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Originally published In Press as doi:10.1074/jbc.M600549200 on March 29, 2006

J. Biol. Chem., Vol. 281, Issue 22, 15485-15495, June 2, 2006
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A Role for AQP5 in Activation of TRPV4 by Hypotonicity

CONCERTED INVOLVEMENT OF AQP5 AND TRPV4 IN REGULATION OF CELL VOLUME RECOVERY*Formula

Xibao Liu{ddagger}, Bidhan Bandyopadhyay{ddagger}, Tetsuji Nakamoto§, Brij Singh, Wolfgang Liedtke||, James E. Melvin§, and Indu Ambudkar{ddagger}1

From the {ddagger}Secretory Physiology Section, Gene Therapy and Therapeutics Branch, NIDCR, National Institutes of Health, Bethesda, Maryland 20892, the §Center for Oral Biology, University of Rochester, Rochester, New York 14642, the Department of Biochemistry and Molecular Biology, University of North Dakota, Grand Forks, North Dakota 58203, and the ||Center for Translational Neuroscience, Duke University Medical Center, Durham, North Carolina 27710

Received for publication, January 19, 2006 , and in revised form, March 21, 2006.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Regulation of cell volume in response to changes in osmolarity is critical for cell function and survival. However, the molecular basis of osmosensation and regulation of cell volume are not clearly understood. We have examined the mechanism of regulatory volume decrease (RVD) in salivary gland cells and report a novel association between osmosensing TRPV4 (transient receptor potential vanalloid 4) and AQP5 (aquaporin 5), which is required for regulating water permeability and cell volume. Exposure of salivary gland cells and acini to hypotonicity elicited an increase in cell volume and activation of RVD. Hypotonicity also activated Ca2+ entry, which was required for subsequent RVD. Ca2+ entry was associated with a distinct nonselective cation current that was activated by 4{alpha}PDD and inhibited by ruthenium red, suggesting involvement of TRPV4. Consistent with this, endogenous TRPV4 was detected in cells and in the apical region of acini along AQP5. Importantly, acinar cells from mice lacking either TRPV4 or AQP5 displayed greatly reduced Ca2+ entry and loss of RVD in response to hypotonicity, although the extent of cell swelling was similar. Expression of N terminus-deleted AQP5 suppressed TRPV4 activation and RVD but not cell swelling. Furthermore, hypotonicity increased the association and surface expression of AQP5 and TRPV4. Both these effects and RVD were reduced by actin depolymerization. These data demonstrate that (i) activation of TRPV4 by hypotonicity depends on AQP5, not on cell swelling per se, and (ii) TRPV4 and AQP5 concertedly control regulatory volume decrease. These data suggest a potentially important role for TRPV4 in salivary gland function.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
The ability of cells to regulate their volume is essential for maintenance of cellular homeostasis under anisotonic environmental conditions (1–3). Changes in osmolarity of the extracellular medium induce water fluxes that result in swelling or shrinkage of cells, depending on the osmotic gradient. Most cells respond to changes in tonicity and cell volume by initiating mechanisms that allow them to recover their original volume in the continued presence of the osmotic stress. Such regulatory volume changes depend on the activation of cation and anion permeabilties that reverse the osmotic gradient and direction of water flow (1–4). Emerging studies demonstrate that regulatory volume changes are critical for cell survival and also for regulation of cellular processes such as gene transcription and proliferation (1). In addition, a variety of cells, such as exocrine gland cells, utilize the mechanism of regulatory volume change to drive fluid secretion (5). Several monovalent cation and anion channels as well as intracellular Ca2+ changes contribute to cell volume regulation. For example, hypo-osmolarity-induced cell swelling has been associated with a rise in cytosolic Ca2+ concentration ([Ca2+]i) in different cell types, which is due to hypotonicity-activated Ca2+ entry pathways. It has also been clearly demonstrated that this Ca2+ entry is critical for regulating the ion fluxes that drive volume decrease (1–4). However, the underlying mechanism(s) that senses the change in osmolarity and/or cell volume to initiate volume regulation is poorly understood (5).

Regulation of transepithelial osmotic forces as well as cell volume critically impacts salivary gland fluid secretion induced by neurotransmitter stimulation of the gland (5). The water channel, AQP5 (aquaporin 5) provides a regulated water permeability across the apical membrane of salivary gland acinar cells that is important not only for fluid secretion but also for regulatory volume changes (5–7). Aqp5–/– mouse salivary glands show decreased salivary secretion in response to muscarinic receptor stimulation. Additionally, dispersed salivary gland acini from these mice have reduced ability to control volume changes in response to hyper- or hyposmotic solutions (7). Thus, fluid secretion and cell volume regulation converge at the level of AQP5, which mediates the final step in both processes (i.e. water efflux). However, it is not clear how cells sense the change in osmolarity and how this signal is transduced to achieve volume regulation.

