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PIP2-DEPENDENT REARRANGEMENT OF TRPV4 CYTOSOLIC TAILS ENABLES CHANNEL ACTIVATION BY PHYSIOLOGICAL STIMULI
Anna Garcia-Eliasa,1 , Sanela Mrkonjića,1 , Carlos Pardo-Pastora , Hitoshi Inadab , Ute A. Hellmichb , Fanny Rubio-Moscardóa , Cristina Plataa , Rachelle Gaudetb , Rubén Vicentea and Miguel A. Valverdea,2
a

Laboratory of Molecular Physiology and Channelopathies,Dept. of Experimental and Health Sciences, Universitat Pompeu Fabra, Barcelona, Spain; b Dept. of Molecular and Cellular Biology, Harvard University, Cambridge, USA. Submitted to Proceedings of the National Academy of Sciences of the United States of America

Most TRP channels are regulated by phosphatidylinositol-4,5biphosphate (PIP2 ), although the structural rearrangements occurring upon PIP2 binding are currently far from being understood. Here we report that TRPV4 activation by hypotonic and heat stimuli requires PIP2 binding to and rearranging of the cytosolic tails. Neutralization of the positive charges within the sequence 121 KRWRK125 , which resembles a phosphoinositide binding site, rendered the channel unresponsive to hypotonicity and heat but responsive to 4α-phorbol 12,13-didecanoate, an agonist that binds directly to transmembrane domains. Similar channel response was obtained by depleting PIP2 from the plasma membrane with translocatable phosphatases in heterologous expression systems or by activation of phospholipase C in native ciliated epithelial cells. PIP2 facilitated TRPV4 activation by the osmotransducing cytosolic messenger 5’-6’-epoxyeicosatrienoic acid and allowed channel activation by heat in inside-out patches. Protease protection assays demonstrated a PIP2 binding site within the N-tail. The proximity of TRPV4 tails, analysed by fluorescence resonance energy transfer, increased by depleting PIP2 , mutations in the PIsite or by co-expression with PACSIN3, a regulatory molecule that binds TRPV4 N-tails and abrogates activation by cell swelling and heat. PACSIN3 lacking the F-BAR domain interacted with TRPV4 without affecting channel activation or tail rearrangement. Therefore, mutations weakening the TRPV4-PIP2 interacting site and conditions that deplete PIP2 or restrict TRPV4 access to PIP2 –in the case of PACSIN3- change tail conformation and negatively affect channel activation by hypotonicity and heat.
TRP | conformational change | heat | osmotic

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INTRODUCTION TRPV4 is a non-selective cation channel that responds to osmotic (1-4), mechanical (5-7) and temperature stimulation (8), thereby contributing to many different physiological functions: cellular (4, 9) and systemic volume homeostasis (10), vasodilation (11, 12), nociception (13), epithelial hydroelectrolyte transport (14), bladder voiding (15), ciliary beat frequency regulation (7, 16, 17), chondroprotection (18) and skeletal regulation (19). Osmotic (20) and mechanical (7, 16) sensitivity of TRPV4 depends on phospholipase A2 activation and the subsequent production of the arachidonic acid metabolites, epoxyeicosatrienoic acids (EET), while the mechanism leading to temperature-mediated activation (only observed in intact cells) it is not known at present (21). Reports also exists claiming EET-independent TRPV4 activation by membrane stretch in excised-patches from oocytes (22), in apparent contradiction with early reports claiming lack of activation by membrane stretch (1). Several studies have characterized TRPV4 domains implicated in channel regulation by calmodulin (23, 24), PACSIN3 (25), intracellular ATP (24) and inositol-trisphosphate receptor (16, 26). However, little is known about the domains relevant for TRPV4 activation by different stimuli, apart from the interaction between the TRPV4 activator 4α-phorbol 12,13-didecanoate (4α−PDD) and transmembrane
