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Article |
IP3 sensitizes TRPV4 channel to the mechano- and osmotransducing messenger 5'-6'-epoxyeicosatrienoic acid
Correspondence to M. Valverde: miguel.valverde{at}upf.edu
Mechanical and osmotic sensitivity of the transient receptor potential vanilloid 4 (TRPV4) channel depends on phospholipase A2 (PLA2) activation and the subsequent production of the arachidonic acid metabolites, epoxyeicosatrienoic acid (EET). We show that both high viscous loading and hypotonicity stimuli in native ciliated epithelial cells use PLA2–EET as the primary pathway to activate TRPV4. Under conditions of low PLA2 activation, both also use extracellular ATP-mediated activation of phospholipase C (PLC)–inositol trisphosphate (IP3) signaling to support TRPV4 gating. IP3, without being an agonist itself, sensitizes TRPV4 to EET in epithelial ciliated cells and cells heterologously expressing TRPV4, an effect inhibited by the IP3 receptor antagonist xestospongin C. Coimmunoprecipitation assays indicated a physical interaction between TRPV4 and IP3 receptor 3. Collectively, our study suggests a functional coupling between plasma membrane TRPV4 channels and intracellular store Ca2+ channels required to initiate and maintain the oscillatory Ca2+ signal triggered by high viscosity and hypotonic stimuli that do not reach a threshold level of PLA2 activation.
Abbreviations used in this paper: 4
-PDD, 4-phorbol 12,13-didecanoate; AA, arachidonic acid; AACOCF3, arachidonyl trifluoromethyl ketone; ANOVA, analysis of variance; CBF, ciliary beat frequency; EET, epoxyeicosatrienoic acid; IP3, inositol trisphosphate; IP3R, IP3 receptor; pBPB, 4-bromophenacyl bromide; PLA2, phospholipase A2; TRP, transient receptor potential; TRPV4, TRP vanilloid 4.
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The nonselective cation channel TRPV4 is a member of the vanilloid subfamily of transient receptor potential (TRP) channels (Montell, 2005). TRPV4 shows multiple modes of activation and regulatory sites, enabling it to respond to various stimuli, including osmotic cell swelling (Strotmann et al., 2000; Liedtke et al., 2000; Wissenbach et al., 2000; Arniges et al., 2004), mechanical stress (Gao et al., 2003; Suzuki et al., 2003; Liedtke et al., 2003; Andrade et al., 2005), heat (Guler et al., 2002), acidic pH (Suzuki et al., 2003), endogenous ligands (Watanabe et al., 2003), and both PKC-activating and nonactivating phorbol esters (Watanabe et al., 2002a; Xu et al., 2003). Besides, TRPV4 can be sensitized by coapplication of different stimuli (Gao et al., 2003; Alessandri-Haber et al., 2006; Grant et al., 2007). Osmotic (Vriens et al., 2004) and mechanical (Andrade et al., 2005) sensitivity of TRPV4 depends on phospholipase A2 (PLA2) activation and the subsequent production of the arachidonic acid (AA) metabolites, epoxyeicosatrienoic acids (EETs). Signaling pathways involving G proteins and/or PLC/IP3 are also activated by osmotic cell swelling (Suzuki et al., 1990; Hoffmann and Dunham, 1995; Felix et al., 1996) and mechanical stimulation (Vandenburgh et al., 1993; Felix et al., 1996; Gudi et al., 1998). However, the contribution of these pathways to the generation of an osmotic or mechanically induced Ca2+ signal by TRPV4 is unknown. Given that both extracellular ATP (Evans and Sanderson, 1999; Morales et al., 2000) and intracellular PLA2 and PLC pathways (Hermoso et al., 2001; Barrera et al., 2004; Andrade et al., 2005) are involved in the regulation of CBF, we explored whether the PLC–IP3 pathway might be involved in the response of the TRPV4 channel to high viscous solutions and hypotonic cell swelling. To do so, we measured TRPV4 activity in both hamster oviductal ciliated cells and TRPV4-expressing HeLa cells. We show here for the first time that IP3, without being an agonist itself, sensitizes TRPV4 to EET but not to other TRPV4 physiological stimuli such as warm temperature, an effect that requires a functional IP3 receptor (IP3R).