As discussed above, hypotonic and hypertonic conditions induce [Ca2+]i increases in different cell types, and Ca2+ entry has been reported to be critical for the volume recovery process (1–4). Neurotransmitter-stimulated Ca2+ mobilization events, including intracellular Ca2+ release and Ca2+ entry as well as Ca2+-dependent activation of cation and anion channels, have been quite extensively studied in salivary and other exocrine gland cells (5, 8). Further, increases in cytosolic Ca2+ due to Ca2+ entry have been correlated with ion channel activation and fluid secretion. However, the Ca2+ entry mechanism(s) involved in the cellular response to anisosmotic conditions has not yet been identified (5). TRPV4 (transient receptor potential vanilloid 4), a member of the TRP superfamily of cation channels (9) has been shown to be activated by hypotonicity and a variety of other stimuli (10–15). This channel is expressed in several cell types and has been reported to be involved in RVD2 in airway epithelial and keratinocyte cell lines (16, 17). TrpV4–/– mice display impaired regulation of systemic tonicity (18), although exactly how TRPV4 activity leads to regulation of cell volume has not yet been directly demonstrated. The molecular mechanism by which tonicity changes regulate TRPV4 has also not yet been established (11, 12, 14, 16–18).

This study was directed toward defining the molecular basis of RVD. Toward this end, we have measured the response of salivary gland cell lines, primary cultures, and dispersed salivary gland acini to hypotonicity. These cells have been previously shown to display robust volume changes as well as efficient volume recovery in response to anisosmotic conditions (5, 7). The exact molecular events involved in these responses are not yet known. The data presented here demonstrate a novel association between TRPV4 and AQP5, which controls RVD in salivary gland cells. We show that AQP5 is required for the activation of TRPV4 by hypotonicity. Further, AQP5 and TRPV4 are concertedly involved in regulating recovery of cell volume.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Cell Culture and Transfection—Cells were cultured in Dulbecco's modified Eagle's medium (rat basophilic leukemia cells; RBL-2H3), Earle's minimal essential medium (human submandibular gland (HSG) and human parotid gland (HSY) cells) supplemented with 10% fetal calf serum, 2 mM glutamine, 1% penicillin/streptomycin at 37 °C in 5% CO2. HSG cells were transiently transfected with 1 µg of required plasmids and 0.2 µg of green fluorescent protein-encoding plasmid or only with green fluorescent protein plasmid using Lipofectamine reagent 2000 (Invitrogen).

Dispersed Cell Preparation from Mouse Parotid and Submandibular Gland Cells—All mice were maintained according to guidelines approved by the NIDCR, National Institutes of Health, Animal Care and Use Committee. Submandibular glands were removed, cleaned, minced, and digested in standard external solution containing 145 mM NaCl, 5 mM KCl, 1 mM MgCl2, 1 mM CaCl2, 10 mM Hepes, 10 mM glucose, pH 7.4 (NaOH), with 0.02% soybean trypsin inhibitor and 0.1% bovine serum albumin containing collagenase P (2.5 mg/8 ml) (7). After a 15–20-min incubation at 37 °C, the digest was washed twice with the normal external solution and resuspended in external solution.

Electrophysiological Recording—The patch pipette had resistances between 3 and 5 megaohms after filling with the standard intracellular solution that contained 145 mM cesium methane-sulfonate, 8 mM NaCl, 10 mM MgCl2, 10 mM HEPES, 10 mM EGTA, pH 7.2 (CsOH). External solutions were composed as follows: 145 mM NaCl, 5 mM CsCl, 1 mM MgCl2, 10 mM CaCl2, 10 mM HEPES, 10 mM glucose, pH 7.4 (NaOH) (Ca2+ and Na+ solution); 170 mM NMDG, 5 mM CsCl, 1 mM MgCl2, 10 mM HEPES, 10 mM glucose, pH 7.4 (HCl) (NMDG solution). Osmolarity for all solutions was adjusted with D-mannitol to 305 ± 5 mmol/kg using a vapor pressure osmometer (Wescor). For measuring swelling-activated currents, an isotonic solution containing 75 mM NaCl, 6 mM CsCl, 5 mM CaCl2, 1 mM MgCl2, 10 mM Hepes, 150 mMD-mannitol, and 10 mM glucose, pH 7.4, with NaOH (305 ± 5 mmol/kg) was used. Cell swelling was induced by omitting mannitol from this solution to reach proper osmolarity. In some experiments, the reduction of osmolarity was achieved by reduction of NaCl from normal external solution where indicated.

Patch clamp experiments were performed in the tight seal whole-cell configuration at room temperature (22–25 °C) using an Axopatch 200B amplifier (Axon Instruments) as described earlier (19–21). Voltage ramps ranging from –90 to 90 mV over a period of 1 s were imposed every 4 s from a holding potential of 0 mV and digitized at a rate of 1 kHz. A liquid-junction potential of less than 8 mV was not corrected, and capacitative currents and series resistance were determined and minimized. For analysis, the first ramp was used for leak subtraction for the subsequent current records. There was no significant increase in the current under these conditions unless cells were exposed to hypotonic external solution (HTS).

Measurement of Intracellular Ca2+ Concentration—Freshly isolated salivary gland cells were loaded with fura-2 for 45–60 min at 30 °C and allowed to attach to a glass bottom dish (Matek Corp.). Ducts and acinar cells were morphologically identified (7). Other cells were cultured overnight in Matek dishes. Fluorescence measurements were made using a Till Photonics-Polychrome IV spectrofluorimeter attached to a Olympus X51 microscope and Metafluor imaging system (Universal Imaging Corp.) (21). Fluorescence traces shown represent [Ca2+]i (values are averages from >50 cells and representative of results obtained in at least 3–5 individual experiments).

