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J. Biol. Chem., Vol. 280, Issue 3, 1771-1781, January 21, 2005

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Orexin-A-induced Ca²+ Entry

EVIDENCE FOR INVOLVEMENT OF TRPC CHANNELS AND PROTEIN KINASE C REGULATION^*

Kim P. Larsson, Hanna M. Peltonen, Genevieve Bart, Lauri M. Louhivuori, Annika Penttonen, Miia Antikainen, Jyrki P. Kukkonen, and Karl E. O. Åkerman¶

From the A. I. Virtanen Institute for Molecular Sciences, Department of Neurobiology, Laboratory of Cell Biology, University of Kuopio, P. O. Box 1627, FIN-70211 Kuopio, Finland and the Department of Neuroscience, Division of Physiology, Uppsala University, BMC, P. O. Box 572, S-75123 Uppsala, Sweden

Received for publication, June 1, 2004 , and in revised form, October 28, 2004.

	ABSTRACT

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
The orexins are peptide transmitters/hormones, which exert stimulatory actions in many types of cells via the G-protein-coupled OX₁ and OX₂ receptors. Our previous results have suggested that low (subnanomolar) concentrations of orexin-A activate Ca²⁺ entry, whereas higher concentrations activate phospholipase C, Ca²⁺ release, and capacitative Ca²⁺ entry. As shown here, the Ca²⁺ response to subnanomolar orexin-A concentrations was blocked by activation of protein kinase C by using different approaches (12-O-tetradecanoylphorbol acetate, dioctanoylglycerol, and diacylglycerol kinase inhibition) and protein phosphatase inhibition by calyculin A. The Ca²⁺ response to subnanomolar orexin-A concentrations was also blocked by Mg²⁺, dextromethorphan, and tetraethylammonium. These treatments neither affected the response to high concentrations of orexin-A nor the thapsigargin-stimulated capacitative entry. The capacitative entry was instead strongly suppressed by SKF96365 An inward membrane current activated by subnanomolar concentrations of orexin-A and the currents activated upon transient expression of trpc3 channels were also sensitive to Mg²⁺, dextromethorphan, and tetraethylammonium. Responses to subnanomolar concentrations of orexin-A (Ca²⁺ elevation, inward current, and membrane depolarization) were voltage-dependent with a loss of the response around –15 mV. By using reverse transcription-PCR, mRNA for the trpc1–4 channel isoforms were detected in the CHO-hOX1-C1 cells. The expression of truncated TRPC channel isoforms, in particular trpc1 and trpc3, reduced the response to subnanomolar concentrations of orexin-A but did not affect the response to higher concentrations of orexin-A. The results suggest that activation of the OX₁ receptor leads to opening of a Ca²⁺-permeable channel, involving trpc1 and -3, which is controlled by protein kinase C.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Orexins act via two G-protein-coupled receptors called OX₁R and OX₂R (1, 2). They activate neurons and secretory cells by mechanisms that are not fully understood (reviewed in Refs. 3 and 4). Interaction of orexin receptors with G-proteins of the G_q/11, G_i, and G_s families has been suggested based on second messenger assays and covalent labeling of G-proteins with azido-GTPS¹ (5). The most typical responses to orexins in neurons include increased excitability, membrane depolarization (5–10 mV), and Ca²⁺ elevation (3, 4). Evidence for several different mechanisms have been proposed to explain these responses, including activation of a nonselective cation current, a decrease in K⁺ conductance, and activation of Na⁺/Ca²⁺ exchange. The orexin-stimulated Ca²⁺ elevation in neurons shows an explicit dependence on extracellular Ca²⁺ and is therefore likely to be due to Ca²⁺ entry into cells (5–7). Activation of recombinantly expressed orexin receptors in nonexcitable cells also leads to Ca²⁺ elevation (1, 8–11). High concentrations (>10 nM) of orexins induce intracellular Ca²⁺ release (9), but with lower concentrations of orexins the Ca²⁺ elevations observed are dependent on extracellular Ca²⁺ (8–11) and do not appear to involve measurable discharge from stores (9). Similar results are observed upon recombinant expression of orexin receptors in neuron-like excitable cells (PC12 and Neuro2A). Activation of a novel Ca²⁺ influx pathway was thus suggested. The existence of such a pathway is also indirectly suggested by the dependence of the Ca²⁺ response on a negative internal membrane potential and the activation of a robust influx of Mn²⁺ ions in the absence of store discharge (9). The identity of this Ca²⁺ influx pathway and the mechanisms involved in its activation remain unresolved. Because the orexin-stimulated [Ca²⁺]_e-dependent Ca²⁺ elevation is relatively insensitive to blockers of capacitative Ca²⁺ entry, such as lanthanides and 2-APB, but blocked by Ni²⁺, a different molecular entity was proposed (10). Several different pathways for receptor-activated Ca²⁺ entry have also been suggested based on functional studies with other receptors. These include store-operated Ca²⁺ channels and second messenger-operated channels as well as Ca²⁺-activated Ca²⁺ channels (reviewed in Ref. 12). A large family of potential receptor-activated channels called transient receptor potential channels (TRP channels) has been identified (reviewed in Refs. 13–15). When recombinantly expressed, the different TRP channel subtypes produce currents that are, to various extents, dependent on extracellular Ca²⁺ and Na⁺. The mechanism by which receptors couple to activate these channels has not been clarified, but recombinantly expressed TRP channels have been shown to modify receptor-activated Ca²⁺ influx, and it has thus been suggested that they represent the molecular entities of receptor-activated pathways (13). It has been shown by using single cell RT-PCR that both orexin receptors are co-expressed with several members of the TRPC channel family in e.g. rat aminergic neurons, but the expression profile varies significantly between different cells (16). Some TRP channel subtypes are regulated by lipid products such as diacylglycerol, i.e. exogenously applied diacylglycerol analogs activate trpc3 (17–20), although endogenously produced diacylglycerol appears to suppress the activation of some of these channels through a protein kinase C (PKC)-dependent mechanisms (21–23). Calyculin A, a protein phosphatase inhibitor, strongly suppresses the activity of TRP and TRPL channels in Drosophila (23) and causes internalization of trpc1, -3, and -4 in human neutrophils and overexpressed trpc3 in HEK293 (24, 25), which suggests that TRP channels are under the control of phosphorylation/dephosphorylation reactions. The TRP channels are widely distributed in different cells and the subtypes appear to represent subunits of larger channel complexes (26, 27). Interaction of expressed channels with endogenous channel complexes and constitutive activation further complicates the assessment of the functional properties of individual TRP channels (13, 14), whereas the identification of the specific function of endogenous TRP channels has, especially in neurons and endocrine cells, been hampered by the lack of specific pharmacological blockers.

The goal of this study was to investigate whether the primary orexin-A-activated Ca²⁺ entry (mediated by OX₁ receptor) involves TRPC channels using the CHO-hOX1-C1 cell line as an experimental system. Because excitable and nonexcitable cells express the same G-proteins and TRP channel subtypes (with the exception of TRPC5), their basic signaling mechanisms are expected to be the same or quite similar. A panel of channel blockers was used to distinguish the orexin-activated Ca²⁺ influx from store-operated influx, and patch clamp recordings were used to define the properties of the pathway. TRPC channel mRNA profiling was used to determine the best targets for interference with the function of the endogenous TRPC channel by using transient expression of truncated TRPC constructs and thus to assess their involvement in the orexin-stimulated Ca²⁺ entry. In addition the regulation of the orexin-activated pathway by PKC was tested by using PKC activation and inhibition.

