TRPM7 Is Involved in Volume Regulation in Salivary Glands

Introduction

Regulatory volume decrease (RVD) is a dynamic cellular process in response to extracellular osmotic (hypotonic) changes. The volumetric regulation mechanism is essentially required for maintenance of homeostasis in biological systems and for induction of biological functions such as proliferation, migration, and differentiation (Okada et al. 2001; Dubois and Rouzaire-Dubois 2012). The RVD response following osmotically induced cell swelling has been ubiquitously reported in a variety of cell types (Hoffmann et al. 2009). The major proportion of the net solute loss during this response is accounted for by the KCI efflux, which is mediated by the activation of separate K⁺ and Cl⁻ channels, with an additional loss of organic osmolytes through a volume-sensitive anion channel (Strange et al. 1996; Okada 1997). In addition to KCl transport, an increase in intracellular Ca²⁺ ([Ca²⁺]_i) has been observed during the volumetric changes of cells under hypotonic stress (McCarty and O’Neil 1991; Negulescu et al. 1992; Wu et al. 1997; Urbach et al. 1999). The ubiquity of the swelling-evoked [Ca²⁺]_i increase makes Ca²⁺ a plausible mediator for an osmo-transducing signal. It has been reported that RVD under osmotic perturbation is blocked by extracellular or intracellular Ca²⁺ chelation (Montrose-Rafizadeh and Guggino 1991; Pan et al. 2008), implying that Ca²⁺ from diverse routes might be involved in the volumetric regulatory process. However, in a few cell types, the RVD response was apparently not associated with any change of [Ca²⁺]_i (Grinstein and Smith 1990; Jorgensen et al. 1997).

In salivary glands, the acinar cells are considered to equip a well-developed volume regulatory system since the cell volume is frequently changed by an imbalance between the basolateral and apical membrane fluxes during saliva secretion (Melvin et al. 2005). Moreover, several stimulants of saliva secretion are known to cause a rapid increase in [Ca²⁺]_i in a single acinar cell, which is followed by rapid and substantial cell shrinkage (Foskett and Melvin 1989; Nakahari et al. 1990). Although such evidence demonstrates the functional relationship between RVD and Ca²⁺ in salivary acinar cells, the precise molecular mechanism of Ca²⁺ in this process remains unclear. Therefore, this study investigated the role of Ca²⁺ in the RVD process upon anisomotic conditions using submandibular gland acinar cells.

Materials and Methods

Results

Regulatory Volume Decrease and [Ca²⁺]_i Changes in Hypotonic Solutions

We first monitored the volumetric changes of SMG acinar cells upon hypotonic stress and evaluated the patterns after alteration of intracellular Ca²⁺ concentration. Acinar cells cultured from rat SMGs were exposed to hypotonic solution with 74% osmolarity compared with isotonic solution (220 mOSm; see Materials and Methods), because lower osmolarity might induce unwanted damage to the cells. After exposure to the hypotonic solution, the cells swelled to a maximal volume ratio (V_max) of 1.15 ± 0.03 (means ± SE) within 100 s. An RVD was observed over the subsequent 500 s. The minimal cell volume ratio (V_min) at the end of 10-min hypotonic exposure was 1.08 ± 0.02, a significant decrease from the V_max (P < 0.05), indicating a typical RVD response (Fig. 1A). We next investigated the effect of BAPTA-AM (intracellular Ca²⁺ chelator) on the RVD. Cells that were pretreated with a high concentration (20 µM) of BAPTA-AM also swelled after hypotonic changes, but the RVD was almost completely inhibited (V_max: 1.12 ± 0.02 and V_min: 1.10 ± 0.02; P > 0.1) (Fig. 1B). These data clearly showed the RVD in SMG acinar cells upon hypotonic stress and the involvement of intracellular Ca²⁺ in the RVD process (Fig. 1C). We next investigated the actual Ca²⁺ response upon osmotic perturbation by recording the changes in [Ca²⁺]_i induced by cell swelling under 250-, 220-, and 190-mOsm hypotonic solutions (Fig. 1D). The magnitude of the [Ca²⁺]_i increase was proportional to the degree of hypotonicity: the magnitude of the [Ca²⁺]_i increase was highest in the 190-mOsm hypotonic solution (F₃₄₀/F₃₆₀ = 1.96 ± 0.12 equivalent to [Ca²⁺]_i = 262 nM) and lowest in the 250-mOsm solution ([Ca²⁺]_i = 54 nM) (Fig. 1E). The time lag of the rise in [Ca²⁺]_i evoked by hypotonic stress was also correlated with the degree of hypotonicity: the mean time lags were 95.1, 102.9, and 203 s with the 190-, 220-, and 250-mOsm hypotonic solutions, respectively. The incidence of the [Ca²⁺]_i response in the cell population was highest in the most dilute solution of 190 mOsm and lowest in the 250-mOsm hypotonic solution (Fig. 1F).