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domains 3 and 4 (27). Analysis of disease-causing mutations modifying channel activity that lay in regions close to the channel pore or within the ankyrin repeats (28) has also contributed to our understanding of relevant protein domains. Most TRP channels are regulated by phosphatidylinositol phosphates, particularly by phosphatidylinositol-4,5-biphosphate (PIP2 ), which is the most abundant phosphoinositide in the inner leaflet of the plasma membrane (29, 30). In general terms, it is proposed that PIP2 modulates TRP channel gating and/or the sensitivity to activating stimuli (29, 30). The interaction of PIP2 with TRPs involves protein regions characterised by the presence of several positively charged residues. Mutations of these positive residues (31-33) and manipulation of the PIP2 levels in intact cells (34) or in excised patches (33) have been the main tools to evaluate PIP2 -mediated channel regulation. The recent report of the crystal structure of K+ channels with bound PIP2 provides the first atomistic description of a molecular mechanism by which PIP2 regulates channel activity (35, 36). PIP2 binding induces a large conformation change in the protein, expanding and bringing the cytosolic domains closer to the transmembrane domains (35). Whether PIP2 modulation of TRP channels involves similar conformational changes is still an open question. We now show that TRPV4 requires the interaction of PIP2 with a stretch of positive charges at the N-tail, prior to the prolinerich domain (PRD, residues 132-144), in order to be activated by hypotonicity and heat. Moreover, we have also demonstrated that the reported lack of channel response to heat in excised patches is fully recovered in the presence of PIP2 , thereby suggesting that TRPV4 is bona fide thermosensitive channel. Finally, reduction of PIP2 levels or disruption of the PIP2 interaction with the channel increased FRET signal between fluorescent probes on the TRPV4 cytosolic tails, consistent with a more compact cytosolic region. This is the first piece of evidence suggesting that, similar to PIP2 -regulated K+ channels, PIP2 interaction with the TRPV4 channel rearranges cytosolic domains. RESULTS AND DISCUSSION A possible phosphoinositide interacting site in the TRPV4 N-tail is required for channel activation by hypotonicity and heat. We Reserved for Publication Footnotes

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Fig. 1. Functional analysis of N-terminal truncations and mutations of TRPV4. (A)Mean current density measured at +100 and -100 mV in response to a 30% hypotonic shock in HEK-293 cells overexpressing TRPV4-WT, TRPV4Δ1-30, TRPV4-Δ1-130, TRPV4-Δ100-130 and GFP.Number of cells recorded is shown for each condition.(B)Ramp current-voltage relations of cationic currents recorded from HEK-293 cells transfected with TRPV4-WT or TRPV4121 AAWAA and exposed to 30% hypotonic shocks. (C) Mean current responses to isotonic and hypotonic stimuli in cells transfected with TRPV4-WT or TRPV4-121 AAWAA. (D) Mean current responses to 4α-PDD stimulation in TRPV4-WT or TRPV4-121 AAWAA expressing cells. (E) Calcium signals (Fura2 ratio) obtained in HeLa cells transfected with GFP (n=47), TRPV4-WT (n=25) or TRPV4-121 AAWAA (n=59) and sequentially stimulated with 30% hypotonic solutions and 10 μM 4αPDD. (F) Calcium signals obtained in HeLa cells transfected with TRPV4-WT (n=335), TRPV4-121 AAWAA (n=318) or GFP (n=254) and stimulated with warm solutions (38ºC). * P<0.05. Fig. 2. . Effect of PIP2 depletion on TRPV4-mediated Ca2+ signals. HeLa cells were transfected with TRPV4-WT, TRPV4-WT plus FRB and FKBP-5phosphatase (PIP2 -ptase) or the inactive phosphatase (D281A). (A-C) Average calcium signals (fura-2 ratio) measured in the presence of the phosphatase translocation inducing agent rapamycin (1 μM) for cells exposed to (A) 30% hypotonicity (TRPV4, n=302; V4+PIP2 -Ptase, n=229; V4+PIP2 PtaseD281A, n=192), (B) heat (TRPV4, n=150; V4+PIP2 -Ptase, n=146; V4+PIP2 PtaseD281A, n=89) and (C) 4α-PDD (TRPV4, n=278; V4+PIP2 -Ptase, n=233; V4+PIP2 -PtaseD281A, n=89). (D-G) Representative intracellular Ca2+ signals obtained from mouse tracheal ciliated cells exposed to a hypotonic solution in the absence (D) or the presence (E) of 20 μM ATP, or exposed to heat (38ºC) in the absence (F) or the presence (G) of 20 μM ATP. Percentage of ciliated cells responding to hypotonicity and heat was >90%. In the presence of ATP the percentages were <5% (hypotonicity) and >90% (heat).