	EXPERIMENTAL PROCEDURES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
The generation of CHO-hOX₁-C1 cells stably expressing the human OX₁R, has been described earlier (9). Cells were grown in Nutrient Mixture (Ham's F-12) medium (Invitrogen) supplemented with 100 units/ml penicillin G (Sigma), 80 units/ml streptomycin (Sigma), 400 µg/ml geneticin (G418; Invitrogen), and 10% (v/v) fetal calf serum (Invitrogen) at 37 °C in 5% CO₂ in an air-ventilated humidified incubator in 260-ml culture flasks (Nunc A/S, Roskilde, Denmark). For Ca²⁺ measurements in suspension, the cells were grown in 800-ml culture flasks (Nunc) in order to obtain a larger quantity of cells.

Materials—2-Aminoethoxydiphenyl borate (2-APB), GF109203X (bisindolylmaleimide), R59022 [GenBank] (diacylglycerol kinase inhibitor), and SKF96365(1-[-(3-(4-methoxyphenyl)propoxy)-4-methoxyphenethyl]-1H-imidazole-hydrochloride) were from Calbiochem. BzATP (2',3'-O-(4-benzoyl-benzoyl)-ATP), calyculin A, dextromethorphan, dioctanoyl glycerol (DOG), 1,2-dicapryloyl-sn-glycerol, EDTA, EGTA, probenecid (p-[dipropylsulfamoyl]benzoic acid), ATP (Mg²⁺ salt), and GTP (Na⁺ salt), NiCl₂, TPA (12-O-tetradecanoylphorbol-13-acetate), and TEA (tetraethylammonium) were from Sigma. Digitonin was from Merck. Fura-2-acetoxymethyl ester and fura-2-penta-potassium salt were from Molecular Probes (Eugene, OR). Thapsigargin was from RBI (Natick, MA). Human orexin-A was from Peninsula Laboratories Europe Ltd. (St. Helens, UK).

Media—The HEPES-buffered Na⁺ medium (HBM) consisted of the following (in mM): 137 NaCl, 5 KCl, 1 CaCl₂, 0.44 KH₂PO₄, 4.2 NaHCO₃, 10 glucose, 1 probenecid, 20 HEPES, and 1.2 MgCl₂; and the pH was adjusted to 7.4 with NaOH. TEA was used by replacing the Na⁺ in HBM. The desired TEA concentration was prepared by mixing the TEA-based HBM with Na⁺-based HBM. In HBM prepared for electrophysiology, unless otherwise specified, MgCl₂ was in general excluded. The intracellular electrode solution used in the whole-cell voltage clamp recordings consisted of the following (in mM): 136 Cs⁺ aspartate, 30 HEPES, 10 NaCl, 4 ATP, and 0.6 GTP. In current clamp recordings a similar intracellular solution was used but with 136 K⁺ aspartate. The [Ca²⁺] in the intracellular electrode solution was optically measured with fura-2-pentapotassium salt and calibrated to 140 nM by addition of 50 µM EGTA and 25 µM fura-2. The effect of a high intracellular Ca²⁺ buffer capacity was tested in some experiments by increasing the concentrations of EGTA and Ca²⁺ to 4 and 1 mM, respectively, or 10 and 2.8 mM, respectively. Finally, the pH was set to 7.25 with CsOH or KOH.

Ca²⁺ Measurements in Suspension—The fluorescent Ca²⁺ indicator fura-2 (28) was used to monitor changes in the intracellular Ca²⁺ concentration ([Ca²⁺]_i) as described previously (9). Briefly, the cells were harvested using phosphate-buffered saline containing 0.2 g/liter EDTA, spun down, and loaded at 37 °C in HBM, 1 mM probenecid, and 4 µM fura-2 acetoxymethyl ester for 20 min. The cells were washed once with Ca²⁺-free HBM and stored on ice as pellets (medium removed). The measurement of [Ca²⁺]_i was carried out as follows: 1 pellet was resuspended in HBM at 37 °C and placed in a stirred quartz microcuvette in a thermostated cell-holder within a fluorescence spectrophotometer. Fluorescence was monitored with a PTI QuantaMaster fluorescence spectrophotometer at the wavelengths 340/360/380 (excitation) and 505 nm (emission). The experiments were calibrated by using 60 µg/ml digitonin, which gives the maximum value of fluorescence (F_max), and 10 mM EGTA, which gives the minimum value of fluorescence (F_min).

Combined Patch Clamp and Ca²⁺ Imaging—Orexin-A-evoked Ca²⁺ currents were studied in voltage clamp mode at 28 °C by using the standard whole-cell configuration (29) while concurrently monitoring the [Ca²⁺]_i by fura-2 imaging. The [Ca²⁺]_i in cells in the vicinity of the patched cell was monitored as controls. Cells were harvested from 260-ml cell flasks and replated on the day of use on 22 x 22-mm coverglasses (Warner Instruments Inc.) to a confluence of 50–70% and rested for a minimum of 3 h before use. Cells on a coverglass were loaded with 2 µM fura-2 acetoxymethyl ester and 1 mM probenecid for 20 min at 37 °C in HBM. The coverglass was then attached to the bottom of an RC-24 fast exchange chamber (Warner Instruments Inc.) and positioned on top of the microscope. Cells were perfused with HBM (2.5 ml/min giving an exchange time of 3 s) by a gravity-controlled drug delivery system. The perfusates were converging in a perfusion manifold and funneled through an SH-27B in-line heater (Warner Instruments Inc.) located just before the chamber inlet to obtain the desired temperature. Patch pipettes (model PG150T, Harvard Apparatus, UK) were prepared with a PC-10 puller and flame-polished with microforge MF-900 (Narishige, UK) to a resistance of 3.6–3.8 megohms measured in the bath solution. The patch clamp amplifier Axopatch 200A was connected to a computer via the AD/DA Digidata1320E SCSI interface (Axon Instruments). Voltage protocols and data acquisition were controlled with pClamp 8.1 (Axon Instruments). Cells were compensated for the pipette capacitance, whereas following whole-cell access the series resistance was analogically compensated to 60–70%. Liquid junction potential was calculated in pClamp 8.1 and subtracted from the recordings giving a more accurate clamping potential of –60 mV. In general, voltage ramps (–80 to +80 mV; 320 ms) were applied every 5 or 7.5 s. Data were digitally sampled at 3.8 kHz and filtered at 2 kHz by using the low pass Bessel filter on the recording amplifier. In current clamp experiments, data were digitally sampled at 5 kHz and filtered at 2 kHz. Combined fluorescence recordings were obtained with a second computer running the TILLvisION Multi-Color Ratio ImagingSystem (TILL Photonics GmbH, Gräfelfing, Germany), and saved for later analysis. The system consisted of a polychrome IV and a 12-bit IMAGO CCD camera under control of an external control unit. An inverted microscope (Nikon) was used to visualize the fluorescence. UV light was guided through an epifluorescence condenser, and cells were excited through a dichroic mirror (DM430, Nikon). The emission was measured through a 510 nm cut-off filter (Nikon). The imaging protocol was designed to acquire images at 340 and 380 nm every 1–3 s. A TTL trigger pulse synchronized the patch clamp and imaging recordings; the TTL pulse was controlled by TILLvisION 4.0 to trigger the voltage clamp data acquisition by using the "digitizer start input" option in the pClamp 8.1. After ending the recordings, fluorescence from 340 and 380 nm of selected regions of interest were analyzed and converted into [Ca²⁺]_i as described previously (9). Voltage clamp and image data were then combined in Microcal Origin^TM 6.0 for visualization and final analysis.