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Figure 1.

Regulatory volume decrease (RVD) and related [Ca²⁺]_i changes of submandibular gland (SMG) acinar cells in hypotonic solutions. (A) Volume changes in the cells after hypotonic changes (n = 7). (B) Volume changes in cells preincubated with 20 µM BAPTA-AM after hypotonic changes (n = 6). V_max: 1.12 ± 0.02 and V_min: 1.10 ± 0.02 (P > 0.05). (C) Relative changes of V_min/V_max. *P < 0.05. (D) Fura-2–based superimposed traces of the [Ca²⁺]_i increase induced by exposure to 3 different hypotonic solutions (250-, 220-, and 190-mOsm hypotonicity). (E) Magnitude of the [Ca²⁺]_i increase induced by hypotonic change. The maximum [Ca²⁺]_i increase was 1.05 (54 nM, n = 1), 1.25 ± 0.09 (89 nM, n = 6), and 1.96 ± 0.12 (262 nM, n = 10) in 250-, 220-, and 190-mOsm hypotonic solutions, respectively. (F) The incidence of cells showing [Ca²⁺]_i increase induced by hypotonic change. n = 6, 13, and 17 in 250-, 220-, and 190-mOsm hypotonic solutions, respectively.

Coupling of [Ca²⁺]_i and Cell Volume in Hypotonic Condition

We next investigated the direct relationship between [Ca²⁺]_i and RVD by simultaneously recording cell volume and [Ca²⁺]_i in Fura-2–loaded cells during exposure to a 190-mOsm hypotonic solution. We first confirmed that the experimental loading condition of Fura-2 did not perturb RVD response (V_max: 1.21 ± 0.04 and V_min: 1.12 ± 0.03; P < 0.05) (Fig. 2A). To determine whether Ca²⁺ mediates the RVD, the temporal relationship between the [Ca²⁺]_i and the RVD was investigated. A [Ca²⁺]_i peak was always observed between the initiation time point of volume increase and that of RVD (in 10 experimental trials), and it required 95.1 ± 13.6 s to reach the peak [Ca²⁺]_i and 150 ± 17.3 s to reach the V_max for RVD initiation (P < 0.05). The result of simultaneous monitoring of changes in cell volume and [Ca²⁺]_i showed an obvious temporal sequence between the parameters: cell volume increase–Ca²⁺ response–RVD (Fig. 2B). Because the incidence of Ca²⁺ response was 58.82% (10 from total 17 cells) upon 190-mOsm hypotonic stress (Fig. 1F), we compared the rates of cell swelling between the Ca²⁺ response and nonresponse groups. Cells in the Ca²⁺ response group showed a significantly faster swelling rate than those in the nonresponse group (0.22 ± 0.03 [Ca²⁺ response] and 0.08 ± 0.02 unit/min [nonresponse]; P < 0.05), indicating the volume-sensitive [Ca²⁺]_i increase (Fig. 2C). Such different reactivity to the hypotonic solution might be due to a mixed cell population including serous, mucous acinar, and granular convoluted tubule (GCT) cells in the primarily cultured SMG. Next, cells were treated with 2 µM ionomycin (Ca²⁺ ionophore) to verify the direct effect of Ca²⁺ on volume reduction. The [Ca²⁺]_i began to increase 2 min after addition of ionomycin, followed by a significant reduction in the cell volume (P < 0.05; Fig. 2D). These data demonstrated that Ca²⁺ is a crucial mediator that links hypotonic cell swelling and RVD response.