Measurement of Cell Volume—Cells were loaded with the fluoroprobe calcein (Molecular Probes, Inc., Eugene, OR) and excited at 490 nm. Emitted fluorescence was measured at 510 nm. In situ calibration of the dye was performed. The relationship between dye fluorescence and the volume change was linear over a volume range from +35 to –35%. In some experiments, cell volume was estimated using an Olympus X51 microscope interfaced with Universal Imaging MetaMorph software. Data are presented as mean ± S.E. Origin 7.5 (OriginLab, Northampton, MA) was used for data analysis and display. Significant difference between individual groups was tested by using analysis of variance.

Immunoprecipitation and Immunoblotting—Immunoprecipitation was done using solubilized crude membranes or from cell lysates as previously described (21). Precleared lysates were incubated with the required antibody (1:200 dilution of anti-TRPV4 or anti-AQP5). Interacting proteins bound to Sepharose beads were separated, released with SDS-PAGE sample buffer, and detected by Western blotting as described previously using anti-TRPV4 (1:500 dilution) or anti-AQP5 antibody (1:1000 dilution).

Immunocytochemistry—Parotid glands were excised from the animals and fixed in 10% formalin solution, embedded in paraffin, and used to prepare 5–10-µm sections (American Histolabs, Gaithersburg, MD). Sections were dewaxed, rehydrated, and permeabilized with 0.5% Triton in PBS, pH 7.5. Streptavidin-peroxidase reactions using the DAB Histostain kit (Zymed Laboratories, San Francisco, CA) were used to detect specific proteins (anti-TRPV4 at 1:70 and anti-AQP5 at 1:100 dilutions). In control sections, rabbit IgG was used instead of primary antibody.

Surface Biotinylation—Cells were treated as required and incubated with 0.5 mg/ml Sulfo-NHS-Biotin (Pierce) on ice (21), washed with buffer containing 0.1 M glycine, and solubilized with 2 ml of cell lysis buffer containing Nonidet P-40, 0.1% SDS, and proteolytic inhibitors. Biotinylated proteins were pulled down with Neutr-Avidin-linked beads (Pierce). The bound fraction was washed and released with SDS-PAGE sample buffer and analyzed by Western blotting.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Hypotonicity Stimulates Changes in Cell Volume and Ca2+ Entry via a TRPV4-like Channel in Salivary Gland Cells—Fig. 1A shows that HTS induced a sustained increase in [Ca2+]i in HSG (human submandibular gland cell line) cells, which was dependent on the osmolarity. Average data representing peak increase in fluorescence from these experiments are shown in Fig. 1B (significant increases above resting [Ca2+]i are indicated). Primary cultures of human submandibular gland cells (22) but not rat basophilic leukemia (RBL) cells displayed a similar response to HTS (Fig. 1C; 150 mmol/kg solution was used here and in subsequent experiments). The sustained [Ca2+]i increase in HSG cells was blocked by 100 µM but not 1 µM Gd3+ (Fig. 1D), indicating that HTS-stimulated Ca2+ entry is unlike store-operated Ca2+ entry, which is blocked by 1 µM Gd3+ in these cells (19). This was further confirmed by whole cell current recordings. HTS increased HSG cell membrane conductance with a 25–40 s delay (Fig. 1E) and generated a weakly outwardly rectifying current that reversed at +6 ± 3 mV (Fig. 1F; HTS-stimulated [Ca2+]i increase, and cation currents were seen in 91% (20 of 22) of cells). With NMDG as the cation in the external solution (Fig. 1, G and H), inward current was greatly reduced, outward current remained relatively unchanged, and reversal potential showed a left shift (this effect was seen in all of the cells that displayed a response to HTS). Together, these data demonstrate that HTS stimulates a nonselective Ca2+-permeable cation conductance that is distinct from the relatively inwardly rectifying store-operated Ca2+ current ISOC previously described in HSG cells (19, 20). However, the characteristics of the HTS-stimulated cation current in HSG cells are similar to those of HTS-activated TRPV4 currents in other cell types (14, 23, 24).


Figure 1
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  		FIGURE 1.
HTS induces volume changes and Ca2+ entry in human salivary gland cells. A and B, HTS-induced [Ca2+]i increase (increase in the 340/380 nm fluorescence ratio) measured in fura-2-loaded HSG cells. Effects of varying osmolarity are shown in A, and average data from these experiments are presented in B. *, values that are significantly different from those in control conditions, p < 0.01, n = 100–200. HTS at 150 mmol/kg was used in all subsequent experiments. C, effect of HTS on [Ca2+]i in human primary submandibular gland (SMG) and rat basophilic leukemia (RBL) cells to HTS. D, Gd3+ sensitivity of HTS-induced sustained [Ca2+]i increase in HSG cells. 1 or 100 µM Gd3+ was added to the external solution where indicated by arrows, and a control trace is shown by the dashed line. The decrease in fluorescence in the presence of 1 µM Gd3+ was not different from that in control cells; 100 µM Gd3+ rapidly decreased fluorescence to that in resting cells. E, activation of a nonselective cation conductance by HTS in HSG cells. Time courses of the current measured at 80 and –80 mV in cells perfused with Ca2+ + Na+-HTS are shown (see "Experimental Procedures" for details). F, I-V relationship of the current at the time indicated by the arrow in E. G, cation permeability of the HTS-stimulated channel in HSG cells. External Ca2+ + Na+-HTS was replaced by NMDG-HTS solution for the period indicated. H, I-V curves of the current at the time points indicated in G. Current traces are representative of results obtained with a minimum of three cells in each case.