Identification of TRP Channel mRNA—For primer design, nucleotide sequences, retrieved from the GenBank^TM data base, were aligned with MacMolly Tetra (version 3.10, align ppc program, Soft Gene GmbH). 0.5 µg of total RNA were reverse-transcribed using SuperscriptII cDNA synthesis kit (Invitrogen) and then amplified by using general trpc-specific degenerate primers 5'-nggvmchytgcagathtc-3' and 5'-nckhgcaaayttccaytc-3'; the PCR conditions were as follows: 95 °C for 5 min, 50 °C for 30 s, 72 °C for 30 s, and 94 °C for 30 s, 30 cycles. Amplified DNA was gel-purified and inserted into PgemTeasy plasmid (Promega) and sequenced. PCR product identification was done using Blast program (30). For expression/comparison analysis, specific primers for each trpc mRNA subtype were designed and tested, PCR conditions were optimized (Table I). 1 or 0.5 µl (trpc1/trpc2) of the 20-µl cDNA reaction were amplified with channel-specific primers using optimized conditions. Identical amounts of PCRs were run on a 1.5% agarose TBE gel, stained with SYBRgreenI (Molecular Probes) according to manufacturer's instructions, and scanned on Storm 860 (Amersham Biosciences). Quantification of signal was done using ImageQuant program.

Transfection and Ca²⁺ Imaging—For experiments, cells were seeded in 35-mm inner diameter Petri dishes (Nunc, Roskilde, Denmark) containing a coverslip (25 mm inner diameter, Merck Eurolab, Espoo, Finland) at a density of about 125,000 cells per plate in 2 ml of medium. After 18–24 h, cells were transfected with 3 µl of FuGENE 6 (Roche Applied Science) and 1 µg of DNA, according to the manufacturer's recommendations. Cells were used within 24 h of transfection. Expression of the GFP-tagged truncated channel isoforms was detected with 450–480 nm UV light and 520 nm barrier filter. The Ca²⁺ imaging experiments were performed, and the data were analyzed by using the intracellular imaging InCyt2^TM fluorescence imaging system (Cincinnati, OH). In brief, the cells were perfused with HBM at 37 °C and excited by alternating wavelengths of 340 and 380 nm by using narrow band excitation filters, and the fluorescence was measured through a 430 nm dichroic mirror and a 510 nm barrier filter with a Cohu CCD camera. Fluorescence from 340 and 380 nm exposures were imported into Microcal Origin^TM 6.0, and the ratios were calculated. Day to day variance in the orexin-A responses was cancelled out by normalizing Ca²⁺ responses in individual cells to a control response evoked by 100 µM 2',3'-O-(4-benzoyl-benzoyl)-ATP (BzATP) at the end of an experiment. CHO-K1 cells have been shown previously to respond to BzATP via activation of P2X (P2_Z/P2X₇) receptors (36), and this response should not be affected by transfection. Cells were divided into responding and nonresponding groups, determined by their response to 0.3 nM orexin-A, and counted for statistical presentation. Nonresponding cells were then discarded in additional analysis, whereas the peaks in responding cells were further processed.

Trp Co-immunoprecipitation—mTrpc1-FLAG and trpc1N-EGFP (tarpon) were co-transfected into CHO-hOX1-C1 cells at a ratio of 0.5 µg/1.5 mg in a 60-mm inner diameter Petri dish. After 24 h, cells were harvested in PBS and lysed in RIPA 1% Triton X-100, incubated for 30 min on ice, and spun for 15 min at 15,000 x g at 4 °C. The supernatant was pre-cleared 15 min with protein G beads (Dynal, Oslo, Norway) and then mixed with 2 µg of polyclonal anti-full-length GFP antibody (BD Biosciences) cross-linked to protein G beads and mixed for 1 h at room temperature. Beads were captured and washed three times with RIPA and then eluted with SDS-PAGE sample buffer. Initially, 1.5 µg of trpc3FLAG was transfected into HEK293 cells using PEI50 (37) (Sigma) in a 35-mm inner diameter plate. The following day, 1.5 µg of trpc3N was transfected into the same cells, and successful transfection was determined by fluorescence microscopy (trpc3FLAG transfected cells are uniformly green; trpc3N transfected cells have localized fluorescence, and dually transfected cells have localized fluorescence in a fluorescent background). Cells were scraped in PBS and resuspended in 50 mM Tris, 120 mM NaCl, 0.5% IGEPAL, Complete^TM protease inhibitors (Roche Applied Science) and spun 15 min at 15,000 x g at 4 °C. After preclearing and antibody-protein G incubation, beads were washed five times in high salt buffer (0.9 M NaCl) and 1 times in 0.1 M NaCl and eluted in SDS-PAGE buffer. Lysate and nonattached and eluted fraction were run on 7.5% acrylamide gel, blotted on polyvinylidene difluoride membrane, and probed with monoclonal anti-GFP (BD Biosciences) and anti-FLAG M2 (Stratagene). Detection was done with ECL-Plus (Amersham Biosciences) and visualized with Storm 860 (Amersham Biosciences) by using the program ImageQuant.

Data Processing—The differences in the responses between two groups were evaluated by the unpaired Student's t test. Between more than two groups the one-way ANOVA test was used followed by Scheffe's test. Significance is presented for p < 0.05 and p < 0.01. Data are expressed as means ± S.E., and n (where indicated) indicates the number of cells or experiments.

	RESULTS

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Effect of Mg²⁺ and Ion Channel Blockers on Orexin-A Evoked Ca²⁺ Elevation in Cell Suspensions—To distinguish the orexin-activated Ca²⁺ influx pathway from intracellular release and capacitative Ca²⁺ entry, we tested the effect of different inhibitors of cation channels on the response to low and high concentrations of orexin-A (Ox-A) and capacitative entry activated by thapsigargin in CHO-hOX₁-C1. Ca²⁺ measurements in suspension are shown in Fig. 1. Mg²⁺ ions have been shown previously to block a variety of Ca²⁺-permeable channels including members of the TRP channel family (38–40). As shown in Fig. 1A (contr) Ox-A at a concentration of 0.3 nM caused a robust elevation of [Ca²⁺]_i. Increasing the extracellular Mg²⁺ from 1.2 to 5 mM caused a reduction in the response (Fig. 1A, Mg²⁺, also see bar diagram in Fig. 1E). A higher concentration of Mg²⁺ (20 mM) did not cause a further inhibition of the response (n = 5, data not shown). The Ca²⁺ elevation seen at higher concentrations of Ox-A (3 nM or above) was unaffected by elevated Mg²⁺.