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Figure 2.

Coupling of [Ca²⁺]_i and cell volume in a hypotonic condition. (A) Regulatory volume decrease (RVD) of Fura-2–loaded submandibular gland (SMG) acinar cells in a 220-mOsm hypotonic solution (n = 8). V_max: 1.21 ± 0.04 and V_min: 1.12 ± 0.03 (P < 0.05). (B) Simultaneous recording of the cell volume (dark circles) and [Ca²⁺]_i (solid line) in a 190-mOsm hypotonic solution. (C) Cell swelling rate in the [Ca²⁺]_i response group (dark circles, n = 10) and the nonresponse group (white circles, n = 7) in the 190-mOsm hypotonic solution. Dotted lines indicate trend lines. (D) Effects of 2 µM ionomycin on [Ca²⁺]_i (solid line) and cell volume (dark circles) in a normal bath solution (n = 6). *P < 0.05. **P < 0.01.

Identification of the Route of Ca²⁺ Response upon Hypotonic Stress

We next performed mechanistic studies of the Ca²⁺ reaction induced by hypotonic stress. To this end, we used a genetically encoded intensiometric Ca²⁺ biosensor (GCaMP6s) with excellent sensitivity and proper kinetics for detecting slow Ca²⁺ signals (Chen et al. 2013). A rat SMG epithelial cell line (SMG-C6) was selected for efficient expression of the biosensor. Upon hypotonic stress, GCaMP6s signals were clearly increased, with 2.5-fold higher magnitude than a Fura-2 system (relative peak intensity: 4.89 ± 0.59; Fig. 3A and Appendix Movie 1). The tracking data of the [Ca²⁺]_i revealed 2 types of Ca²⁺ signals: an initial increase of brief Ca²⁺ patterns and subsequently evoked elongated patterns, reminiscent of Ca²⁺ influx via channels and Ca²⁺ efflux from intracellular stores, respectively (Fig. 3B). This combined (brief + elongated) Ca²⁺ pattern is a major response upon hypotonic stress: 65% of cells showed this pattern, and a minor population of the cells showed a repeated brief (30%) or single elongated (5%) pattern (Appendix Fig. 1). We hypothesized that these signals compose the Ca²⁺-induced Ca²⁺ release (CICR) process and therefore applied various chemical inhibitors of CICR components. Depletion of intracellular Ca²⁺ stores by thapsigargin treatment almost completely abolished the Ca²⁺ response upon hypotonic stress, and the Ca²⁺ efflux was also significantly reduced by treatment with ryanodine (ryanodine receptor [RyR] inhibitor; relative peak intensity: 2.20 ± 0.37; P < 0.01) but not with U73122 (phospholipase C-γ [PLCγ] inhibitor) (Fig. 3C). The involvement of RyR in this response suggested that the [Ca²⁺]_i increase is mediated by CICR. We presumed that stretch-activated Ca²⁺ channels (SACCs) trigger this reaction, based on the finding that SACCs are involved in the Ca²⁺ reaction upon anisosmotic cell volume changes (Hoffmann et al. 2009). As expected, blocking SACCs by GdCl₃ treatment clearly attenuated the Ca²⁺ response (relative peak intensity: 1.27 ± 0.05; P < 0.001; Fig. 3C). We next investigated the messenger RNA (mRNA) expression profile of mammalian SACCs (Wei et al. 2009) and found dominant expression of transient receptor potential melastatin 7 (TRPM7) in SMG-C6 cells and primary SMG acinar cells (Fig. 3D and Appendix Fig. 2A). The expression of TRPM7 was verified at a protein level by immunocyto- and immunohistochemistry (Fig. 3E and Appendix Fig. 2B). Interestingly, TRPM7 expression was dominantly detected in serous acini (specifically labeled by CD44 [Maria et al. 2012]; Appendix Fig. 2B), suggesting the possibility that serous acinus mainly causes positive Ca²⁺ response upon hypotonic stress. Moreover, the hypotonic-induced [Ca²⁺]_i increase was clearly reduced by treatment with a specific TRPM7 inhibitor, FTY720, indicating that TRPM7 channels act as a crucial mediator for initiating CICR related to RVD response (Fig. 3F).