screened TRPV4 for domains that may participate in the channel response to hypotonicity-induced cell swelling. A schematic view of the channel molecule and all deletions/mutations generated is shown in Supporting Information Fig. S1. We focused on the N-terminus tail because a TRPV4 SNP associated with hyponatremia, P19S, generates a channel with reduced response to hypotonic cell swelling (37). We generated three deletions of different length: TRPV4-Δ1-30, TRPV4-Δ1-130 and TRPV4-Δ100-130. TRPV4 sensitivity to hypotonic conditions was assayed using whole-cell patch clamp recordings and intracellular calcium imaging with fura-2. Addition of a 30% hypotonic solution yielded large currents in cells transiently transfected with TRPV4-WT but not in GFP-transfected cells (Fig. S2A). TRPV4-Δ1-130 and TRPV4-Δ100-130 responses to 30% hypotonicity were reduced to the levels recorded in GFP-transfected cells while TRPV4-Δ1-30 displayed a normal response (Fig. 1A and Fig. S2B). Deletion of the first 30 residues did greatly reduce currents generated by 15% hypotonic solutions (Fig. S2D), mimicking the changes induced by the P19S polymorphism (37). Consistent with the electrophysiological experiments, expression of TRPV4-WT increased intracellular [Ca2+ ] in response to 30% hypotonicity while the
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response of TRPV4-Δ100-130 was indistinguishable from that obtained in GFP-transfected cells (Fig. S2C). To evaluate if differences in plasma membrane expression between WT and mutant TRPV4 proteins could explain the reduced responses of truncated TRPV4 proteins we determined surface labeling of HEK293 cells expressing TRPV4-WT and TRPV4-Δ1-130 proteins tagged with a V5 epitope in the first extracellular loop. Confocal microscopy images and quantification by ELISA revealed no apparent differences in membrane expression between TRPV4WT and the protein presenting the longest truncation, TRPV4Δ1-130 (Fig. S3). To pin down the region within residues 100-130 required for the channel response to hypotonic cell swelling we neutralized four positive charges within a sequence (121 KRWRK125 ) that has been proposed to be a possible phosphoinositide binding site (PIsite) (29). Cells transfected with TRPV4-121 AAWAA displayed greatly reduced swelling-induced whole-cell cationic currents (Fig. 1B-C). TRPV4-121 AAWAA generated currents in response to the synthetic agonist 4α−PDD (0.01-10 μM) were undistinguishable from TRPV4-WT currents (Fig. 1D). Sequential addition of a hypotonic solution and 10 μM 4α−PDD generated
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Fig. 3. Effect of PIP2 on TRPV4 channel activity in inside-out patches. (A) Two TRPV4 single-channel openings, which disappeared within seconds, were observed at +80 mV immediately after excision of a HeLa cell membrane patch (top). Addition of diC8-PIP2 (50 μM) after complete channel rundown did not reactivate TRPV4 (middle) while addition of 5,6-EET (1 μM) in the presence of PIP2 activated TRPV4 (bottom). (B) Recordings obtained at +80 mV in a patch sequentially exposed to EET (1 μM) and 4α-PDD (10 μM). (C), Mean open probability (NPo ) calculated from control patches (2 min after excision) and in response to PIP2 , EET, EET+PIP2 and 4α-PDD (number of patches given in the figure). Percentages patches presenting TRPV4 activity: PIP2 20%, EET 20%, EET+PIP2 71% and 4α-PDD 88%. (D-E) Single channel recordings obtained from the same excised patch in response to 24ºC and warm solutions (38ºC) in the presence (D) or in the absence (E) of 50 μM diC8PIP2 . (F), NPo calculated from consecutive 5 sec recordings following exposure to warm solutions and plotted versus time (+PIP2 , n=11; -PIP2 , n=7; +PIP2 +1 μM HC-067047 (in pipette solution), n=4; TRPV4-121 AAWAA + PIP2 , n=5). * P<0.05.

