Hypotonic-induced Stretching of Plasma Membrane Activates Transient Receptor Potential Vanilloid Channels and Sodium–Calcium Exchangers in Mouse Odontoblasts

Basic Research—Biology

Hypotonic-induced Stretching of Plasma Membrane Activates Transient Receptor Potential Vanilloid Channels and Sodium–Calcium Exchangers in Mouse Odontoblasts Masaki Sato, PhD,* Ubaidus Sobhan, BDS, PhD,* Maki Tsumura, PhD,*† Hidetaka Kuroda, DDS, PhD,*z Manabu Soya, DDS,*z Aya Masamura, DDS,*z Akihiro Nishiyama, DDS,§ Akira Katakura, DDS, PhD,§ Tatsuya Ichinohe, DDS, PhD,z Masakazu Tazaki, DDS, PhD,† and Yoshiyuki Shibukawa, DDS, PhD*† Abstract Introduction: A number of transient receptor potential (TRP) channels have been identified as membranebound sensory proteins in odontoblasts. However, the activation properties of these channels remain to be clarified. The purpose of this study was to investigate hypotonic stimulation–induced Ca2+ entry via TRP vanilloid subfamily member (TRPV) 1, TRPV2, and TRPV4 channels, which are sensitive to osmotic and mechanical stimuli, and their functional coupling with Na+-Ca2+ exchangers (NCXs) in mouse odontoblast lineage cells. Methods: We examined TRP channel activity by measuring intracellular-free Ca2+ concentration by using fura-2 fluorescence and ionic current recordings with whole-cell patch-clamp methods. Protein localization and messenger RNA expression were characterized using immunofluorescence and reverse-transcription polymerase chain reaction analyses. Results: Extracellular hypotonic solution–induced stretching of plasma membrane resulted in the activation of Ca2+ influx and inward currents. TRPV1, TRPV2, and TRPV4 channel antagonists inhibited the hypotonic stimulation– induced Ca2+ entry and currents. Their respective agonists activated Ca2+ entry. Although the increase in the intracellular free Ca2+ concentration decayed rapidly after the applications of these TRPV channel agonists, NCX inhibitors significantly prolonged the decay time constant. The messenger RNA expression of TRPV1, TRPV2, and TRPV4 channels; NCX isoforms 2 and 3; and dentin sialophosphoprotein were upregulated after 24 hours of exposure to the hypotonic culture medium. Conclusions: These results indicate that stretching of the odontoblast membrane activates TRPV1-, TRPV2-, and TRPV4-mediated Ca2+ entry, and increased intracellular-free Ca2+ concentration is

extruded via NCXs. These results suggest that odontoblasts can act as sensors that detect stimuli applied to exposed dentin and drive a number of cellular functions including dentinogenesis and/or sensory transduction. (J Endod 2013;39:779–787)

Key Words Hydrodynamic theory, hypotonic stimulation, intracellular-free Ca2+ concentration, Na+-Ca2+ exchanger, odontoblast, transient receptor potentials

O

dontoblasts primarily participate in the physiological and developmental formation of dentin, extending elongated cellular processes into dentinal tubules where they are immersed in dentinal fluid. Exposed dentin is sensitive to a wide range of stimuli, including temperature related, chemical, mechanical, and osmotic (1). These stimuli cause displacement of dentinal fluid within the tubules, thereby driving mechanical disturbance of cellular architectural components within the dentinal tubules, such as free nerve endings (2, 3). The role of odontoblasts in this sequence of events remains to be clarified. However, tubules act as hydraulic links between the exposed dentin surface and nerve endings/odontoblast processes located in the tubule inner pulpal ends (4). Odontoblasts also produce ‘‘reactionary dentin’’ as a protective response to harmful external stimuli such as moderate lesions that result from dental caries or cavity preparation (5). This indicates that odontoblasts detect temperaturerelated, osmotic, or mechanical stimulation based on the movement of fluid within dentinal tubules. Additionally, it has been suggested that sensing of such cellular deformation through the movement of dentinal fluid results in the transmission of sensory signals to the nerves (6–8). However, the precise mechanism of how such dentinal pain is generated remains to be clarified because of the structural complexity of the dentin-pulp interface, including odontoblasts, nerve endings, and the liquid content in the dentinal tubules. Transient receptor potential (TRP) channels are a large and functionally versatile family of cation-permeable transmembrane proteins that act as polymodal cellular sensors and are involved in a diverse variety of cellular processes (9–12). It has been shown that TRP channels in acutely isolated mouse odontoblasts respond to thermal and hypotonic (membrane stretching) stimulation (13). Cultured human dental pulp cells that have been differentiated into the odontoblastic phenotype are capable of detecting thermal stimulation via these channels (14). These cells showed sensitivity

