletion, very strong inward rectification, and an apparent high selectivity for Ca²⁺ over monovalent cations (Hoth and Penner, 1993). Cation selectivity sequences of CRAC vary somewhat between cell types and are possibly altered by intracellular Ca²⁺ buffering (Zhang and McCloskey, 1995; Fierro and Parekh, 2000). Hoth and Penner (1992) and Zweifach and Lewis (1993) reported a CRAC divalent cation selectivity sequence of Ca²⁺ > Ba²⁺ ≈ Sr²⁺ in mast and Jurkat T cells. In RBL cells, Fierro and Parekh (1999) reported a slightly different divalent cation selectivity sequence of Ca²⁺ > Sr²⁺ > Ba²⁺. The selectivity sequence observed in mast and T cells is similar to that of the intestinal cell SOCC (Fig. 10 C).

CRAC monovalent cation selectivity sequences of Na⁺ ≈ Li⁺ > K⁺ > Cs⁺ (Voets et al., 2001) and Na⁺ ≈ Li⁺ > Rb⁺ > Cs⁺ (Bakowski and Parekh, 2002) have been observed in RBL cells. In T cells, Lepple-Wienhues and Cahalan (1996) reported a monovalent cation selectivity for CRAC of Na⁺ > Li⁺ = K⁺ > Rb⁺ ≫ Cs⁺. The monovalent selectivity sequence for the intestinal cell SOCC is Na⁺ ≈ Li⁺ > Rb⁺ ≈ Cs⁺ (Table III) and resembles that reported for RBL cells (Voets et al., 2001; Bakowski and Parekh, 2002). The most significant difference in CRAC and SOCC cation selectivity is their relative Cs⁺ permeabilities. CRAC has a P_Cs/P_Na of ∼0.1, whereas P_Cs/P_Na for SOCC is 0.6 (Table III).

TMA⁺ permeated SOCC nearly as well as Na⁺, whereas TRIS⁺ and NMDG⁺ had very low or negligible permeability (Fig. 10, A and B), suggesting that the channel has a minimum pore diameter of ∼0.6–0.7 nm. TMA⁺ permeation through CRAC in RBL cells is undetectable, and a pore diameter of 0.32–0.55 nm has been estimated (Bakowski and Parekh, 2002). In Jurkat T cells, TMA⁺ permeates CRAC, albeit poorly (Kerschbaum and Cahalan, 1998). Kerschbaum and Cahalan (1998) have estimated a minimum pore diameter for the T cell CRAC of at least 0.58 nm.

When exposed to divalent-free Na⁺ medium, CRAC undergoes a rapid inactivation (Christian et al., 1996; Lepple-Wienhues and Cahalan, 1996; Voets et al., 2001; Kozak et al., 2002; Prakriya and Lewis, 2002). In Jurkat T cells for example, CRAC activity declines up to ∼80% within ∼20 s after removal of bath Ca²⁺ (Prakriya and Lewis, 2002). Inactivation of CRAC may reflect extracellular Ca²⁺-dependent changes in channel gating (Zweifach and Lewis, 1995; Christian et al., 1996).

The C. elegans SOCC also inactivates in divalent-free medium (Fig. 12). However, this inactivation is considerably slower than that observed with CRAC. The SOCC-mediated Na⁺ current inactivates at a rate of −13%/min.

At present, it is not possible to conclude that homologous genes encode CRAC and the intestinal SOCC because the molecular identities of both channels are unknown. However, C. elegans clearly provides unique experimental advantages and opportunities for identifying SOCC-encoding genes. Identification of these genes may ultimately provide clues into the molecular identity of CRAC.

Role of Store-independent and Store-operated Ca²⁺ Channels in Oscillatory Ca²⁺ Signaling

Extracellular agonist–induced Ca²⁺ signaling in nonexcitable cells requires the release of Ca²⁺ from IP₃-regulated intracellular stores and the influx of Ca²⁺ across the plasma membrane via SMOCCs and SOCCs (Elliott, 2001; Zitt et al., 2002). High concentrations of agonists typically trigger sustained elevation of cytoplasmic Ca²⁺ levels. It is generally accepted that SOCCs play important roles in maintaining globally elevated Ca²⁺ concentrations and in refilling depleted Ca²⁺ stores (Parekh and Penner, 1997; Elliott, 2001). However, in the presence of lower, physiologically relevant agonist concentrations, Ca²⁺ changes are more complex, occurring in oscillations and waves and in localized areas of the cell (Shuttleworth, 1999; Berridge et al., 2000). The mechanisms responsible for generating Ca²⁺ oscillations are varied and depend on passive Ca²⁺ buffering, the spatial distribution of Ca²⁺ stores, rates of Ca²⁺ transport across the plasma membrane, and mitochondrial and ER Ca²⁺ uptake (Shuttleworth, 1999; Berridge et al., 2000; Bootman et al., 2001; Petersen, 2002). The specific roles played by SMOCCs and SOCCs in oscillatory Ca²⁺ signaling are unclear. Calcium oscillations in some cell types continue for long periods in the absence of extracellular Ca²⁺ (Lechleiter and Clapham, 1992), whereas oscillatory Ca²⁺ signals in other cell types are strictly dependent on Ca²⁺ influx (Torihashi et al., 2002; Wu et al., 2002).