View larger version (38K): [in this window] [in a new window]   FIG. 1. Effects of Mg2+, dextromethorphan, tetraethylammonium, and SKF96365on Ox-A or thapsigargin evoked Ca2+ elevation. A, fura-2 recordings from cell suspensions are shown in response to addition of 0.3 nM Ox-A in the absence (contr) and presence of 5 mM Mg2+ or 100 µM dextromethorphan (Dex 100). Test substances were added 2 min prior to Ox-A. The response to Ox-A is completely abolished by Dex and partially by Mg2+. B, similar recordings are shown as in A, but with 10 nM Ox-A and 300 µM Dex (Dex 300) additionally tested. C, 100 nM thapsigargin (thaps) was added in the presence of various substances. In Ca2+-free conditions (–Ca2+), no stable phase was observed. Addition of 10 µM 2-aminoethoxydiphenyl borate (2-APB) is shown to demonstrate the capacitative Ca2+ entry. 5 mM Mg2+ or 100 µM Dex is also shown following a 2-min preincubation. Mg2+ is not affecting the response, whereas Dex shows a reduction in the stable phase without affecting the peak. Scale bars in A–C represent 50 s. D shows the effect of increasing concentrations of Dex on Ox-A-evoked peak responses. The Ox-A concentrations tested are indicated by 0.3, 1, and 10 (nM). Data are obtained with similar recordings as in A–C and presented as means ± S.E. following normalization to control responses (contr). Each point is an average of 5–12 experiments. E, a bar diagram is shown for the fura-2 recordings performed as in A–C showing effects of 5 mM Mg2+, 70 mM TEA, 100 µM Dex, and 10 µM SKF96365(SKF). The cells were pretreated with the substances for 2 min before stimulation. The data are normalized to the respective control peak response or stable phase after 100 s, respectively, (n = 6 ± S.E.).

In order to test the effect of channel blockers on capacitative Ca²⁺ entry, the cells were exposed to 100 nM thapsigargin, which releases Ca²⁺ from intracellular stores and causes subsequent activation of store-operated pathways. When thapsigargin was added in the presence of extracellular Ca²⁺, a long lasting elevation of [Ca²⁺]_i was observed (Fig. 1C, contr). Addition of 10 µM 2-APB, a blocker of capacitative Ca²⁺ entry, reversed the response to thapsigargin when added during the stable phase (Fig. 1C, 2-APB). In line with this, removal of extracellular Ca²⁺ immediately prior to thapsigargin addition only evoked a transient Ca²⁺ elevation that returned to base line after 100 s (Fig. 1C, –Ca²⁺). Introduction of 5 mM Mg²⁺ did not significantly affect the response to thapsigargin-induced Ca²⁺ elevation (Fig. 1C, Mg²⁺), whereas in the presence of 100 µM dextromethorphan only a small reduction in the stable phase was observed (Fig. 1C, Dex 100). The effect of 1–300 µM dextromethorphan was further investigated on peak responses evoked with 0.3, 1, and 10 nM Ox-A. As shown in Fig. 1D, dextromethorphan caused a significant inhibition of the response evoked with 0.3 nM Ox-A. This suggests that the effect of dextromethorphan is noncompetitive.

The mean responses (±S.E.) of the inhibitors tested are summarized in Fig. 1E. Elevated Mg²⁺ inhibited the response to 0.3 nM Ox-A by about 70% but had little or no effect on the peak or stable response to 10 nM Ox-A or 100 nM thapsigargin. Dextromethorphan at 100 µM strongly inhibited the effect of 0.3 nM Ox-A and had no effect on the peak response but partially inhibited the stable phase of the response to 10 nM Ox-A. It did not significantly affect the response to 100 nM thapsigargin. The nonspecific potassium channel blocker TEA was also tested under similar conditions and had, at 70 mM, an effect very similar to that seen with Mg²⁺. SKF96365(10 µM), a blocker of Ca²⁺ entry (43), had little effect on the response to 0.3 nM Ox-A, and it partially inhibited the peak and stable phase of the response to 10 nM Ox-A and strongly inhibited the stable response to thapsigargin.

Effect of Protein Kinase C Stimulation and Inhibition of Ox-A Evoked Ca²⁺ Elevation in Cell Suspensions—Ca²⁺ measurements in suspension were used to test the effect of diacylglycerols, which activate some subtypes of TRP channels independently of receptor activation (17–20). Addition of 30 µM DOG caused a slow increase in [Ca²⁺]_i by 100 nM, which was dependent on extracellular Ca²⁺ (data not shown). This indicates the presence of diacylglycerol-activated Ca²⁺ entry in these cells. The [Ca²⁺]_i elevation in response to 0.3 nM Ox-A (Fig. 2A, contr) was considerably attenuated by the presence of DOG (Fig. 2A). At a concentration of 100 nM the phorbol ester TPA also attenuated the response to 0.3 nM Ox-A (Fig. 2B). The response to 100 nM thapsigargin was unaffected by DOG (Fig. 2C). Neither of these PKC activators significantly affected the response to 10 nM Ox-A (Fig. 2, D and E). The data summarized in Fig. 3A show that 10 µM GF109203X, an inhibitor of PKC, caused a small increase in the response to 0.3 nM Ox-A and almost completely reversed the inhibitory effect of DOG. A diacylglycerol kinase inhibitor R59022 [GenBank] (30 µM) (Fig. 3A, DAGKI) also reduced the response to 0.3 nM Ox-A. Likewise, this response was partially rescued by GF109203X. The peak [Ca²⁺]_i elevation and stable phase in response to 100 nM thapsigargin were unaffected by DOG or GF109203X (Fig. 3B). TPA inhibited the response to low concentrations of Ox-A (0.3 and 1 nM) but had little effect on the peak or stable response at 10 nM (Fig. 3C). As shown in Fig. 3D, preincubation with 100 nM calyculin A, a protein phosphatase blocker, for 10 min inhibited the effect of 0.3 nM Ox-A but did not affect the response to 10 nM Ox-A.

View larger version (24K): [in this window] [in a new window]   FIG. 2. Effects of DOG and TPA on Ox-A or thapsigargin evoked Ca2+ elevation. Fura-2 recordings were performed as in Fig. 1. The cells were preincubated with 30 µM DOG or 100 nM TPA for 2 min before challenge with 0.3 nM orexin-A (Ox-A) (A and B), 100 nM thapsigargin (thaps) (C), or 10 nM Ox-A (D and E), respectively. Controls traces are denoted as contr. Scale bars represent 50 s.

View larger version (30K): [in this window] [in a new window]   FIG. 3. Effects of PKC modification on Ox-A and thapsigargin evoked Ca2+ elevation. Results are obtained under same experimental condition as in Figs. 1 and 2. A, cell suspensions were challenged with 0.3 nM orexin-A (0.3 Ox-A), and the effects of the protein kinase C inhibitor 10 µM GF109203X (GF), 30 µM dioctanoylglycerol (DOG), and 30 µM of the DAG kinase inhibitor (R59022 [GenBank] ) (DAGKI), as well as the effect of GF on the two latter, are shown. The inhibitory effect of DOG and DAGKI is reversed by GF. B, the effect of GF and DOG on the peak (thaps peak) and stable response (thaps stable) to thapsigargin is shown. C, the effect of 100 nM TPA on the response to increasing concentrations of Ox-A is shown (0.3, 1, and 10 nM). D, cells were preincubated for 10 min in the absence or presence of 100 nM protein phosphatase inhibitor calyculin A (cycA), and the cells were challenged either with 0.3 or 10 nM Ox-A.