significant increases in intracellular Ca2+ levels in cells expressing TRPV4-WT while cells expressing TRPV4-121 AAWAA only responded to 4α−PDD (Fig. 1E). TRPV4 is also activated by moderate heat (above 25ºC) (8, 21), although the mechanism of its temperature sensitivity is not fully understood (8). Ca2+ imaging on cells exposed to warm temperatures (38ºC) revealed a typical transient response in cells transfected with TRPV4-WT channels. Neutralization of the positive charges in the TRPV4-121 AAWAA decreased the Ca2+ response to the levels obtained in GFP-transfected cells (Fig. 1F). We hypothesized that the sequence 121 KRWRK125 may form a PI-site required for phosphatidylinositol-4,5-biphosphate (PIP2 ) interaction with TRPV4 to respond to hypotonic and heat stimulation. Different TRP protein sequences containing several positively charged amino acids have been proposed to interact with phosphoinositides, particularly PIP2 (29, 38). To examine how specific was the neutralization of the 121 KRWRK125 positive charges, we neutralized three positive charges of a nearby reFootline Author

Fig. 4. PIP2 binding to the TRPV4 N-tail. (A), Coomassie-stained SDSPAGE showing protection from limited papain digestion by PIP2 but not PI. Purified protein corresponding to residues 1-397 of human TRPV4 (150 μM) was digested with papain (38 nM) in the absence or presence of lipid (PI and PIP2 at 10 μM). The cleavage positions corresponding to each isolated band, determined by N-terminal sequencing, are indicated. (B-E) The four indicated bands were scanned, quantified and plotted versus digestion time. Significant changes were observed in the presence of PIP2 for bands 1 and 3 at all times while band 4 showed significant differences at time 45 and 60 min. Mean±S.D. (n=3). * P<0.05 control vs PIP2 ; ** P<0.01 control and PI vs PIP2 .

gion (114 RHH116 ). Expression of TRPV4-114 AAA produced hypotonicity and heat-induced Ca2+ increases similar to those obtained with TRPV4-WT (Fig S4A). Together, these experiments suggested that residues 121 KRWRK125 are critical for TRPV4 activation by hypotonicity and heat, but not necessary for channel activation by 4α−PDD. Depletion of PIP2 levels prevents channel activation by physiological stimuli. We assessed whether deletion or mutation of residues 121 KRWRK125 may be related to a PIP2 -dependent mode of TRPV4 gating. For that purpose, we evaluated the impact of reducing PIP2 levels on channel activation. We used a rapamycin-induced translocatable 5-phosphatase to deplete PIP2 (39). The membrane-localized rapamycin-binding protein FRB and the cytoplasmic enzyme construct FKBP-5-phosphatase were co-transfected with TRPV4-WT in HeLa cells. Addition of
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Fig. 5. PIP2 -dependent rearrangment of TRPV4 cytosolic tails. (A-B), Mean current density of hypotonicity-activated (A) or 4α-PDD-activated (B) currents recorded from HEK-293 cells transfected with GFP, TRPV4, TRPV4+PACSIN3 or TRPV4+PACSIN3-ΔF-BAR. (C) FRET efficiency was determined at the plasma membrane of HEK-293 coexpressing soluble YFP and CFP-fused TRPV4, CFPand YFP-fused TRPV4-WT, CFP- and YFP-fused TRPV4-121 AAWAA, CFP- and YFP-fused TRPV4-Δ1-130, or CFP- and YFP-fused TRPV4-WT coexpressed with either PACSIN3 or PACSIN3-ΔF-BAR. (D) FRET efficiencies between CFP- and YFP-fused TRPV4-WT determined at the plasma membrane in the absence or presence of tetracyclin. TRPV4 constructs were transiently cotransfected in HEK-293 cells expressing a tetracyclin-inducible 5-phosphatase. Number of cells recorded is shown for each condition. Mean±S.E.M. * P<0.05 versus TRPV4-WT, one way ANOVA and Bonferroni post hoc (A-C) or Student’s t test (D).