From the *Oral Health Science Center, †Department of Physiology, ‡Department of Dental Anesthesiology, and §Department of Oral Medicine, Oral and Maxillofacial Surgery, Tokyo Dental College, Chiba, Japan. Supported by Oral Health Science Center grant (hrc8) and grant-in-aid (grant no. 23592751/90582346) for scientific research from the MEXT of Japan. Address requests for reprints to Dr Yoshiyuki Shibukawa, Oral Health Science Center and Department of Physiology, Tokyo Dental College, Chiba 261-8502, Japan. E-mail address: [email protected] 0099-2399/$ - see front matter Copyright ª 2013 American Association of Endodontists. http://dx.doi.org/10.1016/j.joen.2013.01.012

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Basic Research—Biology when extracellular stimulants such as capsaicin were used as ligands (13, 14). Additionally, TRP vanilloid subfamily member (TRPV) 1 channels in acutely isolated rat odontoblasts were shown to be sensitive to not only extracellular stimulants (6, 7, 15) but also endogenous agonists such as endocannabinoids and endovanilloids (7). Both endocannabinoids and endovanilloids activate TRPV1 channels via intracellular cyclic adenosine monophosphate signals through the activation of the G-protein–coupled cannabinoid receptor in odontoblasts (7). Thus, the expression and thermal/pharmacologic sensitivity of TRP channel subfamilies in odontoblasts are well documented. However, details regarding their mechanosensing properties and functional significance remain to be clarified in odontoblasts. Within the TRP channel subfamily, TRPV1, TRPV2, and TRPV4 have been shown to function in the transduction of osmotic and mechanical stimuli (9, 10, 16). In the present study, we investigated cell membrane stretch-induced activation of TRPV1, TRPV2, or TRPV4 channels in cultured mouse odontoblast lineage cells (OLCs) and examined the functional coupling between the Ca2+ extrusion pathway via Na+-Ca2+ exchangers (NCXs) and these channels.

Materials and Methods Cell Culture Mouse OLCs (17, 18) were cultured in an alpha-minimum essential medium containing 10% fetal bovine serum, 1% penicillinstreptomycin, and 1% Fungizone (Invitrogen, Carlsbad, CA) at 37 C and 5% CO2. The OLCs were positive for various odontoblastrepresentative transcripts from the small integrin binding ligand Nlinked glycoprotein family such as dentin sialophosphoprotein, dentin matrix protein-1, and nestin and were generously provided by Dr Masayuki Tokuda, Kagoshima University, Kagoshima, Japan. Solutions Standard extracellular solution (ECS) was composed of (in millimeters) 135 NaCl, 5 KCl, 2.5 CaCl2, 1 MgCl2, 10 NaH2PO4, 10 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), and 10 glucose, and the pH was adjusted to 7.4 by tris(hydroxymethyl) aminomethane (Tris) (310 mOsm/L). For the extracellular Ca2+free test and standard solution, extracellular Ca2+ was removed (0 mmol/L) from the solutions, and 1 mmol/L EDTA-2Na was added. To prepare test solutions of varying osmolarity values, the extracellular NaCl concentration in standard ECS was reduced to 35 mmol/ L Na+ to prepare a 110-mOsm/L solution (low-NaCl solution). Isotonic, hypotonic, and hypertonic solutions were prepared by adding various concentrations of mannitol to the low-NaCl solution. To obtain the isotonic solution (310 mOsm/L), 200 mmol/L mannitol was added to the low-NaCl solution. To obtain various hypotonic test solutions, between 40 mmol/L (to make 150 mOsm/L) and 190 mmol/L (to make 300 mOsm/L) mannitol was added to the low-NaCl solution. Reagents Fura-2 acetoxymethyl ester was obtained from Dojindo Laboratories (Kumamoto, Japan). Pharmacologic agents, TRPV channels agonist/antagonists (except probenecid and 5-benzyloxytryptamine hydrochloride [5-BOT]), TRP ankyrin subfamily member 1 (TRPA1) channel antagonist (2-[1,3-dimethyl-2,6-dioxo-1,2,3,6-tetrahydro7H-purin-7-yl]-N-[4-isopropylphenyl] acetamide [HC030031]), and NCX inhibitors (except SEA0400) were obtained from Tocris Cookson (Bristol, UK). 5-BOT (a TRP melastatin subfamily member 8 [TRPM8] channel antagonist) and Tris were obtained from Wako Pure Chemicals (Osaka, Japan). SEA0400 was synthesized by Taisho Pharmaceutical Co 780