 
We therefore examined the possibility that TRPV4 contributes to HTS-induced Ca2+ entry. 1 µM ruthenium red (RuR), a concentration used for maximum inhibition of TRPV4 currents (23, 25), substantially blocked both inward and outward currents stimulated by HTS (Fig. 2A). Although the current decayed relatively fast under the control conditions (see Fig. 1E), the decay was faster when RuR was added to cells after the current developed (Fig. 2A). Furthermore, washout of RuR induced a significant (p < 0.05) recovery (41 ± 6%, n = 4) of the current to a value not significantly different from the current in control cells. This inhibitory effect of RuR can be more clearly seen in the I-V curves shown in Fig. 2B (inhibition of inward (>90%) and outward (50–60%) currents, p < 0.05, n = 4). Although the outward current in RuR-treated cells was also decreased compared with that in the untreated cells, at membrane potentials >+70 mV, there was a relative increase in the current (Fig. 2B). Thus, the outward current appears to be less sensitive to RuR. There was larger current at the positive membrane potentials relative to that at more negative potentials, consistent with the RuR block being voltage-dependent (23–25). Further, 4{alpha}PDD, an activator of TRPV4 (12, 14, 16, 23–25), induced a sustained increase in [Ca2+]i in Ca2+-containing medium (Fig. 2C) but not in Ca2+-free medium (data not shown), which was attenuated by pre-exposure of the cells to HTS (Fig. 2D). 4{alpha}PDD-induced membrane conductance was similar to that induced by HTS (Fig. 2, E and F). In a number of previous studies, 4{alpha}PDD activation of endogenous TRPV4 generated currents similar to those seen in HSG cells (i.e. relatively more linear and somewhat outwardly rectifying) (14, 23, 26–28). However, in other studies, 4{alpha}PDD stimulation of exogenously expressed TRPV4 generated currents that displayed marked double rectification (12, 14, 23, 29). It should be noted that although HTS and 4{alpha}PDD are efficient activators of TRPV4, they activate the channel via distinct mechanisms (12, 14). Consistent with HTS activation of Ca2+ entry, TRPV4 expression was detected in HSG cells but not in RBL cells (Fig. 2G; membranes from brain and Madin-Darby canine kidney cells were used as controls). In aggregate, the data in Fig. 2 suggest that TRPV4 might be involved in HTS-induced Ca2+ entry in salivary epithelial cells.


Figure 2
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  		FIGURE 2.
Possible role of TRPV4 in hypotonicity-activated Ca2+ entry. A, effect of ruthenium red (included where indicated in A) on the HTS-induced cation current measured at 80 and –80 mV. I-V curves of the current at the indicated time points are shown in B. Amplitude of the current in the presence of RuR was significantly lower (50–60% lower for outward and >90% for inward currents) than in control cells (p < 0.01, n = 4). All current recordings are representative of results obtained from at least four different cells. 4{alpha}PDD activated [Ca2+]i increases (C and D) and cation current as HTS in HSG cells (E and F). G, expression of TRPV4 in HSG and RBL cells; crude membranes from brain and Madin-Darby canine kidney cells (MDCK) were was used as a control. The bands around 97 kDa represent TRPV4.

 


Figure 3
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  		FIGURE 3.
RVD depends on HTS-stimulated Ca2+ entry. A, HTS-induced HSG cell volume changes in HSG cells (measured using Universal Imaging MetaMorph software; representative images acquired at the times indicated by arrows are shown). Other conditions were as described in the legend to Fig. 1. B–D, dependence of RVD on HTS-stimulated Ca2+ entry (volume was determined based on measurements of calcein fluorescence). Traces represent average values obtained from >80 cells; similar results were obtained in three or four separate experiments. Additions and other details are indicated in the figure.

 
Ca2+ Entry Is Required for Regulatory Volume Decrease—The role of Ca2+ entry in the cellular response to HTS was further determined by measuring volume changes. HTS induced an immediate relatively slow increase in cell volume, which was maintained for over 1 min and then decreased to a steady state volume similar to that in resting cells (Fig. 3A; the data calculated based on the area of the cell). Comparable results were obtained using calcein fluorescence measurements (Fig. 3B). The decrease in cell volume but not the initial increase was attenuated by removal of external Ca2+ and/or the addition of 100 µM Gd3+ (Fig. 3, C and D, respectively). These data strongly suggest that Ca2+ entry triggered by HTS regulates RVD. Cell volume regulation is driven by osmotic gradients generated via activation of K+ and Cl– channels (3, 5). In HSG cells, HTS stimulated a charybdotoxin-sensitive KCa channel and an as yet unidentified Cl– channel that was blocked by niflumic acid, a relatively nonspecific blocker of Cl– channels (supplemental Fig. 1). We suggest that the K+ channel is activated via TRPV4-mediated Ca2+ entry and contributes to RVD. The regulation of the Cl– channel is yet to be determined. Additionally, HTS-induced TRPV4 currents were not altered by inclusion of niflumic acid in the bath, suggesting that there is minimal contribution of the Cl– current to the TRPV4 current. Note that K+ currents are abolished under the conditions used to measure TRPV4 currents.