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rapamycin to translocate the phosphatase to the plasma membrane, locally depleted membrane PIP2 (Fig. S5A-C) and prevented the increase of the Ca2+ signal following hypotonic cell swelling (Fig. 2A) and heat stimulation (Fig. 2B) without affecting the response to 0.1-10 μM 4α-PDD (Fig. 2C; and Fig. S4B). Application of rapamycin to cells cotransfected with TRPV4 and a phosphatase-dead mutant (D281A) (39) did not affect the Ca2+ response to any of the stimuli tested (Fig. 2A-C). We also analyzed whether phospholipase C (PLC)-induced depletion of PIP2 decreased TRPV4 channel activity in native cells. For that purpose we used primary cultures of ciliated epithelial cells obtained from trachea and oviduct, which express functional TRPV4 channels (7, 16, 17). Figure 2D shows typical oscillatory Ca2+ signals generated by hypotonic solutions in tracheal ciliated epithelial cells. However, Ca2+ response to hypotonicity was abrogated following the activation of PLC with ATP, which leads to the hydrolysis of PIP2 and the generation of an IP3 -mediated Ca2+ signal (Fig. 2E). The heat response of epithelial cells was also reduced following the addition of ATP (Fig. 2F-G). The reduction in hypotonicity- and heat-induced Ca2+ signal were not due to Ca2+ -dependent inhibition of TRPV4 as two consecutive stimuli elicited similar responses (Fig. S6A-B). Similarly, mouse ciliated oviductal cells responses to hypotonicity were reduced following addition of ATP (Fig. S6C-D). Although we could not asses directly whether PIP2 remained depleted at the time cells were challenged with TRPV4 activating stimuli, the fact that there was no Ca2+ response to a second ATP stimulation within minutes of the first ATP application (Fig. S6E) may reflect a condition of PIP2 depletion.
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Activity of PIP2 -regulated channels typically decreases in excised inside-out patches and recovers upon addition of exogenous PIP2 (40). In those excised patches in which TRPV4 channel activity was present immediately after excision, channel activity decreased with time and addition of the water soluble diC8-PIP2 (50-200 μM) or long acyl chain PIP2 (10 μM) did not recover initial channel activity (Fig. 3A and C). The fact that PIP2 was not able to activate TRPV4 in excised patches may indicate that following patch excision another, yet unidentified, modulator required for channel activity is lost. Hypotonicity-mediated activation of TRPV4 in excised patches can not be directly evaluated. Instead, the osmotransducing cytosolic messenger 5’-6’epoxyeicosatrienoic acid (EET) has been used (20, 41). Addition of EET (1 μM) in the presence of PIP2 activated TRPV4 in 71% of patches (Fig. 3A and C). However, addition of EET in the absence of PIP2 only activated 20 % of patches (Fig. 3B and C), even though TRPV4 channel activity in the same patches was demonstrated using 4α−PDD (Fig. 3B). Next, we tested channel activation by heat in excised insideout patches obtained from HeLa cells overexpressing TRPV4. In the presence of PIP2 TRPV4-WT channel activity was detected within seconds after application of warm solutions (Fig. 3D) while in the presence of PIP2 and the TRPV4 blocker HC-067047 (42) or with the TRPV4-121 AAWAA no channel activity was elicited by heat (Fig. 3F). We discarded a shear-stress dependent component under our experimental conditions for heat activation of TRPV4 (Fig. S7A). In the absence of PIP2 , and consistent with previous reports (8, 21, 43), no significant change in channel activity was elicited by heat (Fig. 3E). Fig. 3F shows mean channel activity in response to heat and plotted versus time after addition of warm solutions in the presence or absence of PIP2 . The