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Figure 3.

Identification of the route of Ca²⁺ response upon hypotonic stress. (A) Representative images of GCaMP6s-expressing submandibular gland (SMG)–C6 cells in isotonic (left) and hypotonic (right) solution. (B) GCaMP6 intensity changes of the cells in panel A after hypotonic changes (arrows). White and black arrowheads indicate brief and elongated Ca²⁺ transients, respectively. A.U., arbitrary unit. (C) Relative peak intensities of GCaMP6s induced by hypotonic stress in cells pretreated with 1 µM thapsigargin (Tg, sarco/endoplasmic reticulum Ca²⁺-ATPase inhibitor), 10 µM U73122 (phospholipase C-γ inhibitor), 50 µM ryanodine (ryanodine receptor inhibitor), and 500 mM GdCl₃ (Gd, stretch-activated Ca²⁺ channel inhibitor). n = 16 (control), 15 (Tg), 16 (U73122), 13 (ryanodine), and 16 (Gd). Analysis of variance, P = 2.73 × 10^–13, **P < 0.01, ***P < 0.001. (D) Messenger RNA (mRNA) expression profile of stretch-activated Ca²⁺ channels in SMG-C6 cells. P: transient receptor potential polycystic; C: transient receptor potential canonical; V: transient receptor potential vanilloid; M: transient receptor potential melastatin. (E) Immunolabeling of transient receptor potential melastatin 7 (TRPM7) in SMG-C6 cells. (F) Relative peak intensities of GCaMP6 (SMG-C6 cells, black) and Fura-2 (primary rat SMG acinar cells, white) induced by hypotonic stress in cells pretreated with 1 µM FTY720 (TRPM7 inhibitor). Data are presented as median ± maximum/minimum. ***P < 0.001. Scale bars: 20 µm.

Ca²⁺-Activated Cl⁻ Conductance Is Increased by Hypotonic Stress

We examined whether Ca²⁺-dependent conductance could be generated during hypotonic stress. Using a conventional whole-cell configuration, a gradual increase in the Cl⁻ current was recorded in isolated SMG acinar cells by superfusion with a 220-mOsm hypotonic solution (Fig. 4A). At the peak of the Cl⁻ current, addition of 100 µM 5-nitro-2-(3-phenylpropylamino)benzoic acid (NPPB, an inhibitor of calcium-sensitive chloride currents) partially inhibited the increasing currents, and the remnant currents were further decreased after the return to isotonic condition (Fig. 4A). The current profile showed a marked outward rectification with voltage and time dependence (Fig. 4B). It was slowly activated by the depolarizing voltages and rapidly inactivated by the hyperpolarizing voltages, which is a characteristic of the Ca²⁺-activated Cl⁻ channels in lacrimal and salivary acinar cells (Park and Brown 1995; Arreola et al. 1996). The mean current-voltage (I-V) relationship of these currents was recorded with a reversal potential of −10.3 ± 0.3 mV, which is very close to the calculated E_Cl (–9.8 mV) (Fig. 4C). Based on these results, we pretreated cells with NPPB under a hypotonic condition to confirm its inhibitory effect on the RVD. In the presence of 100 µM NPPB, the V_max in the hypotonic solution was 1.38 ± 0.07, while the V_min was 1.38 ± 0.09, which were not significantly different (P > 0.1; Fig. 4D), indicating that the Ca²⁺-induced Cl⁻ current functionally mediates the RVD response.