View larger version (30K): [in this window] [in a new window]   FIG. 4. Orexin A-activated Ca2+ elevation and membrane current: effects of extracellular Ca2+, Mg2+, dextromethorphan, and tetraethylammonium. The membrane potential was clamped in the whole-cell mode to –60 mV. Cs+-based intracellular solution was used. The currents in unchallenged conditions ranged from 9 to 22 pA. Scale bars indicate 25 pA and 25 s, respectively. A, the effect of extracellular Ca2+ removal on the current and fura-2 response (340/380 nm ratio) to 0.3 nM Ox-A using combined patch clamp and Ca2+ imaging is shown. Removal of extracellular Ca2+ reversibly abolishes both responses. B, the relation of the elevation in [Ca2+]i (depicted on the left y axis) and the concomitant current response (depicted on the right y axis) is shown as a function of increasing Ox-A concentration. Points are an average of 5 experiments ([Ca2+]) whereas the average currents are obtained from 5 to 27 experiments. Data are presented as means ± S.E. C and D, the effect of Mg2+ on the current responses to 0.3 nM Ox-A is shown. C, current reduction when Mg2+ is elevated to 5 mM (in a HBM-buffer including 1.2 mM Mg2+) is shown, and in D an increase in the current when 5 mM Mg2+ is removed is shown. E, the effect of 100 µM Dex and in F the effect of 70 mM TEA are shown. Dex and TEA completely and reversibly abolish the response to 0.3 nM Ox-A.

Ox-A-evoked Current-Voltage Relation, Depolarization, and Ca²⁺ Elevation Using Patch Clamp and Ca²⁺ Imaging—In order to analyze the voltage dependence of the Ox-A-activated membrane current, experiments were conducted with a voltage protocol introducing voltage ramps (–80 to +80 mV; 320 ms) every 5 or 7.5 s. Fig. 5A shows a current recorded at –60 mV (ramp traces not shown) in response to 0.3 nM Ox-A. The current-voltage profiles of ramps extracted before and during applications of Ox-A are shown in Fig. 5B (indicated by numbers 1 and 2). There was an approximately linear increase in inward current with increasing negative intracellular polarity.

View larger version (28K): [in this window] [in a new window]   FIG. 5. Current-voltage relationship and depolarization evoked by Ox-A. Currents were recorded as described in Fig. 4, and voltage ramps were introduced every 5 or 7.5 s (–80 to +80 mV; 320 ms). A, current response to 0.3 nM Ox-A is shown. Scale bars indicate 30 pA and 50 s, respectively. Arrows with numbers 1 and 2 indicate the position of extracted current-voltage traces shown in B. The current deflection during the ramp was removed from the trace for clarity. B, current-voltage traces are shown in the absence (trace 1) and presence of Ox-A (trace 2). C, whole-cell current clamp recording is shown in which 0.3 nM Ox-A evokes a reversible 7-mV depolarization. The intracellular solution was K+-based. Scale bar represents 50 s. D, the correlation of the resting membrane potential and the depolarization evoked by 0.3 nM Ox-A are shown. Note that the decline in depolarization with increasing membrane potential is correlating well with the current-voltage profile shown in B. The resting membrane potential and the depolarizing effect of substituting Na+ with K+ in the extracellular solution are shown in the bar diagram (E). Data are presented as means ± S.E. (n = 16). F and G, Ca2+ imaging on individual cells on coverslips is shown. F, Ca2+ responses to 0.3 and 3 nM are shown in cells on a coverslips in the absence and presence of a K+ substitution. In this series the following were used: 1) 0.3 nM Ox-A was added as control; 2) Ox-A 0.3 nM was added in the presence of K+;3)3nM was added as control; and 4) 3 nM was tested in the presence of K+. Washout between each application was 5 min. Data are presented as mean ± S.E. (n = 14 cells). Scale bar represents 100 s. G, summarizes the mean ± S.E. of six different experiments (98 cells) run under same conditions as in F. The majority of the response to low Ox-A (0.3 nM) disappeared during the K+ depolarization, whereas the response to a high concentration (3 nM) was only marginally affected.

Substitution of extracellular Na⁺ with K⁺ caused a considerable depolarization of the cells from a resting potential of –40 ± 5 to –13 ± 3 mV (n = 16, Fig. 5E). The membrane depolarization was rapidly reversed when cells again were exposed to the Na⁺-based external medium (data not shown). In line with this, Ca²⁺ imaging of intact cells showed that extracellular K⁺ substitution considerably reduced the response to 0.3 nM Ox-A but had little effect on the peak elevation at 3 nM Ox-A (Fig. 5, F and G).

In order to exclude that the effects of the blockers used above would be due to changes in membrane potential, their effects were investigated on the resting membrane potential using current clamp that was found to be –41 ± 2 mV (n = 14). Neither 100 µM dextromethorphan (n = 3) nor 20 µM 2-APB (n = 3) affected the membrane potential, whereas 5 mM Mg²⁺ and 70 mM TEA caused marginal depolarization of 0.3 ± 0.6 mV (n = 4) and 0.4 ± 0.9 mV (n = 4), respectively.

Detection of TRPC Channel Isoforms Using RT-PCR—In order to investigate the possible role of TRPC channels in the responses to Ox-A, we first investigated the presence of the mRNA for different channel subtypes (subunits). For identification of trpc mRNA, primers were designed by using alignment of mammalian trpc1–7 nucleotide sequences, available from the GenBank^TM data base. In the putative pore region, two very conserved sequences of 18 and 19 nucleotides, 350–400 bp apart (depending on the channel type), were identified, encompassing channel type-specific sequences that were conserved between different species. Initial RT-PCR amplification products were cloned and analyzed by restriction digestion. Several different clones were subsequently sequenced and identified as Chinese hamster homologues of trpc1, trpc2, and trpc3. In order to confirm that significant amounts of mRNA for trpc1, trpc2, and trpc3 were present in the cell line used, as well as to clarify whether other types of trpc channel mRNA were present, channel specific primers were designed. All available sequences from each individual channel from a wide range of mammals (including partial sequences and our sequences) were aligned. Specific primers sequences were selected from the same pore region (except for trpc4 primers that were located at the 5' end) and had the following features, the amplification product sequence would be highly conserved between organisms and channel type-specific sequences. This way trpc4 was detected, and the presence of trpc1, trpc2, and trpc3 mRNAs was confirmed. Quantification of relative amounts of the PCR product for each channel-specific primer pair (Fig. 6) indicates that trpc1 and trpc2 mRNA are the most abundant, whereas trpc3 and trpc4 mRNAs are present in lesser quantities. trpc5, trpc6 and trpc7 mRNAs were not detected, although several primer pairs were used (which produced fragments of the expected size from other cell lines and rat brain).