TRPV4 Q10 obtained from excised patches containing TRPV4-WT in the presence of PIP2 was 21±5 (n=3) (Fig. S7B), consistent with previous values obtained from TRPV4 whole-cell recordings (21, 43). Together these experiments confirm that PIP2 is required for TRPV4 activation by physiological stimuli, probably acting as an allosteric modulator. However, at present we do not have a comprehensive model to incorporate all factors involved in TRPV4 gating, i.e., why TRPV4 gating by 4α−PDD is not affected by PIP2 depletion or why PIP2 is unable to activate TRPV4 on its own. PIP2 interacts with the TRPV4 N-tail. To further characterize PIP2 interaction with the TRPV4 N-tail, we carried out limited proteolysis assays on the purified TRPV4 N-terminal region (residues 1-397), which includes the N-terminal tail and the ankyrin repeats. Papain digestion led to cleavage at four positions within the N-tail (Fig. 4A). Quantification of the bands obtained (Fig. 4B-E) showed that proteolysis of TRPV4 N-terminus is reduced in the presence of PIP2 but not PI. PIP2 -dependent proteolysis protection was not observed with the isolated TRPV4 ankyrin repeats (residues 136-397), the TRPV1 ankyrin repeats or the TRPV4-121 AAWAA N-terminal region (Fig. S8), ruling out non-specific inhibition of papain by PIP2 . These biochemical data therefore support a direct interaction of PIP2 with the N-tail region of TRPV4-WT. PACSIN3 F-BAR domain is required for TRPV4 channel regulation. The effect of neutralizing the positively charged residues, or depleting PIP2 levels, on channel activity resembled the response of TRPV4 when coexpressed with PACSIN3, i.e., reduced channel response to hypotonicity and heat but normal response to 4α-PDD (25, 44). PACSIN3 belongs to a family of proteins that contain a Bin-Amphiphysin-Rvs (BAR) domain required to penetrate and remodel the plasma membrane (45, 46). PACSIN3 binds through its SRC homology 3 (SH3) domain to the PRD of TRPV4 (44), in the close proximity of the PI-site. Two competing hypotheses are that membrane-bound PACSIN3 binding to the PRD may either promote or physically block the interaction
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of the PI-site with membrane PIP2 . To test these hypotheses, we generated a PACSIN3 lacking the F-BAR domain. Similar deletion in PACSIN1 renders the protein unable to interact with the lipids of the plasma membrane (47). The F-BAR domain of PACSIN3 is not required for interaction with TRPV4 (44). Accordingly, we detected interaction of PACSIN3-ΔF-BAR with TRPV4 (Fig. S9). Coexpression of TRPV4 with PACSIN3-ΔFBAR, unlike coexpression with PACSIN3, did not reduce the whole-cell currents generated by hypotonic challenges (Fig. 5A). The channel response to 4α-PDD was not affected under any of the experimental conditions tested (Fig. 5B). These results were therefore consistent with the hypothesis that PACSIN3 interferes with the interaction of TRPV4 with PIP2 , an effect that was lost when a membrane-unbound PACSIN3-ΔF-BAR was used. PIP2 rearranges TRPV4 cytosolic tails. Together, our findings underscore the involvement of PIP2 in TRPV4 gating by physiological stimuli. However, an important question remained that has not been resolved for any PIP2 -modulated TRP channel yet. Does PIP2 binding affect the structural conformation of TRPV4? We approached this question studying the impact of TRPV4 deletions and mutations on the conformation of cytosolic tails. For that purpose we evaluated the proximity of the intracellular C-tails of CFP- and YFP-tagged TRPV4 proteins, which we assumed formed a random population of heteromeric channels, by fluorescence resonance energy transfer (FRET). We tagged C-tails, which remained unmodified in all the TRPV4 deletions/mutations generated, to