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-tubulin (Fig. 1 B, bottom right, yellow), although the strongest TRPV4 (green) and -tubulin (red) signals were at the base (apical side of the cell) and the tip of cilia, respectively. A weak intracellular and basolateral membrane TRPV4 signal was also present in all cells analyzed. In controls where the primary antibody was omitted or in the presence of antigen-reabsorbed TRPV4 antibody, no signal was observed either at the cilia or apical location (Fig. S1, available at http://www.jcb.org/cgi/content/full/jcb.200712058/DC1).
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Under conditions favorable to measuring inward cationic currents (see Materials and methods), 20% dextran (Fig. 1 C) and 30% hypotonic (70% of normal osmolality) solutions (Fig. 1 D) evoked whole-cell TRPV4-like currents in isolated actively beating hamster oviduct cells. Inhibition of PLA2 with 100 µM 4-bromophenacyl bromide (pBPB) or PLC with 1 µM U73122 totally blocked high viscosity–induced (20% dextran) TRPV4 current activation (Fig. 1 C), whereas hypotonicity (30%)-activated TRPV4 currents were completely inhibited only by pBPB (Fig. 1 D). Inhibition of PLA2 with arachidonyl trifluoromethyl ketone (AACOCF3; 50 µM) also blocked dextran-induced TRPV4 currents (20% dextran: –14.8 ± 0.8 pA/pF, n = 7; vs. dextran + AACOCF3: –1.9 ± 0.6 pA/pF, n = 5; P < 0.05). In the presence of U73122, significant hypotonicity-activated TRPV4 currents were recorded, although of smaller magnitude (Fig. 1 D). Mean normalized current responses obtained in all conditions described in Fig. 1 (C and D) are shown in Fig. 1 (E and F). The dramatic impact of the PLC inhibitor upon TRPV4 response to dextran-containing solutions contrasted with its modest effect upon hypotonic stimulation. This observation prompted us to analyze the signaling pathways up- and downstream of PLC activation. The participation of intracellular Ca2+ stores in the activation of TRPV4 channel by 20% dextran solutions was discarded, as 1 µM thapsigargin, a blocker of the ER calcium pump, did not modify the TRPV4 response (Fig. 2 A). Transient cationic currents were observed after thapsigargin addition (not depicted) but disappeared within 5 min, the time at which cells were exposed to dextran solutions in the presence of thapsigargin (Fig. 2 A). High viscosity–induced currents were prevented in cells loaded with 500 µM GDPβ-s, which locks G proteins in their inactive state (Fig. 2 B), or treated with 100 µM of the P2 receptor antagonist suramin (Fig. 2 C). Mean normalized current responses obtained in the conditions described in Fig. 2 (A–C) are shown in Fig. 2 D. GDPβ-s also reduced the hypotonicity-induced currents to the same levels recorded in the presence of U73122 (Fig. S2 A, available at http://www.jcb.org/cgi/content/full/jcb.200712058/DC1) but significantly different from those obtained in the presence of GDPβ-s and dextran solutions (P < 0.05; Fig. 2 D), confirming the differences in sensitivity of high viscosity– and hypotonicity-induced TRPV4 response to the G protein–PLC pathway. Dialysis of cells with a pipette solution containing 30 µM IP3 restored TRPV4 activation by dextran solutions in the presence of U73122 (PLC inhibitor) except when both U73122 and pBPB (PLA2 inhibitor) were used (Fig. 2, E and F). Similarly, the presence of IP3 in the pipette solution also restored full TRPV4 activation by 30% hypotonic solutions (Fig. S2 A). The combination of pipette solutions containing IP3 and cell stimulation with 20% dextran solutions showed a modest but significant potentiation of the response, which was not modified by treatment with thapsigargin (Fig. S2 B), therefore confirming the little impact of intracellular stores to the IP3-mediated sensitization of TRPV4 response to highly viscous loads.
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We first tested whether the level of PLA2 activation is responsible for the different response seen under high viscous and hypotonic solutions. In the absence of a reliable method to directly test PLA2 activity in hamster oviductal ciliated cells, we measured the Ca2+ signal and its dependence on PLC–IP3 in ciliated cells under milder hypotonic stimuli (15%), aiming to elicit less PLA2 activation. Fig. 5 (A and B) shows that, unlike the Ca2+ response to 30% hypotonic solutions (Fig. 3 F), the response to 15% hypotonicity is completely abolished by U73122. These results suggest that PLC–IP3 pathway also becomes crucial to the generation of the Ca2+ signal under conditions of lower PLA2 activation by milder hypotonic stimuli.