View larger version (35K): [in this window] [in a new window]   FIG. 6. Expression of trpc channel mRNA in CHO-hOX1-C1 cells. 6 µl of each PCR-amplified DNA was loaded on 1.5% agarose gel and post-stained with SYBRgreenI. Bands (upper panel) were visualized by UV transillumination (image captured using Bio-Rad Gel Doc 2000 with program Quantity One version 4.1.1). Lower panel shows the quantification of blue fluorescence from gel (as described under "Experimental Procedures"). Channel of trpc subtypes are listed on the x axis, and the y axis is the arbitrary units representing fluorescence per bp.

Fig. 7A shows a representative mean (±S.E.) of the Ca²⁺ response of the cells on a coverslip challenged with 0.3 and 3 nM Ox-A as well as 100 µM BzATP. In cells expressing the trpc1N construct (fluorescent cells), the response to 0.3 nM Ox-A was considerably attenuated. The response to BzATP was unaffected by the transfection. This suggests that the driving force (membrane potential) is comparable in transfected (fluorescent) and nontransfected cells (see below) because the response to BzATP mainly acts at endogenous expressed P2X₇ ion channels (36). BzATP was thus used as an internal control for the ability of the cell in question to respond. BzATP at this concentration gives a robust Ca²⁺ elevation in virtually all cells. Day to day variations and interference of the transfection procedure may also alter the quantification of the fura-2 signals. In some batches of cells a reduction in the responsiveness in fluorescent cells (regardless if the cells expressed fusion proteins or GFP alone) was observed in comparison to nonfluorescent cells. The responses were therefore normalized to the response to BzATP. No significant difference between GFP fluorescent and nonfluorescent cells was found, when the responses to 3 nM Ox-A were normalized to the BzATP response (Fig. 7B). Furthermore, the expression of GFP alone did not significantly affect the Ca²⁺ response to 0.3 nM Ox-A and was found to be 96.0 ± 1.5% of the nonfluorescent cells (one-way ANOVA test, p = 0.34, n = 80 experiments, 583 cells). Expression of the trpc7N construct evoked a similar response as expression of GFP alone. The trpc1N and trpc3N constructs caused a significant reduction of the Ca²⁺ response to 0.3 nM Ox-A, whereas trpc2N and trpc4N only had a marginal effects. The inhibitory effect of overexpressing truncated trpc channels was also reflected in the percentage of cells responding to 0.3 nM Ox-A. Although there was no difference in the number of cells responding to 3 nM Ox-A (and also BzATP) in the two groups (fluorescent versus nonfluorescent cells), the reduction in the number of cells responding to 0.3 nM Ox-A with the trpc1N construct was found to be 21%, trpc2N = 3%, trpc3N = 11%, trpc4N = 13%, and trpc7N = 5%.

View larger version (35K): [in this window] [in a new window]   FIG. 7. Effects of expression of truncated trpc channels in CHO-hOX1-C1 cells. Cells were transfected with truncated constructs or GFP, and Ca2+ imaging was performed. Before each recording of the fura-2 responses, cells that were fluorescent at 450–480 nm excitations were identified, and the cells were on this basis divided into fluorescent and nonfluorescent groups, respectively. A, a typical experiment with cells transfected with the trpc1N construct is shown. The cells were challenged with 0.3 and 3 nM Ox-A and 100 µM BzATP as indicated. B, data obtained from experimental conditions from A are summarized showing the effects of transfection with the different trpcN constructs. The data were processed for 0.3 nM Ox-A by normalization of the response of the individual cells to BzATP. The results are presented as the ratio between the responses in fluorescent and nonfluorescent cells, respectively, and expressed as % response ± S.E. Statistical significance was established by the one-way ANOVA test and followed by the Scheffe's test. **, p < 0.01. C, co-immunoprecipitation of trpc3FLAG/eyfp-trpc3N, trpc1FLAG/trpc1N-egfp, trpc3FLAG/trpc7N-egfp, trpc3FLAG/egfp-n3, and trpc7NFLAG/trpc7N-egfp transfected cells with polyclonal anti-GFP antibody cross-linked to protein G-magnetic beads (contr = untransfected cells) and detection with anti-FLAG M2 antibody (dilution 1:2000).

The co-precipitation data in Fig. 7C demonstrates that TRPC1N, TRPC3N, and TRPC7N are capable of binding their intact counterpart and furthermore that TRPC7N can bind full-length TRPC3, but EGFP alone could not precipitate any TRPC channels. TRPC3N, TRPC7N, and TRPC2N were not co-precipitated with TRPC1 (data not shown).

Effect of Channel Blockers on TRPC3 Channel Current— Several TRPC channels are constitutively active when overexpressed in commonly used cell lines, and this asset has been used previously to characterize their properties (21). CHO-hOX1-C1 cells were thus transfected with TRPC-cDNA constructs, and cells expressing constructs were identified by GFP fluorescence. Cells were clamped at –80 mV. In patched cells, no change in the basal membrane current was observed upon overexpression of TRPC1 (n = 6), which is consistent with previous findings (47). Because the primary current response to Ox-A was highly sensitive to TEA, Mg²⁺, and dextromethorphan, we tested their effect on cells expressing TRPC3FLAG. Fig. 8A (left panel) shows a representative current of a cell expressing TRPC3FLAG. Inward currents were in general transient and were followed by a more steady current level ranging from around –250 to –600 pA (48). Ramp analysis (Fig. 8A, right panel, indicated by numbers 1–3) shows that the voltage profiles, following whole cell access and during the transient and the more steady phase of the currents, are similar. The reversal potential was found to be 6.6 ± 0.5 mV (n = 17) ranging from around 4 to 8 mV. Thus, to determine their blocking effect, TEA, Mg²⁺, and dextromethorphan were applied during the more steady current level. Fig. 8B shows a representative recording in which TEA (70 mM) blocks the trpc3 current by 91%, Mg²⁺ (5 mM) by 47%, and dextromethorphan (100 µM) by 81%. TEA reduced the currents to less than 50 pA regardless of the magnitude of the basal steady current level. The magnitude of the block was in average found to be 85.6 ± 3.9% (n = 6) with respect to zero current. Dextromethorphan (100 µM) reduced the trpc3 current by 71.2 ± 4.5% (n = 6). Application of 5 mM Mg²⁺ was less effective and blocked the trpc3 current by 49.6 ± 2.0% (n = 6).

View larger version (18K): [in this window] [in a new window]   FIG. 8. Effects of tetraethylammonium, Mg2+, and dextromethorphan on membrane currents induced by TRPC3 overexpression. CHO-hOX1-C1 were transfected with trpc3cDNA inserted into pIRES-hrGFP-1a, which transcribes trpc3 and GFP as a single bi-cistronic mRNA (TRPC3-expressing cells are fluorescent). The membrane potential was clamped in whole-cell mode to –80 mV, and voltage ramps were introduced every 5 or 7.5 s (–80 to +80 mV; 320 ms). A Cs+-based intracellular solution was used. A, left panel shows a representative current response in a TRPC3-transfected cell. The current deflection during introduction of the ramps was removed from the trace for clarity. Scale bars indicate 200 pA and 100 s, respectively. Arrows with numbers 1–3 indicate the position of extracted current-voltage traces shown in A, right panel. Reversal potential and current-voltage profiles were similar in all traces, and thus blockers were introduced during the more steady state current. B shows a TRPC3 steady state current recorded under same conditions as in A. The presented current trace shows the effect of 70 mM TEA, 5 mM Mg2+, and 100 µM Dex in a HBM buffer including 1.2 mM Mg2+. The dotted line above current traces in A and B indicates zero current.