Involvement of TRPV4 in Hypotonicity-stimulated Ca2+ Entry and RVD—To conclusively establish the involvement of TRPV4 in salivary cell volume regulation, we used dispersed submandibular gland acinar cells from TrpV4+/+ and TrpV4–/– mice (18). Robust [Ca2+]i increase was stimulated by HTS in cells from control mice (Fig. 4A). This HTS-induced [Ca2+]i increase was >70% lower in cells isolated from TRPV4 null animals (Fig. 4A; see Fig. 4C for average data). 4{alpha}PDD-induced [Ca2+]i increase was about 20% lower than that seen with HTS in cells from TrpV4+/+ mice but >90% lower in cells from TrpV4–/– animals (Fig. 4B; see Fig. 4C for average values). Similar loss of 4{alpha}PDD-induced TRPV4 activity in cells from TrpV4–/– mice has been previously shown (14, 23). HTS-stimulated cation current was also inhibited >70% (Fig. 4, D–F). Thus, TRPV4 contributes to HTS- and 4{alpha}PDD-induced [Ca2+]i increase and cation currents in submandibular gland acinar cells. Importantly, cells lacking TRPV4 displayed a lack of RVD response in hypotonic conditions; note that cell swelling was not altered (Fig. 4G). A key molecular component required for the RVD response to HTS in salivary acinar cells is AQP5 (7). Fig. 4H shows that TRPV4 expression is lost in submandibular glands from TrpV4–/– mice but that of AQP5 is not altered. These data provide strong evidence that TRPV4-mediated Ca2+ entry is required for AQP5-dependent RVD. The relatively low level of HTS-stimulated, non-TRPV4-dependent, Ca2+ entry detected in TrpV4–/– cells does not support RVD.

Requirement of AQP5 for Activation of TRPV4 by Hypotonicity—The functional association between TRPV4 and AQP5 was further assessed by examining the response of salivary gland acinar cells from Aqp5–/– and Aqp5+/+ mice to HTS (7). A major, and somewhat unexpected, finding was that HTS-stimulated Ca2+ entry was significantly decreased in submandibular gland acini and almost completely abolished in parotid acini isolated from Aqp5–/– mice (Fig. 5, A and B, respectively; see Fig. 5C for average data obtained from parotid gland cells). 4{alpha}PDD-induced Ca2+ entry was about 50% lower than HTS-induced Ca2+ entry in Aqp5+/+ cells and was not significantly different from 4{alpha}PDD-induced Ca2+ entry in Aqp5–/– cells (data from parotid glands are shown in Fig. 5C). Consistent with these findings, HTS-stimulated current was almost completely abolished in parotid acini (Fig. 5, D and E; >90% decrease at –80 and +80 mV, respectively, n = 4). AQP5 was expressed only in glands from Aqp5+/+ mice, whereas TRPV4 expression was similar in control and Aqp5 null mice (Fig. 5F). Thus, changes in TRPV4 expression do not account for the loss of HTS-stimulated Ca2+ entry. Previous studies with parotid and sublingual acini from Aqp5–/– mice showed that the magnitude of HTS-induced swelling was similar to that in Aqp5+/+ cells, although the rate was slower. Importantly, RVD was almost fully inhibited in these cells (7). We noted a similar decrease in the rate (Fig. 5G) but not in the magnitude of swelling (Fig. 5H) in submandibular gland cells from Aqp5–/– mice. Importantly, RVD was significantly decreased in cells lacking AQP5 (Fig. 5I, >80% decrease, p < 0.01). The results of cell volume measurements with parotid cells were similar to those reported earlier (7) and are not shown here. Together these findings suggest that TRPV4-mediated Ca2+ entry regulates RVD in salivary gland acinar cells. More interestingly, these novel findings demonstrate that AQP5 is required for activation of TRPV4 by hypotonicity rather than swelling per se.


Figure 4
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  		FIGURE 4.
Loss of HTS-induced Ca2+ entry and RVD response in salivary gland cells from TRPV4 null mice. Dispersed salivary gland cells were prepared as described under "Experimental Procedures." HTS- and 4{alpha}PDD-induced [Ca2+]i increases in fura-2-loaded submandibular gland acinar cells isolated from TrpV4+/+ and TrpV4–/– mice (A and B, respectively; each trace is an average of at least 20–30 cells; average values from three or four separate cell preparations are shown in C). D and E, membrane current recorded at –80 mV and representative I-V plots of the maximum current generated in each case. F, average current densities. G, HTS-induced cell volume changes. H, expression of TRPV4 (upper blot) and AQP5 (lower blot) in submandibular glands from TRPV4–/– and TRPV4+/+ mice. **, values significantly different from control values in each set (p < 0.01; n = 4–6 for whole cell current measurements and n = 80–100 for Ca2+ imaging experiments).

 
Association of TRPV4 and AQP5 in Salivary Gland Cells—Since AQP5 appeared to determine TRPV4 function, we examined the localization of AQP5 and TRPV4 in mouse submandibular gland. TRPV4 (Fig. 6A, right) was localized in the apical region of mouse submandibular gland acinar cells, as was AQP5 (Fig. 6A, left, also reported previously; see Refs. 30–33). Further, immunoprecipitation with anti-AQP5 antibody pulled down AQP5 with TRPV4 and vice versa (Fig. 6B). AQP3 was present in HSG cells but did not co-immunoprecipitate with TRPV4 (AQP1 and AQP8 were not detected in HSG cells; data not shown). Neither conventional (assessing N- and C-terminal interactions between the two proteins) nor split ubiquitin membrane-based (using the full-length proteins) yeast two-hybrid analysis techniques revealed significant interaction between the two proteins (data not shown). Thus, although TRPV4 and AQP5 are associated with each other, it is likely that they do not directly interact with each other. Alternatively, other regulatory mechanisms govern their interaction under hyposomotic conditions.