Dolmetsch and Lewis (1994) have proposed that Ca²⁺ oscillations in T lymphocytes are driven primarily by pulsatile Ca²⁺ influx via CRAC. These investigators suggest that the main function of the intracellular Ca²⁺ stores is to control the extent and timing of CRAC activity (Dolmetsch and Lewis, 1994). Pharmacological studies in rat hepatocytes (Gregory and Barritt, 2003) and astrocytes (Pizzo et al., 2001) also suggest that Ca²⁺ oscillations are dependent on CRAC-mediated Ca²⁺ influx.

Shuttleworth and coworkers (Shuttleworth, 1999; Mignen et al., 2003) have noted that evidence for the involvement of CRAC specifically and SOCCs in general in generating Ca²⁺ oscillations is limited. Shuttleworth has also argued that CRAC possesses neither the sensitivity to store depletion nor the activation kinetics required for oscillatory Ca²⁺ signaling (Shuttleworth, 1999). Instead, he has suggested that the function of CRAC may be primarily to mediate plasma membrane Ca²⁺ influx required for refilling ER stores under conditions of sustained store depletion (Shuttleworth, 1999).

Recently, an arachidonic acid–regulated Ca²⁺ channel (ARC) has been described in many cell types (Mignen and Shuttleworth, 2000; Moneer and Taylor, 2002). It has been proposed that ARC is a major Ca²⁺ entry pathway required for oscillatory Ca²⁺ signaling (Shuttleworth, 1999; Mignen et al., 2001, 2003; see also Lankisch et al., 1999). In contrast, Luo et al. (2001) have suggested that an influx pathway distinct from both ARCs and SOCCs mediates Ca²⁺ entry that drives Ca²⁺ oscillations in HEK cells. The disparate conclusions of these studies underscore the need for extensive additional work to define the mechanisms of Ca²⁺ entry in nonexcitable cells, to define the role of SMOCCs and SOCCs in oscillatory Ca²⁺ signaling, and to identify molecular mechanisms of SMOCC and SOCC regulation.

As noted earlier, intracellular Ca²⁺ levels in the nematode intestine oscillate with a periodicity of 45–50 s (Dal Santo et al., 1999). These oscillations drive rhythmic contraction of body wall muscles and are dependent on IP₃ receptor function (Dal Santo et al., 1999). Both the ORCa and SOC channels could play central roles in generating and maintaining oscillatory Ca²⁺ signaling in the intestine. However, determination of their physiological functions requires in vitro and in vivo characterization of intestinal cell Ca²⁺ signaling events and identification of the genes that encode both channels. These studies are currently underway and will likely be facilitated by the genetic and molecular tractability of C. elegans as well as by the physiological accessibility of cultured intestinal cells.

Genetic and Molecular Analysis of the C. elegans Defecation Cycle

Mutagenesis and forward genetic analysis in C. elegans has to date identified ∼12 genes that disrupt normal defecation rhythm when they are mutated (Iwasaki et al., 1995). itr-1 and flr-1, which encodes a putative DEG/ENaC cation channel (Take-Uchi et al., 1998), are the only genes that have been mapped and characterized in detail. In addition, mutations in calcium/calmodulin-dependent serine/threonine kinase type II (CaMKII) have been shown to disrupt the defecation cycle (Reiner et al., 1999). CaMKII plays essential roles in oscillatory Ca²⁺ signaling (De Koninck and Schulman, 1998; Dupont and Goldbeter, 1998) and is expressed in multiple C. elegans cell types, including intestinal epithelial cells (unpublished observation cited in De Koninck and Schulman, 1998).

Mapping and characterization of mutant genes that disrupt defecation rhythm as well as isolation of other defecation mutants will likely provide unique insights into intestinal cell oscillatory Ca²⁺ signaling. It will also be important to utilize reverse genetic approaches to identify genes that encode the ORCa and SOC channels. At present, it is reasonable to postulate that the channels are encoded by one or more TRP genes (Montell, 2001; Clapham, 2002). The C. elegans genome contains 13 predicted TRP-encoding genes (3 TRPC, 5 TRPV, 4 TRPM, and 1 TRPN). Gene function in C. elegans can be rapidly and economically disrupted either by the use of chemical deletion mutagenesis or RNA interference (Barr, 2003; Strange, 2003). Deletion mutagenesis should allow definitive testing of the hypothesis that TRP genes encode the ORCa and SOC channels.

In conclusion, we have identified two highly Ca²⁺-selective cation conductances in C. elegans intestinal epithelial cells. One conductance is store independent, and the other is activated by store depletion. Our studies provide the first detailed electrophysiological characterization of voltage-independent and store-operated Ca²⁺ conductances in C. elegans. The ability to combine patch clamp electrophysiological measurements on intestinal epithelial cells with forward and reverse genetic analyses provides a powerful new approach for defining the cellular and molecular mechanisms of IP₃-dependent oscillatory Ca²⁺ signaling and its role in controlling rhythmic biological processes.