	DISCUSSION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

The results of the present study suggest that the primary pathway for OX₁ receptor-mediated Ca²⁺ elevation is activation of a nonstore-operated Ca²⁺-permeable channel. The molecular entity of this channel has, however, remained unresolved. A pharmacological distinction of nonstore-operated channels from store-operated mechanisms is difficult due to the lack of specific blockers or other specific means to distinguish the pathways. Thus, we screened a panel of channel inhibitors on the response to Ox-A and thapsigargin, in order to find compounds that would show preference for blocking a putative noncapacitative Ca²⁺ entry over typical capacitative entry. The Ca²⁺ response and inward currents evoked with 0.3 nM Ox-A were inhibited by Mg²⁺, dextromethorphan, and 70 mM TEA. Because Cs⁺ was used to substitute internal K⁺ in voltage clamp experiments, the action of TEA would not be expected to stem from K⁺ channel modulation but rather represents a direct channel block. This conclusion was further supported by our current clamp recordings that were unaffected by TEA. Sensitivity to Mg²⁺ has been demonstrated previously for certain TRP channels (38–40). We only observed the effects of extracellular Mg²⁺, on the Ox-A evoked current, when Mg²⁺ was above 1mM. The Mg²⁺ block by 5 mM left a residual Ox-A current that was not further blocked even by a higher Mg²⁺ concentration of 20 mM. Ramp analysis of Ox-A-stimulated membrane currents indicated that the voltage profile of the Ox-A-dependent current does not change in the presence of Mg²⁺, i.e. Mg²⁺ blocks in a voltage-independent manner.² The results presented here are strikingly similar to effects of Mg²⁺ reported by Hardie et al. (39) using Drosophila trp and TRPL cation channels. They showed a similar threshold at 1 mM Mg²⁺, similar concentration dependence, lack of voltage dependence, and a residual current in the presence of high Mg²⁺. They also demonstrated that the Mg²⁺ block was virtually voltage-independent. Like the Ox-A current, the constitutive current induced by TRPC3 overexpression was also sensitive to Mg²⁺, dextromethorphan, and TEA, which also argues for an action at the level of channels rather than blocking the receptor mechanisms. The apparent lack of competition with respect to dextromethorphan is in agreement with this.

On the other hand the peak and stable phase of Ca²⁺ elevation seen with high concentrations of Ox-A and the thapsigargin-induced Ca²⁺ entry phase were relatively insensitive to these inhibitors. In line with this, the store-operated pathway has also been shown previously (40) to be insensitive to Mg²⁺. Conversely, SKF96365inhibited the stable phase of the Ox-A response and the thapsigargin-mediated Ca²⁺ entry more effectively than the response to subnanomolar concentrations of Ox-A. Thus, the data discussed above strongly suggest that the response to subnanomolar concentrations of Ox-A is due to activation of a pathway for Ca²⁺ entry, which is distinct from the store-operated entry. This is in agreement with previous data showing robust Ca²⁺ elevation with subnanomolar concentrations of Ox-A with no appreciable emptying of Ca²⁺ stores (9). It has been shown previously with fura-2 in suspension recordings that the [Ca²⁺]_e-dependent response to orexins is inhibited by Ni²⁺ and is relatively insensitive to lanthanides or 2-APB, which in contrast strongly blocked the capacitative entry (10, 11). In line with this, voltage clamp recordings similarly showed that 0.3 nM Ox-A evoked an inward Ca²⁺-dependent current, which were completely insensitive to 2-APB, whereas Ni²⁺ almost completely blocked the current.

The involvement of a specific channel in the Ca²⁺ elevation by subnanomolar concentrations of Ox-A was further substantiated by the voltage dependence of the response. The Ox-A-activated current and membrane depolarization showed a steep dependence on a negative membrane potential. No appreciable current or depolarization was seen when the membrane potential was about –15 mV. In line with this, the Ca²⁺ elevation in intact cells showed a similar dependence on the membrane potential as depolarization with high K⁺ almost totally abolished the response to 0.3 nM Ox-A but did not affect the response at higher concentrations of Ox-A. In previous reports, orexins have similarly been shown to induce depolarization in native cells of comparable magnitude as observed in this study (3).

Activation of PKC by DOG, the diacylglycerol kinase inhibitor R59022 [GenBank] , or TPA caused a considerable reduction in the response to subnanomolar concentrations of Ox-A. In the same way as with some of the channel inhibitors Mg²⁺, dextromethorphan, and TEA (as discussed above), PKC activation did not significantly affect the peak or stable phase of the response to high concentrations of Ox-A or to thapsigargin. PKC activation has been shown previously to inhibit Ca²⁺ entry in response to receptor stimulation (49, 50). It has recently also been shown that certain isoforms of TRPC channels, most notably TRPC3, are blocked by activation of PKC (21–23). The Drosophila TRP channels have also been shown to be sensitive to PKC activation as judged from stimulatory effects of PKC inhibitors and inhibition by protein phosphatase inhibitor calyculin A (23). In agreement with previous studies (21) the store-operated pathway (stable phase of the response to high Ox-A concentrations and thapsigargin) was unaffected by PKC activation. An attractive hypothesis may be that PKC functions as a negative feedback to regulate these Ca²⁺-permeable channels and thus prevent massive intracellular Ca²⁺ elevation (21, 22). A negative feedback would explain the steep concentration dependence of the orexin-activated current response as compared with the far less steep concentration response curve for Ca²⁺ elevation. One possibility could be that the action of PKC is on the OX₁ receptor itself. However, this appears unlikely because the Ca²⁺ peak elevation with higher Ox-A concentration was relatively insensitive to PKC treatment. In the case of calyculin A, its action may be related to its ability to cause internalization of TRPC1, -3, and -4 channels (24).

A variety of nonstorage-activated Ca²⁺ channels are present in cells (12), and they are frequently observed when challenging cells with low agonist concentrations (51) as was also the case in this study. The characteristics of the pathway for Ca²⁺ entry described here including regulation by PKC, sensitivity to Mg²⁺, and inhibition by low intracellular Ca²⁺ are similar to those observed with expressed TRP channels (13). RT-PCR suggests the presence of four functional TRPC channels subtypes in our cells (trpc1–4). Trpc1 and -2 have been identified previously in CHO-K1 cells (52, 53). Comparison of the amount of PCR product obtained with each primer pair also indicates trpc1 and trpc2 to be the major trpcs in CHO-hOX1-C1 cells. This is in line with our results. We have additionally detected trp1 and -4 mRNA, which currently nobody to our knowledge has studied in CHO cells.