avoid possible FRET artifacts generated by the different lengths of the N-tails. The relative CFP and YFP fluorescence intensities in the plasma membrane were determined for every single cell and used to calculate FRET efficiencies in transiently transfected HEK-293 cells (Fig. 5C). TRPV4-WT generated a FRET ratio similar to that previously reported (48) while TRPV4-Δ1-130 and TRPV4-121 AAWAA doubled the FRET ratio, indicating a more compacted tail conformation. Similarly, coexpression of TRPV4-WT with PACSIN3 markedly increased the FRET signal, an effect that was lost when coexpressed with PACSIN3-ΔF-BAR. We reasoned that the increased FRET observed with mutant TRPV4 proteins or when coexpressed with PACSIN3 was due to the inability of TRPV4 to interact with membrane PIP2 . To test this hypothesis, we studied how the reduction in PIP2 levels affected FRET efficiency of TRPV4-WT. We overexpressed CFP- and YFP-tagged TRPV4WT channels in HEK293 cells engineered with tetracyclineinducible expression of 5-phosphatase IV (33). Induction of this enzyme depleted PIP2 from the plasma membrane (Fig. S5D) and significantly increased FRET ratio (Fig. 5D). This observation further supported the hypothesis that conditions that prevented the N-tail access to membrane PIP2 (by deletion/mutation of the PI-site or by overexpression of PACSIN3) or depleted PIP2 from the plasma membrane rearranged the cytosolic TRPV4 tails into a more compacted conformation (increased FRET ratio). Thus, in the presence of PIP2 and an intact PI-site the intracellular tails appeared in an expanded conformation. Conclusions. Together, our data provide several new findings. First, we have demonstrated that TRPV4, as many other TRP channels, is regulated by PIP2, a process that involves PIP2 binding to a PI-site (121 KRWRK125 ) in the N-tail. Second, we have showed that PIP2 regulates channel activity in a stimulus-dependent manner. Third, TRPV4 is bona fide thermosensitive channel, providing there is PIP2 to interact with the N-tail. Fourth, the interaction of the TRPV4 PI-site with plasma membrane PIP2 favors an expanded conformation of the intracellular tails and channel activation by hypotonicity and heat. Conditions that reduced PIP2 levels (inducible phosphatase) or interfere the interaction of TRPV4 with PIP2 (mutations in the PI-site or coexpression with PACSIN3) promote a compacted tail conformation and prevent channel activation by hypotonicity and heat. Our study

provides the first piece of evidence suggesting that, similar to PIP2 regulated K+ channels, PIP2 interaction with TRPV4 channel rearranges the cytosolic domains. Whether the intracellular tail rearrangement occurring upon PIP2 binding to TRPV4 facilitates the access of stimuli-generated messengers (e.g., EET) to their binding sites or favors the stimulus-dependent opening of the gates themselves it is not known at present. MATERIALS AND METHODS
Cells and transfection For electrophysiological or calcium imaging experiments HeLa or HEK293 cells were transiently transfected as previously described (16, 48). Primary cultures of tracheal and oviductal ciliated cells were obtained as previously described (7, 17). Animals were maintained and experiments were performed according to the guidelines issued by the Institutional Ethics Committee of the Universitat Pompeu Fabra. Solutions Isotonic bath solutions used for imaging experiments contained (in mM): 140 NaCl, 2.5 KCl, 1.2 CaCl2 , 0.5 MgCl2 , 5 glucose and 10 HEPES, pH 7.3 with Tris. Bath solutions for whole cell recordings contained (in mM): 100 NaCl, 1 MgCl2 , 6 CsCl, 10 HEPES, 1 EGTA and 5 glucose, pH 7.3 with Tris. Osmolarity was adjusted to 310 mOsm using mannitol. 