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IP3R mediates IP3-dependent sensitization of TRPV4 to 5',6'-EET
Next, we further investigated the mechanism of IP3-mediated sensitization. A previous study has localized IP3R3 (not IP3R1) to the plasma membrane of hamster oviduct ciliated cells and suggested its participation in Ca2+ influx (Barrera et al., 2004). To test whether IP3-mediated sensitization was a general mechanism rather than circumscribed to ciliated cells, we expressed human TRPV4 in HeLa cells that endogenously expressed IP3R1 and IP3R3 (Tovey et al., 2001).
Dialysis, through the patch pipette, of HeLa cells expressing human TRPV4 with 1 nM 5',6'-EET and 30 µM IP3 resulted in an increase in current that reached a maximum within 3–5 min (Fig. 6 A). The current-voltage relationship of the corresponding TRPV4 currents is shown in Fig. 6 D. Dialysis with EET and/or IP3 alone elicited no significant TRPV4-like currents (Fig. 6 D). The cationic currents shown in Fig. 6 (A and D) illustrate that the IP3-mediated sensitization observed in hamster ciliated cells is reconstituted in HeLa cells expressing human TRPV4. Mean normalized currents are shown in Fig. 6 H. Increasing 5',6'-EET concentration to 100 nM in the presence of 30 µM IP3 augmented TRPV4 current amplitude (Fig. S4, available at http://www.jcb.org/cgi/content/full/jcb.200712058/DC1), although it did not reach statistical significance compared with 1 nM 5',6'-EET + 30 µM IP3 (Fig. 6 H) or 100 nM 5',6'-EET alone (Fig. S4). No sensitization was observed in HeLa cells transfected with rat IP3R3 (Fig. 6, C and F) or EGFP alone (Fig. 6 G). Coexpression of TRPV4 and IP3R3 (Fig. 6, B, E, and I) tripled IP3 potentiation as compared with TRPV4 expression alone, an effect that was completely inhibited in the presence of 1 µM of the IP3R inhibitor xestospongin C. Neither EET (1 nM) nor IP3 alone activated TRPV4 currents in HeLa cells expressing any combination of EGFP, TRPV4, or IP3R3 (P > 0.05).
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-phorbol 12,13-didecanoate (4-PDD; 10 µM). Similar results were obtained in CHO, HEK, and COS cells transfected with human TRPV4 (n > 200; unpublished data). HeLa cells expressing IP3R3 showed no response to dextran solutions containing 1 µM ATP (Fig. 7 G). Mean increases in the Ca2+ signal obtained in the conditions described in Fig. 7 (A–G) are shown in Fig. 7 (H–J). Altogether, these data suggest that ATP–PLC–IP3 signaling sensitizes TRPV4 response to high-viscosity and hypotonic solutions (calcium imaging data) or to the mechano- and osmotransducing TRPV4-activating messenger 5',6'-EET (patch-clamp data), an effect that depends on the presence of a functional IP3R.
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Different pieces of evidence have pointed to TRPV4 as the Ca2+ entry channel in response to high viscous and hypotonic solutions in native hamster oviductal ciliated epithelial cells (Andrade et al., 2005; Teilmann et al., 2005): (1) TRPV4 mRNA and protein have been identified in oviductal ciliated cells; (2) electrophysiological characterization of high-viscosity– and hypotonicicty-induced cationic currents in ciliated cells coincides with the features of cationic currents induced by the TRPV4-specific agonist 4-PDD; (3) functional inhibition of the high viscosity–induced cationic current with an antibody against TRPV4 in oviductal ciliated cells; and (4) high-viscosity solutions evoked cation currents and Ca2+ signals in TRPV4-expressing HeLa cells but not in cells transfected with vector alone. The present study adds new evidence supporting the participation of TRPV4 in the response of ciliated cells to high-viscocity and hypotonic solutions: the presence of TRPV4 at the base of the cilia, where oscillatory Ca2+ signals are needed to modulate CBF (Tamm, 1994; Evans and Sanderson, 1999; Lansley and Sanderson, 1999), and the convergence of signaling pathways in the TRPV4 activation by high viscosity and hypotonic solutions in both native ciliated cells and in cells heterologously expressing the channel. All together, these observations are consistent with the role of TRPV4 in the transduction of mechanical stimulation in ciliated epithelial cells (Andrade et al., 2005). Ciliated epithelia of the oviduct also express TRPP1-2 (Teilmann et al., 2005), although their functional significance is still unresolved.