In the present study, expressing truncated trpc1 and trpc3 subtypes caused a clear inhibition of the response to subnanomolar concentrations of Ox-A. None of the constructs had any effect on the response to higher concentrations of Ox-A. These data suggest that TRPC1 and TRPC3 have a central role in the signaling via the OX₁ receptor. TRPC1 may also interact with other members of the TRP channel family like TRPC4 and TRPP channels (54). RT-PCR shows the presence of mRNA for polycystin2, mucolipin1, trpm2–7, and trpv1, -2, and -4 in CHO-hOX1-C1 cells.² The data in this study thus indicate that the Ox-A-activated pathway for Ca²⁺ entry involves TRPC1. Interestingly, a physical interaction of TRPC1 with the mGlur1 receptor has also been demonstrated (55). The stimulation of Ca²⁺ entry by DOG indicates that the cells express diacylglycerol-activated channels. Of the TRPC channel subtypes expressed in these cells, only TRPC3 has been shown to be activated by diacylglycerol (13). These data taken together with the similar sensitivity of currents activated by TRPC3 overexpression to the Mg²⁺, dextromethorphan, and TEA strongly suggest that TRPC3 channels are expressed in the membrane and are activated by orexin receptors. The sensitivity to protein kinase C activation (21–23) and calyculin A (24) is also a property typical of TRPC3 channels. As mentioned above TRPC3N expression also had a marked effect on the Ox-A response. We could not detect mRNA for the typical partners of TRPC3, namely TRPC6 and -7. Therefore, TRPC3 must be present as homomeric channels or then it interacts with as yet undefined partners. TRPC7N, which could interact with TRPC3, did also not affect the response. An explanation could be that the truncated channel subunits, even though they can bind the normal partners of their intact homolog, only have a dominant negative effect if they bind an intact endogenous homolog. Previous studies have also indicated that truncated channels may act by preventing insertion of native channel subunit into the membrane (45).

In some embryonic tissues, TRPC3 has been shown to be able to bind TRPC1 (27). A functional link between TRPC1 and TRPC3 is also suggested by findings demonstrating that these channel subtypes promote differentiation of hippocampal cells (56). Co-expression of TRPC1 and TRPC3 has further been shown to produce a novel membrane current indicating a functional interaction between these two channel subtypes (57). However, our co-immunoprecipitation data do not support a significant direct interaction. Because both TRPC1 and TRPC3 have been detected in caveolae and shown to bind caveolin-1, which is of importance for channel assembly (58, 59), the possibility also exist that TRPC1 and TRPC3 can collaborate without direct physical interaction. TRPC3 channels have in many studies been shown to be strongly stimulated by intracellular Ca²⁺ (13, 14, 22). Overexpression of TRPC3 produces constitutively active membrane currents in CHO-K1 cells (48). These currents are strongly regulated by Ca²⁺. An attractive hypothesis may thus be that TRPC1 and TRPC3 are activated by subnanomolar concentrations of Ox-A, which subsequently evoke a delayed Ca²⁺-dependent Ca²⁺ enhancement via subsequent TRPC3 channel stimulation. This hypothesis would also explain the delay in the response time and why the Ox-A-activated currents are abolished in a medium with strong intracellular Ca²⁺ buffer capacity. The lack of response to orexins in cells with high initial intracellular Ca²⁺, which also do not respond to Ox-A, would also be explained as the TRPC3 current would be already active/inactive.

Whether the results obtained here have relevance for the action of orexins in neurons is difficult to prove at present. As discussed above, the same G-protein mechanisms and TRPC channel subunits are functional in neuronal and non-neuronal cells. Thus one would expect that the basic signaling mechanisms are similar, although in neurons downstream pathways (e.g. different types of ion channels) may complicate the interpretation of data. Functional studies with recombinantly expressed orexin receptors in neuron-like cells (PC12 and Neuro2A) show the same basic features as those described in CHO-K1 cells (10). An inward Ca²⁺-dependent current and depolarization is also activated in PC12 cells.² In these cells like in neurons, however, several other mechanisms are additionally activated. Several mechanisms have been proposed for the actions of orexins in neurons, e.g. nonselective cation channels, Na⁺/Ca²⁺ exchange, and a reduction in potassium conductance or combinations of these (3, 4). The latter two mechanisms do not operate in CHO-K1 cells. It should be noted that the intracellular Ca²⁺ dependence as well as protein kinase regulation of the responses here may conceal signals in native cells. With neurons 1,000–10,000 higher, orexin concentrations have been used so the results may also not be directly comparable. Furthermore the orexin receptors are promiscuous and may interact with several different G-proteins (4). Therefore, the actions of orexins may be very dependent on the cellular microenvironment. The functions appearing at low concentrations of ligands are expected to be the primary responses of the receptor (33). Therefore, the mechanisms observed here are highly likely to be operating in neurons.

In conclusion, the data presented here show that the response to low concentrations of Ox-A acting at the OX₁ orexin receptor results in opening of a Ca²⁺-permeable channel distinct from the typical store-operated channels. Activation of this channel is sufficient to depolarize the cells by about 10 mV, so this mechanism may be of significance in excitatory cells as well. This pathway of Ca²⁺ entry can be distinguished from other pathways of Ca²⁺ mobilization on the basis of its sensitivity to inhibitors, interference with TRPC1 and TRPC3 channels, and its regulation by PKC.

	FOOTNOTES

 
^* This work was supported by European Union Contracts ERBBIO4CT960699 and QLG3-CT-2002-00826, the Academy of Finland, the Sigrid Jusélius Foundation, the Magnus Ehrnrooth Foundation, the Lars Hierta Foundation, the Göran Gustafsson Foundation, and the Novo Nordisk Foundation. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

The nucleotide sequence(s) reported in this paper has been submitted to the GenBank^TM/EBI Data Bank with accession number(s) AJ566614 [GenBank] , AJ566615 [GenBank] and AJ566613 [GenBank] .

^¶ To whom correspondence should be addressed: A. I. Virtanen Institute for Molecular Sciences, Dept. of Neurobiology, Laboratory of Cell Biology, University of Kuopio, P.O. Box 1627, FIN-70211 Kuopio, Finland. E-mail: karl.okerman{at}uku.fi.

¹ The abbreviations and trivial names used are: azido-GTPS, azidoguanosine 5'-3-O-(thio)triphosphate; PKC, protein kinase C; TRPC, transient receptor potential channel; 2-APB, 2-aminoethoxydiphenyl borate; GF109203X, bisindolylmaleimide; R59022 [GenBank] , diacylglycerol kinase inhibitor; SKF96365 1-[-(3-(4-methoxyphenyl)propoxy)-4-methoxyphenethyl]-1H-imidazole hydrochloride; BzATP, (2',3'-O-(4-benzoyl-benzoyl)-ATP); DOG, dioctanoyl glycerol; TPA, 12-O-tetradecanoylphorbol-13-acetate; probenecid, p-(dipropylsulfamoyl)benzoic acid; Ox-A, orexin-A; CHO-K1, Chinese hamster ovary cells; [Ca²⁺]_i, intracellular free Ca²⁺ concentration; ANOVA, analysis of variance; GFP, green fluorescent protein; EGFP, enhanced GFP; TEA, tetraethylammonium; Dex, dextromethorphan; RT, reverse transcription.

² K. P. Larsson, H. M. Peltonen, G. Bart, L. M. Louhivuori, A. Penttonen, M. Antikainen, J. P. Kukkonen, and K. E. O. Åkerman, unpublished observations.

	ACKNOWLEDGMENTS

 
We are grateful for the laboratory assistance provided by Veera Pevgonen (A. I. Virtanen Institute). The generous material and scientific support from Dr. Michel Detheux (Euroscreen) is also gratefully acknowledged.

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TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 

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