30% and 15% hypotonic solutions (255 and 220 mOsm) were obtained by removing mannitol. Wholecell pipette solution contained (in mM): 20 CsCl2 , 100 CsAcetate, 1 MgCl2, 0.1EGTA, 10 HEPES, 4 Na2 ATP and 0.1 NaGTP; 300 mOsm, pH 7.25. Bath and pipette solutions for excised inside-out single channel recordings contained (in mM): 130 CsCl, 1 MgCl2 , 1 Na2 ATP, 0.034 CaCl2 , 5 EGTA, 10 HEPES (310 mosmol/liter, pH 7.25). When required, solutions were warmed using a water jacket device (Warner Instruments). All chemicals were obtained from Sigma-Aldrich except diC8-PI and diC8-PI(4,5)P2 (Echelon Biosciences Inc.), HC-067047 (Tocris Biosciences) and Fura-2 (Invitrogen). Electrophysiological and Ratiometric Ca2+ recordings Patch-clamp whole-cell and single single-channel currents were recorded at room temperature (∼24ºC, unless otherwise indicated) as previously described (16, 48). Cells/excised patches were perfused at 0.8 ml/min. Cytosolic Ca2+ signals, relative to the ratio (340/380) measured prior to cell stimulation, were obtained from cells loaded with 4.5 μM fura-2 AM as previously described (4). FRET measurements FRET measurements were carried out in a Leica TCS SP2 confocal microscope (Leica) attached to an inverted microscope. FRET efficiencies expressed as the increase of the FRET donor CFP after bleaching the FRET acceptor YFP (48). Lipid protection assay Human TRPV4 ankyrin repeats (136-397) and N-tail (1-397) were cloned using NdeI and NotI into pET21-C6H (49). Recombinant proteins were produced and purified as described (50), except the size exclusion chromatography buffer was 10 mM Tris-HCl pH 7.0, 300 mM NaCl, 10 % glycerol, and 1 mM DTT for TRPV4 N-tail, and 10 mM Tris-HCl pH 7.0, 150 mM NaCl and 1 mM DTT for TRPV4 ankyrin repeats. Lipid protection assay by limited proteolysis was performed at 4°C (on ice) in reaction buffer containing (in mM): 180 NaCl, 20 Tris-HCl pH 7.0, 1 % glycerol and 1 DTT (for TRPV4-1-397) or 150 NaCl, 20 Tris-HCl pH 7.0 and 1 DTT (for TRPV4-136-397 and TRPV1-ARD). Proteins were pre-incubated in the absence or presence of PI or PIP2 at 4°C for 60 min and then digested with papain. Final concentrations of protein, lipid, and papain were 10 μM, 150 μM and 38 nM, respectively. Digestion was stopped at 15, 30, 45 and 60 min by adding SDS sample buffer, and samples separated by SDS-PAGE and visualized by Coomassie staining. The gels were scanned and signals were quantified with ImageJ. Statistical analysis Data are expressed as mean±SEM (or mean±S.D. in Fig. 4) of n experiments. Statistical analysis was assessed with Student’s unpaired test or oneway analysis of variance (ANOVA) using Sigma-Plot software. Acknowledgments. We thank Dr T. Voets (KU Leuven, Belgium) for the gift of HEK-293 tetracycline-inducible phosphatase expressing cells, Dr. T. Meyer (Stanford, U.S.A.) for rapamycin-inducible phosphatases and Dr. M. Schaefer (Leipzig, Germany) for help with initial FRET experiments. This work was supported by the Spanish Ministry of Science and Innovation (SAF2012-38140); Fondo de Investigación Sanitaria (Red HERACLES RD12/0042/0014); FEDER Funds; Generalitat de Catalunya (SGR05-266); and National Institutes of Health (R01GM081340). M.A.V. is the recipient of an ICREA Academia Award and U.A.H. of an EMBO Long-Term Fellowship. Footnotes 1 A.G-E. and S.M. contributed equally to this work. 2 To whom correspondence should be addressed. E-mail: miguel.valverde@upf.edu Author contributions: A.G-E., R.G., R.V. and M.A.V. designed research; A.G-E, S.M., C.P-P, H.I., F.R-M., C.P.,U.A.H. and R.V. performed research; A.G-E, S.M., C.P-P., H.I., U.A.H. and M.A.V. analyzed data; and M.A.V. wrote the paper. All authors collaborated in paper edition. The authors declare no conflict of interest. This article contains supporting information online

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