We now demonstrate for the first time that: (1) PLC–IP3 signaling participates in TRPV4 activation by high-viscosity solutions in hamster oviductal ciliated cells downstream to the activation of P2 receptors after mechanically induced ATP release (Felix et al., 1996; Okada et al., 2006; Winters et al., 2007); under our experimental conditions, the suramin-sensitive receptor implicated is most likely of the P2Y2 type, which has been associated to the mechanosensitivity of ciliated epithelial cells (Winters et al., 2007) and is present in hamster oviductal ciliated cells (Morales et al., 2000); (2) that IP3 alone is able to compensate the inhibitory effect of U73122; (3) that the effect of IP3 requires a functional IP3R (as the sensitizing effect is inhibited by xestospongin C) although it does not require the release of Ca2+ via IP3R, as the response is maintained in cells in which ER was calcium-depleted using thapsigargin; and (4) the possibility that IP3-mediated potentiation of TRPV4 response to dextran solutions involves positive feedback via a Ca2+-calmodulin–dependent mechanism (Strotmann et al., 2003) is unlikely, as TRPV4 currents were recorded in the absence of extracellular and intracellular Ca2+ (including 5 mM EGTA) and in the presence of thapsigargin.
Both hypotonic and mechanical stimulation activates PLC and/or PLA2 in different cell types (Lehtonen and Kinnunen, 1995; Pedersen et al., 2000; Moore et al., 2002; Zholos et al., 2005), although, to date, only the latter has been implicated in TRPV4 regulation. Activation of both signaling pathways has been associated with direct sensing by the phospholipid bilayer of physical stimuli and activation of membrane-bound G proteins in the case of PLC (Gudi et al., 1998, 2003) or direct activation of PLA2 (Lehtonen and Kinnunen, 1995; Pedersen et al., 2000). Moreover, crosstalk between PLC and PLA2 has been demonstrated in several cell types (Vandenburgh et al., 1993). Activation of TRPV4 under hypotonic (Vriens et al., 2004) and high viscosity conditions (Andrade et al., 2005) depends on the activity of PLA2 and appears to be ultimately related to the production of 5',6'-EET via the metabolism of AA by P450 enzymes. Thus, 5',6'-EET is the only physiological, diffusible molecule known to directly activate TRPV4 (Watanabe et al., 2003). Other TRPV4 stimuli such as temperature and the synthetic 4-PDD are independent of 5',6'-EET production (Vriens et al., 2004).
The impact of the ATP–PLC–IP3 pathway on TRPV4 activity depends on the stimuli used, being more relevant in the case of channel activation by 20% dextran solutions than in the case of 30% hypotonic solutions, probably reflecting a higher level of PLA2 activity in the latter. However, using a milder hypotonic stimuli (15%) turned the response fully PLC dependent. IP3 also potentiated TRPV4 response to low EET concentrations measured by whole-cell patch clamp of both native ciliated epithelia and cells heterologously expressing TRPV4, which is consistent with the observation that convergence of ATP–PLC–IP3 and PLA2–AA–EET signaling is essential for the activation of TRPV4 by high viscous and hypotonic solutions that do not reach a threshold level of PLA2 activation.