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Figure 4.

Ca²⁺-activated Cl⁻ conductance is increased by hypotonic stress. (A) Swelling-activated Cl⁻ currents evoked by exposure to a 220-mOsm hypotonic solution. The inset on the upper right side shows a voltage pulse protocol. (B) Current profiles recorded at the time indicated by * in panel A. The pulse protocol involved a −40-mV holding potential with 20-mV step potentials applied from −100 mV to +100 mV. (C) Mean I-V relationship of the volume-activated Cl⁻ conductance. (D) Effects of the Ca²⁺-activated Cl⁻ channel inhibitor (5-nitro-2-(3-phenylpropylamino)benzoic acid [NPPB]) on the regulatory volume decrease response. V_max: 1.39 ± 0.06 and V_min: 1.39 ± 0.09 (P > 0.05). n = 6.

Discussion

Through this work, we sought to reveal the role of Ca²⁺ in the RVD response upon hypo-osmotic perturbation. Our experiments show that a typical RVD response takes place with a concurrent increase in [Ca²⁺]_i in rat SMG acinar cells. We found that intracellular Ca²⁺ mobilization was mediated by a CICR process, and the magnitude and incidence of [Ca²⁺]_i increase were closely related to the volume changes of acinar cells. A previous study reported that Ca²⁺ influx was a prerequisite for initiating the RVD response in the lacrimal gland, but the cells did not show a remarkable change in [Ca²⁺]_i during hypotonic exposure (Speake et al. 1998). In contrast, Ca²⁺ efflux from intracellular stores mainly occurred during RVD in renal cells (Tinel et al. 2000). This study suggests a more complex interaction between Ca²⁺ influx and efflux during the RVD response in which inward Ca²⁺ signals are amplified via RyR-mediated intracellular Ca²⁺ release, known as the CICR process. These results reflect the context-dependent role of Ca²⁺ in the RVD according to different biological systems. However, inhibition of RyR could not totally abolish the intracellular Ca²⁺ response (Fig. 3C), implying that another route for Ca²⁺ efflux might also be involved in this reaction. It can be assumed that inositol trisphosphate receptor partially mediates the Ca²⁺ reaction through phospholipase activity, as application of PLCγ inhibitor slightly reduced the [Ca²⁺]_i increase upon hypotonic stress (Fig. 3C).

We applied immortalized submandibular gland acinar cells (SMG-C6 cells) to dissect the origin of Ca²⁺ transients by applying a genetically encoded Ca²⁺ indicator (GECI). A SMG-C6 cell line was generated from primarily cultured acinar cells from rat SMGs by transduction of a replication-defective simian virus (SV40) genome, and initial validation demonstrated that the cell line possessed diverse signaling properties of acinar origin (Quissell et al. 1997). Therefore, this cell line has been widely used as an in vitro model for studying SMG physiology. Corresponding to our results showing Cl⁻ conductance in primary acinar cells (Fig. 4B, C), a previous characterization study reported that SMG-C6 cells also showed Ca²⁺-induced Cl⁻ conductance (Castro et al. 2000). Moreover, the functionality of RyR, one of the key components in the CICR mechanism, was reported in SMG-C6 cells in accordance with RyR expression in parotid acinar cells (Zhang et al. 1997; Liu et al. 2000). This body of evidence clearly demonstrates the relevance of using an SMG-C6 cell line for this work.