Figure 5
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  		FIGURE 5.
Loss of HTS-induced Ca2+ entry and RVD response in salivary gland cells from AQP5 null mice. HTS-induced [Ca2+]i changes in submandibular (A) and parotid acinar cells (B) from Aqp–/– and Aqp+/+ mice (average data are shown in C). The number of cells tested is indicated in the figure. D, HTS-stimulated membrane currents in Aqp5–/– and Aqp5+/+ parotid acinar cells recorded at –80 mV; I-V curves of the maximum current are shown in E. The current recordings represent results obtained with at least four cells in each case. F, expression of TRPV4 (upper blot) and AQP5 (lower blot) in submandibular (SG) and parotid (P) glands from Aqp5–/– and Aqp5+/+ mice. Changes in cell volume in response to HTS were measured in calcein-loaded submandibular gland cells from AQP5–/– and Aqp+/+ mice. G, t1/2 of cell swelling (time required for half-maximal swelling); H, percentage increase in cell volume; I, extent of cell volume recovery. **, values that are significantly different from unmarked values (p < 0.05; n = 29–34 for imaging experiments).

 
Effect of N- or C-terminal Deletion of AQP5 on Hypotonicity-induced TRPV4 Activation and RVD—To further characterize the regulation of TRPV4 by AQP5, we utilized constructs of AQP5 in which either the C or N terminus of AQP5 has been deleted (34). Expression of N terminus-deleted AQP5 (Fig. 6, D–F) but not wild type AQP5 or C terminus-deleted AQP5 (Fig. 6C) blocked HTS-induced but not 4{alpha}PDD- or thapsigargin-induced (data not shown) Ca2+ entry and HTS-stimulated cation currents. Importantly, as seen in cells lacking AQP5, the extent of HTS-induced swelling was not altered, although RVD was significantly reduced (Fig. 6D). C-terminal deletion of AQP5 has been shown to disrupt its trafficking to the apical membrane, resulting in retention of the protein in the cell, whereas N-terminal deletion does not affect its intracellular targeting (34). Based on the effects of these mutant AQP5 proteins on the response of cells to HTS, it seems likely that AQP5 lacking the C terminus does not exert any effect, since it does not interact with endogenous AQP5 or reach the apical membrane. The N terminus-deleted AQP5, on the other hand, induces a dominant negative effect. We suggest that this protein can interact with the endogenous AQP5 but does not form a functional channel. This further indicates that the N terminus of AQP5 might not be involved in homomeric interactions of AQP5. Additionally, the AQP5 N terminus might be involved in the HTS regulation of TRPV4. Whereas the mechanism by which the aquaporin mutant suppresses HTS responses has to be further established, these data strongly suggest that TRPV4 and AQP5 are associated functionally and physically (although probably via indirect interactions). These data also reveal a novel regulation of TRPV4 (i.e. that channel activation by hypotonicity is dependent on the presence of functional AQP5). These data are also consistent with previous suggestions that TRPV4 is not activated by cell swelling per se (16).


Figure 6
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  		FIGURE 6.
Association and functional interaction between TRPV4 with AQP5. A, localization of AQP5 (left) and TRPV4 (right; magnified image included) in serial sections of mouse submandibular glands (reactivity was not detected in the absence of primary antibody; image not shown). B, IP with anti-TRPV4 antibody (second panel; IB antibodies indicated below the blots) or AQP5 antibody (third panel; inputs shown in the first panel). IP with TRPV4 does not pull down AQP3 (first lane, IP fraction; second lane, unbound fraction). Shown is the HTS-induced increase of [Ca2+]i in HSG cells transiently expressing AQP5-wt or AQP5-{Delta}C (C) or AQP5-{Delta}N (D). The effect of AQP5-{Delta}N expression on HTS-activated cation current (E) and on HTS-induced volume changes (F) is shown (average trace measured in 20–30 cells). 4{alpha}PDD was included in the external solution where indicated.

 
Involvement of Cytoskeleton in Activation of TRPV4 and Regulatory Volume Decrease—AQP5-mediated regulation of cellular volume during agonist stimulation of salivary gland cells has been associated with translocation of the water channel protein to the plasma membrane (31, 32). This recruitment depends on [Ca2+]i increase and is blocked by depolymerization of actin with cytochalasin D (cyto-D). Additionally, TRPV4 has also been shown to interact with the cytoskeleton (35). Thus, we examined the effect of depolymerization of the cytoskeleton on the responses of HSG cells to HTS. HTS-but not 4{alpha}PDD-stimulated Ca2+ entry in HSG cells was blocked by actin depolymerization (Fig. 7A). Interestingly, 4{alpha}PDD induced a transient activation of Ca2+ entry (compare Fig. 7A with Fig. 2C) and cation currents (compare Fig. 7B with Fig. 2E). Furthermore, HTS-induced swelling of HSG cells was not affected by cyto-D, but RVD was blocked (Fig. 7C), and the addition of 4{alpha}PDD to the swollen cells induced a small transient decrease in cell volume consistent with the transient Ca2+ entry.