Our data also addresses the impact of Ca2+ entry upon agonist-induced Ca2+ oscillations (Yule and Gallacher, 1988; Shuttleworth, 1999). The dependency on Ca2+ entry for continued oscillations has been interpreted in terms of the Ca2+ dependency of the IP3R (Shuttleworth, 1999). Under conditions of low activation of the PLC–IP3 pathway (usually associated to oscillatory Ca2+ signals), IP3 will bind to IP3R and release little or no stored Ca2+, a response that is magnified by the sensitizing effect of Ca2+ entry via plasma membrane channels situated in close proximity to the IP3R. Using this model, we propose that the PLC–IP3 pathway is required for PLA2-dependent TRPV4 activation by dextran solutions and that both active TRPV4 and the PLC–IP3 pathway are needed to maintain the oscillatory Ca2+ signal. In the case of 30% hypotonic stimuli, TRPV4 activation is largely independent of PLC–IP3 pathway but, again, both active TRPV4 and the PLC–IP3 pathway are needed to maintain oscillations. In this sense, it is worth mentioning that, although the basic features of the TRPV4 response in native epithelia are reproduced in cells heterologously expressing the channel, the overall Ca2+ signal recorded in response to 20% dextran and 30% hypotonic solutions was not fully reproduced in the cell expression systems. Ciliated epithelial cells responded with oscillatory Ca2+ signals to both stimuli (Fig. 3), whereas HeLa cells expressing TRPV4 responded with single, transient Ca2+ increases (Fig. 7). Occasionally, additional peaks were observed in HeLa cells (Fig. 7 B). Another difference between the response of ciliated epithelia and HeLa cells expressing TRPV4 is the impact of the ATP–PLC–IP3 pathway on TRPV4 activation by 20% dextran solutions. Although ciliated cells responded to dextran solutions in the absence of added ATP, HeLa cells required the presence of 1 µM ATP to respond to dextran solutions. The difference may reflect a higher efficiency of ciliated cells to release ATP in response to mechanical/osmotic stimuli, a higher sensitivity of the ATP–PLC–IP3 pathway to extracellular ATP, or more efficient coupling between the ATP–PLC–IP3 and PLA2–AA–EET pathways to activate TRPV4. At present, we cannot discriminate between these three possibilities, and this remains an interesting issue for future studies.
Conclusions
We have delineated a novel regulatory mechanism through which IP3, via its receptor, potentiates TRPV4 sensitivity to the mechano- and osmotransducing messenger 5',6'-EET but not to thermal stimulation. However, at present it is not known whether the association between TRPV4 and IP3R demonstrated by the coimmunoprecipitation studies is required to support the potentiating effect of IP3. IP3Rs are, themselves, capable of mediating plasma membrane Ca2+ entry (Dellis et al., 2006) or interacting with and modulating the TRPC and TRPP channels (Boulay et al., 1999; Kiselyov et al., 2005; Li et al., 2005), although no description of such mechanisms exists for the subfamily of TRPV channels (Clapham, 2003). Thus, IP3R, without being a channel itself or a being direct activator of plasma membrane ion channels, modulates Ca2+ influx via TRPV4. This mechanism is another example of the complexity of TRP channel gating, most likely reflecting the physiologically relevant convergence of different signaling pathways into channel gating. In conclusion, the functional coupling between IP3R and TRPV4 ensures channel gating under conditions of mechanical stimulation that do not reach a threshold level of PLA2–EET pathway activation.
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Cells
Primary cultures and single ciliated cells were obtained and maintained as described previously (Lock and Valverde, 2000; Hermoso et al., 2001; Andrade et al., 2005). Animals were maintained and experiments were performed according to the guidelines issued by the Institutional Ethics Committees of the Institut Municipal d'Investigació Mèdica of the Universitat Pompeu Fabra. All experiments were performed only in beating ciliated cells. HeLa cells were transfected with different combinations of the following cDNAs as described previously (Arniges et al., 2006): 0.3 µg pEGFPN1, 1.5–3 µg pcDNA-3-TRPV4-human, and 1.5–3 µg rat pcDNA-3-rat IP3R3s (provided by H. DeSmedt, Katholieke Universiteit Leuven, Leuven, Belgium; and G.I. Bell, University of Chicago, Chicago, IL).
Laser confocal immunodetection
Hamster oviducts were fixed overnight at 4°C with 4% paraformaldehyde + 3.7% sucrose, embedded into 7.5% gelatin, and sectioned with a criostat (
8 µm). Isolated cells were fixed in suspension with 4% (wt/vol) paraformaldehyde + 3.7% sucrose in PBS for 30 min at 4°C and attached to 1.5% gelatin-coated coverslips by spinning at 500 rpm for 3 min using a cytospin (Shandon; Thermo Fisher Scientific). After the spinning, fixation procedure continued for an additional 10 min at RT. Tissue sections and single cells were permeabilized with Tween 20 (0.1% or 0.05%, respectively) in PBS (1 h or 15 min at RT) and nonspecific interactions were blocked with 1.5% BSA + 5% FBS + Tween 20 (0.1% or 0.05%) in PBS. Sections and isolated cells were incubated overnight at 4°C with the primary antibodies diluted in the same blocking solution. For competition experiments, the antigen was incubated with TRPV4 antibody (5:1) for 1 h at RT.