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(Saitama, Japan) and generously provided by Professors Akemichi Baba and Toshio Matsuda, Osaka University, Osaka, Japan. Except where indicated, all reagents (including probenecid) were obtained from Sigma-Aldrich (St Louis, MO). Stock solutions were prepared in dimethyl sulfoxide and later diluted to the appropriate concentrations in either extracellular solutions or culture medium.

Measurement of Peripheral Cell Length Detached and resuspended OLCs in standard ECS were placed into culture dishes at 37 C and 5% CO2 for 1 hour. Images of OLCs in isotonic or various hypotonic test solutions were acquired by using an intensified charge-coupled device camera mounted on a microscope (Olympus, Tokyo, Japan). The OLC radius was determined from the images (Aquacosmos; Hamamatsu Photonic, Shizuoka, Japan). The calculated peripheral cell lengths in the hypotonic test solutions were normalized to that found in the isotonic solution. The peripheral length of cells in the isotonic test solution was found to be 18.5  2.24 mm (N = 30). Measurement of Ca2+-sensitive Dye Fluorescence OLCs were incubated for 60 minutes (37 C) in standard ECS containing 10 mmol/L fura-2 followed by rinsing with fresh standard ECS. The intracellular free Ca2+ concentration ([Ca2+]i) was measured using the Aquacosmos system and Hamamatsu Photonic software with an excitation wavelength selector and an intensified chargecoupled device camera incorporated onto a microscope (Olympus). The [Ca2+]i was expressed as the fluorescence ratio (RF340/F380) at excitation wavelengths of 380 and 340 nm. Whole-cell Recording Techniques Whole-cell recordings were performed using a conventional patch-clamp recording configuration under voltage-clamp conditions. Patch pipettes (2–5 MU) were pulled from capillary tubes by using a DMZ Universal Puller (Zeitz Instruments, Martinsried, Germany), and the pipettes were filled with an intracellular solution. The intracellular solution contained 140 mmol/L KCl, 10 mmol/L NaCl, and 10 mmol/L HEPES (pH = 7.2 using Tris). Whole-cell currents were measured using a patch-clamp amplifier (L/M-EPC7+; Heka Elektronik, Lambrecht, Germany). The current traces were monitored and stored using pCLAMP (Axon Instruments, Foster City, CA) after digitizing the analog signals at 10 kHz (DigiData 1440A, Axon Instruments) and filtering the signals digitally at 30 Hz using pCLAMP. The data were analyzed offline by using pCLAMP and the technical graphics/analysis program ORIGIN (MicroCal Software, Northampton, MA). Real-time Reverse-transcription Polymerase Chain Reaction Total RNA was extracted from the OLCs by using a modified acid guanidinium–phenol chloroform method. Reverse-transcription, complementary DNA amplification, and polymerase chain reaction (PCR) were performed using the One-Step SYBR Primescript RT-PCR Kit with Thermal Cycle Dice for semiquantitative real-time reverse-transcription (RT)-PCR (TaKaRa-Bio, Shiga, Japan). The forward and reverse primer sets used and the real-time RT-PCR conditions are described in Supplemental Table S1 (available online at www. jendodon.com). The real-time RT-PCR results were quantified using the comparative threshold (2[DDCt]) method (Ct is the cycle threshold) (19, 20). The relative messenger RNA expression level of the target gene was normalized against that of glyceraldehyde-3phosphate dehydrogenase (GAPDH). The Ct for each primer set was JOE — Volume 39, Number 6, June 2013

Basic Research—Biology measured from the samples and averaged. The change in the Ct was then calculated as the difference between the average Ct for the target gene and for GAPDH as the control for the total starting RNA quantity. The delta-delta Ct calculation was then used to assess the fold change in gene expression relative to that of the GAPDH gene.