Effect of Hypotonicity on Cell Surface Expression and Association of TRPV4 and AQP5—Fig. 7, D–F, shows the effect of HTS on the surface expression of TRPV4 and AQP5 in control and cyto-D-treated HSG cells. HTS induced a marked increase in the level of AQP5 and TRPV4 in the biotinylated fraction, which was attenuated in cells treated with cyto-D (Fig. 7D; plasma membrane Ca2+ pump in this fraction was not changed; Supplemental Fig. 2). Additionally, HTS increased co-immunoprecipitation of TRPV4 with AQP5, which was also blocked by cyto-D pretreatment of cells (Fig. 7E, left upper blots). Since anti-AQP5 was used to pull down AQP5, the level of AQP5 is similar in both samples (right panels, which also indicate similar protein load; note that AQP5 and TRPV4 were detected using the same blot). Input levels of TRPV4 and AQP5 in lysates of control and cyto-D-treated cells are shown in Fig. 7F (note that these lysates were used for IP with either avidin or anti-AQP5 (i.e. blots shown in Fig. 7, D and E) and thus serve as a control for both). The increased association of TRPV4 and AQP5 was verified using dispersed mouse submandibular gland cells. HTS induced a similar increase in the co-immunoprecipitation of TRPV4 with AQP5 in these cells (Fig. 7G; IP was done using anti-AQP5 antibody; blots show TRPV4 and AQP5 in IP samples in the top panels and cell lysates, input, in the lower panels). An increase was seen only in the levels of TRPV4 in HTS-treated cells. Thus, the effects of HTS and cyto-D on the association of TRPV4 with AQP5 are not due to differences in protein load. In aggregate, these data suggest that HTS (i) increases the association between TRPV4 and AQP5 and (ii) regulates plasma membrane trafficking/insertion of these channels. Cytoskeletal rearrangements appear to be involved in regulating both of these events. It is important to note that both AQP5 and TRPV4 have been suggested to be regulated via interactions with the cytoskeleton (31, 32, 34, 35). Further studies are required to determine whether the two proteins associate with each other prior to translocation to the plasma membrane or after they are individually trafficked to the surface membrane.


Figure 7
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  		FIGURE 7.
HTS-induced, cytoskeleton-dependent association and surface expression of TRPV4 and AQP5 regulates RVD. Effect of cytochalasin D treatment (CytoD; 1 µM for 30 min) on HTS-induced [Ca2+]i increase (A), cation current (B), and regulation of cell volume (C). Changes in the external medium and time (horizontal bar) are indicated in the figure. D–G, effect of HTS on surface expression and association of TRPV4 and AQP5 in HSG (D–F) and freshly dispersed submandibular gland cells (G). Cells were treated with cytochalasin D as described above. Control and cyto-D cells were then exposed to HTS and biotinylated on ice. Cell lysates from control and HTS-treated cells were incubated with either avidin (D) or anti-AQP5 (E), and the blots were probed with anti-TRPV4 or anti-AQP5 antibodies as indicated (TRPV4 and AQP5) in HSG cell lysates (F). G, effect of HTS on co-IP of TRPV4 and AQP5 from dispersed mouse submandibular glands. Cells were prepared as described under "Experimental Procedures." Cyto-D treatment and other experimental conditions were similar to those used for HSG cells. Data are representative of results obtained in three independent experiments.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
The data presented above provide evidence that TRPV4 and AQP5 concertedly control RVD in salivary epithelial cells. Hypotonic conditions induce cell swelling and activation of Ca2+ entry via TRPV4. Subsequent activation of RVD is dependent on this Ca2+ entry as well as AQP5, which regulates water permeability in these cells. Importantly, we demonstrate that TRPV4 activation by HTS is dependent on the presence of functional AQP5. TRPV4 is not activated by HTS in cells lacking AQP5, although cells undergo swelling of a similar magnitude as, albeit at a slower rate than, that in control cells. The slower rate of swelling probably reflects a lack of contribution by AQP5 to plasma membrane water permeability. Further, we also show that HTS stimulates the association and plasma membrane trafficking of both of these channels. Conditions that block these processes also block RVD. Thus, our data reveal for the first time an important functional link between TRPV4 and the water channel AQP5. Despite a number of recent studies that demonstrate the involvement of TRPV4 in cell volume regulation and mechanosensation (10–18), the molecular basis for the regulation of TRPV4 has not yet been established. Further, the exact mechanisms by which AQP5 is regulated are also unclear (5). Our data provide evidence of a novel association between TRPV4 and AQP5 that is involved in activation of TRPV4 by hypotonicity and regulation of cellular response to the osmotic stress. We suggest that TRPV4 and AQP5 are assembled in a signaling complex that responds to anisosmotic conditions and coordinates cellular volume regulation.

Our data also suggest that TRPV4 can have a critical role in salivary gland fluid secretion. A fundamental role for AQP5 in regulation of salivary gland fluid secretion has been shown earlier (7). Aqp5–/– mice display pronounced decreases in salivary fluid secretion, and salivary gland cells from these mice exhibit diminished RVD response to HTS (5, 7). Thus, it was suggested that AQP5 regulates salivary secretion by controlling the water permeability across salivary acinar cell apical membrane. However, the molecular basis of RVD in salivary gland and other cell types is not yet clear. Although it has been long recognized that regulatory volume changes are determined via regulation of [Ca2+]i, the mechanisms involved in sensing and transducing signals related to changes in the osmolarity of the cell medium have not yet been determined. Our data clearly demonstrate that, like AQP5, TRPV4 has a central role in regulating RVD in salivary epithelial cells. TRPV4 channels are activated in response to hypotonicity in freshly dispersed salivary gland cells and salivary cell lines. Further, the Ca2+ entry via TRPV4 is required for activation of a Ca2+-activated K+ channel and subsequent RVD. Importantly, we show that dispersed salivary gland cells from TrpV4–/– mice (18) display decreased responses to HTS; Ca2+ entry is greatly reduced, and the ability to regulate cell volume after swelling is lost. These data demonstrate that TRPV4 contributes to HTS-activated Ca2+ entry and regulates RVD via activation of the K+ channel. Together with an as yet unknown Cl– channel, these K+ and Cl– fluxes generate the osmotic gradient that drives water flow, via AQP5, to induce shrinkage of cells to their original volume. Thus, TRPV4 has a critical role in the regulation of RVD. Since regulation of transepithelial osmotic forces as well as cell volume critically impacts salivary gland fluid secretion, TRPV4 is likely to have a central role in regulating salivary gland function. This remains to be confirmed, since salivary secretion could not be measured reliably in TrpV4–/– mice due to the sensitivity of the mice to general anesthesia.3