The anti-TRPV4 polyclonal antibody (Arniges et al., 2004, 2006; Andrade et al., 2005) was used at 6.4 µg/ml. A commercial anti–-tubulin (Sigma-Aldrich) was diluted to 1:500. For immunodetection, we used goat anti–rabbit IgG Alexa 488 (Invitrogen) and goat anti–mouse IgG Alexa 555 (Invitrogen) diluted 1:750 in the same solution used with the primary antibodies. Samples were counterstained with 1 µg/ml TO-PRO-3 in PBS for nuclear localization. Images were taken at RT with an inverted confocal microscope (SP2; Leica) using an HCX Pl APO 63x 1.32 NA oil Ph3 confocal scanning objective (Leica), LCS confocal software (Leica) and argon (488 nm; JDS Uniphase Corporation) and HeNe (555 and 633 nm; JDS Uniphase Corporation and LASOS Lasertechnik GmbH, respectively) lasers. Original images were not further processed except for adjustments of brightness, contrast, and color balance.
Measurement of intracellular Ca2+
Cytosolic Ca2+ signal was determined at 30–37°C in cells loaded with 4.5 µM fura-2AM as described previously (Fernandez-Fernandez et al., 2002; Arniges et al., 2004, 2006). Cytosolic Ca2+ increases are presented as the increment in the ratio of emitted fluorescence (510 nm) after excitation at 340 and 380 nm relative to the baseline.
Electrophysiology
Ionic currents were recorded in the whole-cell patch-clamp mode (Fernandez-Fernandez et al., 2002). Patch pipettes were filled with a solution containing 140 mM N-methyl-D-glucamine chloride, 1 mM MgCl2, 5 mM EGTA, 10 mM Hepes, 4 mM ATP, and 0.1 mM GTP (300 mosmoles/liter, pH 7.3). Occasionally, pipette solutions contained different concentrations of 5',6'-EET and IP3 (as shown in the corresponding figures). Cells were held at 0 mV and ramps from –140 mV to +100 mV (400 ms) were applied at a frequency of 0.2 Hz. Ramp data were acquired at 10 KHz and low-pass filtered at 1 KHz. Experiments were performed at RT (22–26°C). In case of TRPV4 activation by heat, temperature of the bathing solution was changed within 30 s.
Immunoprecipitation assay, SDS-PAGE, and Western blotting
TRPV4-YFP and IP3R3 receptor transfected and nontransfected HeLa cells (48 h) were washed twice with cold PBS and detached using a cell scraper. Whole cell extracts were obtained by resuspension in BNP40 lysis buffer (1% Nonidet P-40, 10% glycerol, 150 mM NaCl, 5 mM EDTA, 1 mM Na3VO4, 1 mM DTT, 1 mM PMSF, 0.05% aprotinin, pH 7.4, and 50 mM Tris-Cl, pH 7.4) containing 12% of additional protease inhibitor cocktail (Roche). Cell extracts were agitated for 1 h at 4°C, aspirated three times through a 25-gauge needle, and centrifuged at 13,000 rpm for 1 h at 4°C. Protein concentration was quantified by the Bradford method. 620 µg of total protein samples from transfected and nontransfected HeLa cells were incubated overnight at 4°C, gently mixed with 4 µl of anti-rGFP polyclonal antibody (rabbit; Clontech Laboratories, Inc.) or 2,700 µg of total protein and 27 µl of anti-IP3R3 antibody (mouse; BD Biosciences) for the reverse coimmunoprecipitation. After that, 15 µg of protein G was added to the samples and mixtures were incubated for 2 h at room temperature. Protein G immunocomplexes were collected by centrifugation, washed four times with PBS, and resuspended in Laemmli sample buffer with 5% β-mercaptoethanol. Samples were boiled for 6 min at 100°C and centrifuged for 10 min at 13,000 rpm to remove protein G. Supernatants were collected, boiled again for 3 min at 100°C, electrophoresed in 8% Tris-HCl polyacrylamide gels, and transferred to nitrocellulose membranes using a dry blotting system (iBlot; Invitrogen). Membranes were blocked overnight at 4°C in Tris Base solution 1x–0.1% Tween 20 containing either 5% skim milk or 3% BSA. Membranes were washed again and subjected to chemiluminescence analysis using SuperSignal West Pico chemiluminescent substrate (Thermo Fisher Scientific) and detected on ECL films (GE Healthcare). Primary antibodies used for Western blotting were anti-rGFP for TRPV4 detection (1:1,000, rabbit; Clontech Laboratories, Inc.), anti-IP3R3 (1:3,000, mouse; BD Biosciences), and anti-IP3R1 (1:2,000, rabbit; Millipore). Mouse and rabbit secondary antibodies (GE Healthcare) from a sheep and donkey source, respectively, were used at 1:2,000.