Using this model, we found that TRPM7, a stretch-activated channel, is a crucial component triggering the CICR. To date, the 8 types of TRP channels in mammalian cells are known to be SACCs (Nilius et al. 2007). Among these channels, transient receptor potential vanilloid 4 (TRPV4) is widely known for its role in swelling-mediated Ca²⁺ transduction in diverse biological systems such as tracheal, renal, and corneal epithelial cells (Arniges et al. 2004; Cohen 2005; Pan et al. 2008). In salivary acinar cells, TRPV4 was reported to be associated with aquaporin-5 for volume regulation, implying that multiple regulatory mechanisms might be involved in the acinar cell RVD process (Liu et al. 2006). TRPM7 is a mechanosensitive nonselective cation channel (Nilius et al. 2007), and its role in volume regulation is just beginning to be understood. In this study, we demonstrated the TRPM7-mediated Ca²⁺ response under hypotonic conditions by profiling mRNA expression and applying a specific inhibitor, thereby elucidating a previously unknown role of TRPM7 in the volume regulatory process in SMG acinar cells. It is noteworthy that TRPV4 inhibitor did not show a remarkable effect on [Ca²⁺]_i changes upon hypotonic stress (data not shown).

We also provided additional evidence for the role of [Ca²⁺]_i in the RVD through patch-clamp experiments. The current profile induced by hypotonic stress in acinar cells was different from that of the volume-activated Cl⁻ channels. The electrophysiological characteristic of the volume-activated Cl⁻ channels in the salivary acinar cells is that membrane depolarization beyond +60 mV inactivates the conductance in a time- and voltage-dependent manner (Arreola et al. 1995; Majid et al. 2001). It has been well documented that activation of the Ca²⁺-activated Cl⁻ and Ca²⁺-activated K⁺ channels is osmotically induced by cell swelling in lacrimal cells (Kotera and Brown 1993). Our findings that NPPB inhibited not only channel activation but also the RVD strongly suggest that Cl⁻ conductance, which is activated by Ca²⁺, plays a putative role in the RVD.

However, it is unlikely that the Ca²⁺-activated channels are the only channels involved in the RVD in SMG acinar cells. It was reported that 3 types of Cl⁻ channels were expressed simultaneously in rat salivary acinar cells, and each type could be activated independently by intracellular Ca²⁺, cell volume, and membrane potential (Arreola et al. 1996). It seems that the Cl⁻ channel type involved in RVD depends on the [Ca²⁺]_i buffering capacity of the cell. When the intracellular Ca²⁺ buffering capacity is normal, the Ca²⁺-activated Cl⁻ channel plays a role in initiating the RVD. When the Ca²⁺ buffering capacity is high (e.g., application of BAPTA at high concentration), a volume-activated Cl⁻ channel might play a role (Majid et al. 2001). The possibility that the role of the channels throughout the RVD process can be switched according to physiological conditions also cannot be ruled out. For example, with cell swelling, the Ca²⁺-activated Cl⁻ channels are first activated by a rise in the [Ca²⁺]_i. When the [Ca²⁺]_i falls below the threshold level, volume-activated Cl⁻ channels, which are independent of [Ca²⁺]_i, take over the role in the RVD. Chloride channel (ClC)–3, a representative swelling- and Ca²⁺-activated Cl⁻ channel in mammals, has been proposed to be a possible mediator in cell volume regulation. However, ClC-3–deficient mice presented normal RVD responses and Cl⁻ currents equivalent to wild-type mice (Arreola et al. 2002). Therefore, the remaining question is to identify the core mediator of RVD among Cl⁻ channel subfamilies.

In conclusion, our findings suggest the precise signaling process in SMG acinar cells from hypo-osmotic changes to the RVD as follows: mechanistic stimulation of TRPM7 by cell swelling evokes triggering Ca²⁺ signals, and the resultant CICR response generates Ca²⁺-activated Cl⁻ currents, which are essentially required for RVD (Fig. 5). These results provide fundamental evidence for understanding the molecular mechanism underlying volume regulation in salivary glands.

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Figure 5.

Schematic representation of the physiological changes in submandibular gland acinar cells upon hypotonic stress. TRPM7, transient receptor potential melastatin 7; RyR, ryanodine receptor; ER, endoplasmic reticulum; CACC, Ca²⁺-activated Cl⁻ channel.

Author Contributions

J.M. Kim, S. Choi, contributed to data acquisition, analysis, and interpretation, critically revised the manuscript; K. Park, contributed to conception and design, drafted the manuscript. All authors gave final approval and agree to be accountable for all aspects of the work.