A pivotal finding of the present study is that hypotonicity-induced activation of TRPV4 is dependent on the presence of functional AQP5 and not on the osmolarity of the medium or magnitude of cell swelling per se. Further, we show that the association between AQP5 and TRPV4, as well as their surface expression, is increased upon exposure of the cells to hypotonic conditions. Although the molecular scaffold mediating these events is not yet known, based on our data, we suggest that sustained RVD is mediated by a cytoskeleton-dependent increase in the association and cell surface expression of AQP5 and TRPV4. This suggestion does not exclude the possibility that other HTS-activated signals, such as arachidonic acid metabolites or protein kinase (12, 36), could contribute to this channel regulation. Consistent with our findings, Arniges et al. (16) previously reported that reduced cell swelling does not account for lack of TRPV4 activation and loss of RVD in airway epithelial cells exogenously expressing a mutant cystic fibrosis membrane regulator. Cystic fibrosis membrane regulator has been known to interact with several other ion transport proteins and regulate fluid secretion. Cystic fibrosis airway epithelia have defective swelling-activated K+ and Cl– channel activities and therefore display loss of RVD response (16). Although the role of aquaporins in cystic fibrosis membrane regulator-dependent regulation of cell volume has not yet been described, it has been suggested that alveolar aquaporins might be more involved in cell volume regulation than transepithelial fluid flux (6). In this regard, it is interesting that AQP5 and cystic fibrosis membrane regulator appear to be colocalized apically in pancreatic ductal cells and salivary epithelial cells (37). TRPV4 is also found in the apical region of salivary gland cells (30–33). Recently, TRPV4 has been reported to form a functional signaling complex with large conductance Ca2+-activated K+ channels that are involved in vasodilation (27). Additionally, and consistent with our data, TRPV4 has been reported to interact with MAP-7 via its C terminus; the interaction links the channel to the cytoskeleton and regulates surface expression of the channel (35). Together our present data and these previous studies suggest that TRPV4 and AQP5 are components of a larger signaling platform that can not only sense osmotic and mechanical signals but also transduce these signals and coordinate the regulation of key ion channels that are involved in controlling cell shape and volume recovery. It is also interesting that TRPV4, like TRPV1 and TRPV2, is translocated to the membrane in response to the stimulus. Thus, regulated trafficking appears to be a common mechanism underlying regulation of TRPV channels by specific sensory signals (38–41).

In conclusion, our data demonstrate a role for AQP5 in the activation of TRPV4 by hypotonicity in salivary gland cells. Further, we suggest that AQP5 and TRPV4 are concertedly involved in regulation of cell volume and salivary gland fluid secretion. AQP5 also mediates fluid secretion in mucosal glands and epithelial cells in the lung as well as in cells from pancreas and sweat glands (6, 7). TRPV4 has been suggested to regulate RVD in airway epithelial cells, although the functional interaction between TRPV4 and AQP5-mediated volume changes in these tissues remains to be clarified. Thus, based on the role of TRPV4 in osmosensation and cell volume regulation demonstrated here and previously (16–18, 27), it can be suggested that TRPV4 can function as a transducer of osmotic stimuli and in concert with AQP5 contribute to the regulation of cellular volume. Further studies are required to determine the critical molecular components that are involved in detection of these signals and coordination of the cellular responses.


   FOOTNOTES
 
* This work was supported in part by National Institutes of Health Grant DE013823 (to Anil Menon, University of Cincinnati, and J. E. M.). 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. Back

Formula The on-line version of this article (available at http://www.jbc.org) contains supplemental Figs. 1 and 2. Back

1 To whom correspondence should be addressed: Secretory Physiology Section, Gene Therapy and Therapeutics Branch, NIDCR, Bldg. 10, Rm. 1N-113, National Institutes of Health, Bethesda, MD 20892. Tel.: 301-496-1478; Fax: 301-402-1228; E-mail: indu.ambudkar{at}nih.gov.

2 The abbreviations used are: RVD, regulatory volume decrease; HTS, hypotonic solution; NMDG, N-methyl-D-glucamine, 4{alpha}PDD, 4{alpha}-phorbol-12,13-didecanoate; IP, immunoprecipitation; cyt-D, cytochalasin D; RuR, ruthenium red. Back

3 W. Liedtke, unpublished observations. Back


   ACKNOWLEDGMENTS
 
We thank Drs. Michael Caterina and Jon Levine for generously providing the TRPV4 antibody, Drs. Bruce Baum, Robert Wellner, and Anil Menon for anti-AQP5 antibodies and AQP5 plasmids, and David Wang for invaluable help with immunocytochemistry.



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 ABSTRACT
 INTRODUCTION
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 RESULTS
 DISCUSSION
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