Statistics
Data are expressed as the mean ± SEM. Student's t test or analysis of variance (ANOVA) were performed with the SigmaPlot (Systat Software, Inc.) and SPSS (SPSS, Inc.) programs. Bonferroni's test was used for post hoc comparison of means.
Online supplemental material
Fig. S1 shows immunofluorescence images of an antigen-preabsorbed TRPV4 antibody. Fig. S2 shows the effect of GDPβ-s, U73122, IP3, and thapsigargin on cationic currents recorded from hamster oviductal ciliated cells stimulated with 20% dextran or 30% hypotonic solutions. Fig. S3 shows calcium imaging data obtained from hamster oviductal ciliated cells in response to 20% dextran or 30% hypotonic solutions in the presence of U73343 or thapsigargin. Fig. S4 shows potentiation of the TRPV4 current by IP3 at different EET concentrations in TRPV4-expressing HeLa cells. Fig. S5 shows reverse coimmunoprecipitation. Online supplemental material is available at http://www.jcb.org/cgi/content/full/jcb.200712058/DC1.
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Acknowledgments |
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This work was supported by grants from the Spanish Ministries of Education and Science (SAF2006-04973 and SAF2006-13893-C02-02), and Health (Fondo de Investigación Sanitaria, Red HERACLES RD06/0009), the Generalitat de Catalunya (SGR05-266), and Fundació la Marató de TV3 (061331). J.M. Fernández-Fernández is a Ramón y Cajal Fellow. The authors declare that they have no competing financial interests.
Submitted: 11 December 2007
Accepted: 14 March 2008
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ratio, 340/380) obtained from different primary cultures of hamster oviductal ciliated cells stimulated with: (A) 20% dextran solution (n = 21; inset shows a recording obtained from a single cell); (B) dextran + 2 µM U73122 (n = 22); and (C) dextran + apyrase (10 U/ml; n = 19). (D) Mean [Ca2+] increases (
6 for each concentration). (D) Potentiation of the TRPV4 response in ciliated cells loaded with 1 nM 5',6'-EET and 30 µM IP3. Data are expressed as the mean ± SEM; *, P < 0.001 for EET + IP3 versus EET or IP3 alone (one way ANOVA and Bonferroni post hoc).
) in HeLa cells transfected with human TRPV4 (A), TRPV4 and rat IP3R3 (B), or IP3R3 (C) and dialyzed with 1 nM 5',6'-EET and 30 µM IP3. At the time indicated by the bars, the NaCl bathing solution was replaced by N-methyl-D-glucamine chloride solution (NMDGCl) in A and B. Current-voltage relations of peak whole-cell cationic currents were recorded from different HeLa cells transfected with TRPV4 (D) or TRPV4 and IP3R3 (E) and dialyzed with either 1 nM 5',6'-EET, 30 µM IP3, or EET+IP3. (F) Current densities at –100 mV in HeLa cells transfected with IP3R3 and dialyzed with 1 nM 5',6'-EET (n = 4), 30 µM IP3 (n = 5), and EET+IP3 (n = 4). (G) EGFP transfected cells dilayzed with 1 nM 5',6'-EET (n = 3), 30 µM IP3 (n = 4), and EET+IP3 (n = 4). (H) Current densities at –100 mV in HeLa cells expressing TRPV4: control (n = 4); 1 nM 5',6'-EET (n = 7); 30 µM IP3 (n = 8); and EET + IP3 (n = 8). (I) Current densities at –100 mV in HeLa cells expressing TRPV4 and IP3R3: control (n = 6); 1 nM 5',6'-EET (n = 10); 30 µM IP3 (n = 11); EET + IP3 (n = 8); and EET + IP3 + 1 µM xestospongin C (n = 4). Data are expressed as the mean ± SEM; **, P < 0.001 (one-way ANOVA and Bonferroni post hoc).