Immunofluorescence Microscopy Mandibles were dissected from anesthetized 6-week-old Wistar rats perfused with 4% paraformaldehyde in 0.1 mol/L cacodylate buffer (pH = 7.4) through the cardiac left ventricle. Dissected mandibles free of soft tissue were immersed in the same fixative for an additional 12 hours at 4 C. After decalcification with a 5% EDTA-2Na solution for 2–3 weeks at 4 C, selected specimens were dehydrated through a graded series of ethanol, embedded in paraffin, and sagitally sectioned (together with mandibular incisors) to 5 mm in thickness. We performed double immunofluorescence staining of the TRPV2 and TRPV4 channels. To inhibit endogenous peroxidase, dewaxed paraffin sections were treated with 0.5% H2O2 in phosphate-buffered saline (PBS) for 20 minutes. Blocking of nonspecific staining was then performed using 1% bovineserum albumin in PBS for 30 minutes at room temperature. The sections were pretreated by adding 1 mg/mL trypsin for 30 minutes for antigen retrieval and then incubated with a rabbit anti-TRPV2 channel antibody (dilution, 1:100, #KM019; TransGenic, Kumamoto, Japan) or an anti-TRPV4 channel antibody (dilution, 1:100; #ab39260; Abcam, Cambridge, UK) at room temperature for 2 hours. The sections were then incubated with secondary antibodies (Alexa Fluor 488 goat antirabbit; Invitrogen) overnight at 4 C followed by rinsing with PBS. The samples were examined using a conventional fluorescence microscope (Zeiss, Jena, Germany). Statistical Analysis Data are expressed as the mean  standard error or deviation of the mean of N observations, where N represents the number of separate experiments. Statistical differences were evaluated using 1-way analysis of variance or the Student’s t test. A P value <.05 was considered significant.

Results Hypotonic Stimulation–induced Ca2+ Entry in Odontoblasts To measure changes in cell size after exposure to 4 different hypotonic (300, 250, 200, and 150 mOsm/L) or isotonic test solutions (310 mOsm/L), the cell peripheral length was calculated. The normalized peripheral length of cells significantly increased as extracellular osmolarity decreased from 250 to 150 mOsm/L (Fig. 1A, P < .05), indicating that hypotonic stimulation induced stretching of the odontoblast plasma membrane. The [Ca2+]i response in OLCs was expressed as F/F0; the change in the RF340/F380 value (F) was normalized to its resting value (F0) to determine the fluorescence ratio. Each hypotonic test solution (ie, 300, 250, 200, and 150 mOsm/L) elicited a transient increase in [Ca2+]i (Fig. 1B). The semilogarithmic plot in Figure 1C shows F/F0 values as a function of applied osmolarity in the test solutions. The osmolarity dependence of increased [Ca2+]i was obtained by fitting the data to the following Boltzmann function: A ¼ ðA max  A min Þ= 1 þ eðxK=dxÞ þ A min




where K is the half-maximal osmolarity of activation for the applied test solution (301.6 mOsm/L) and Amax is the maximal and Amin the minimal JOE — Volume 39, Number 6, June 2013

F/F0 response. The x indicates applied osmolarity. When extracellular Ca2+ was removed from all test solutions by using 1 mmol/L EDTA, hypotonic stimulation–induced [Ca2+]i responses were completely abolished, indicating that plasma membrane stretching induced by hypotonic stimulation activates Ca2+ influx (Fig. 1D). In the presence of extracellular Ca2+, the addition of hypertonic solution (320 and 370 mOsm/L) elicited a transient increase in [Ca2+]i (Fig. 1E). However, the [Ca2+]i response was small compared with that induced by hypotonic stimulation (Fig. 1F). TRPV channels reportedly exhibit Ca2+-dependent desensitization (21). The hypotonic test solution (200 mOsm/L) was applied repeatedly for 1 minute at 2-minute intervals in the presence of extracellular Ca2+, and the [Ca2+]i response was measured. Repeated application of the hypotonic test solution did not result in a significant desensitizing effect on Ca2+ entry (Fig. 1G). In the presence of external Ca2+, the increased [Ca2+]i after the third application was 98.5%  0.6% (N = 3) compared with that after the first application.

TRPV Channel Antagonists Inhibit Hypotonic Stimulation–induced Ca2+ Influx in Odontoblasts To further examine the Ca2+ influx pathway activated by hypotonic stimulation, the effects of selective antagonists, including TRPV1 (capsazepine [CZP] [Fig. 2A] and AMG9810 [Fig. 2B]) (22), TRPV2 (tranilast [Fig. 2C], tetraethylammonium-chloride [TEA] [Fig. 2D]) (23, 24), TRPV4 (RN-1734 [Fig. 2E]) (25), TRPM8 (5-BOT [Fig. 2F]) (26), and TRPA1 (HC030031 [Fig. 2G]) channels (7), on hypotonic stimulation–induced Ca2+ influx were investigated. In the presence of external Ca2+ (2.5 mmol/L), [Ca2+]i in OLCs showed a rapid and transient increase after exposure to the hypotonic test solution (200 mOsm/L), which was significantly and reversibly inhibited by 1 mmol/L CZP to 49.7%  2.33% (N = 7), 10 nmol/L AMG9810 to 57.9%  5.29% (N = 5), 1 mmol/L tranilast to 76.4%  3.44% (N = 6), 1 mmol/L TEA to 57.0%  1.78% (N = 6), and 1 mmol/L RN-1734 to 50.6%  2.70% (N = 6) (Fig. 2A–E) but not by 1 mmol/L 5-BOT (97.0%  5.12%, N = 4, Fig. 2F) or 10 mmol/L HC030031 (99.3%  0.92%, N = 5, Fig. 2G and I). In the presence of a mixture of TRPV1, TRPV2, and TRPV4 channel antagonists (ie, CZP, tranilast, and RN-1734 at 1 mmol/L each), hypotonic stimulation–induced [Ca2+]i responses were significantly, reversibly, and nearly completely abolished (13.0%  2.70% of the control value [N = 6], Fig. 2H and I). The results indicate that plasma membrane stretching activates Ca2+ influx via TRPV1, TRPV2, and TRPV4 channels but not via TRPM8 and TRPA1 channels. NCX Inhibitors Reduced Ca2+ Extrusion Efficiency after TRPV Channel-mediated Ca2+ Entry in Odontoblasts To investigate the Ca2+ extrusion pathway of increased [Ca2+]i by TRPV1, TRPV2, or TRPV4 channel activation, the contribution of the NCXs was investigated. Ca2+ extrusion efficiency was determined by comparing the time constant of decay calculated on the basis of a single exponential function. In the presence of extracellular Ca2+, the application of 1 mmol/L capsaicin (TRPV1 channel agonist) (7) (Fig. 3A–C), 100 nmol/L probenecid (TRPV2 channel agonist) (23) (Fig. 3D–F), or 1 mmol/L RN-1747 (TRPV4 channel agonist) (25) (Fig. 3G–I) elicited a transient increase in [Ca2+]i to peak F/F0 values. This increase in [Ca2+]i decayed rapidly after the application of the TRPV channel agonists (black lines in Fig. 3A–I). When NCX inhibitors KB-R7943 (1 mmol/L) (Fig. 3A, D, and G), SN-6 (3 mmol/L) (Fig. 3B, E, and H), or SEA0400 (1 mmol/L) (Fig. 3C, F, and I) (27) were applied after the activation of each channel, the decay time Mechano-/Osmo-sensitive TRP Channels in Odontoblasts

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Figure 1. Hypotonic stimulation–induced cell stretching and activated Ca2+ entry in odontoblasts. (A) The relationship between the change in the peripheral length of cells and extracellular osmolarity. Normalized cell size increased with decreasing extracellular osmolarity (normalized cell size was 1.0 in isotonic solution, 1.04  0.08-fold in 300 mOsm/L, 1.47  0.21-fold in 250 mOsm/L, 1.62  0.24-fold in 200 mOsm/L, and 1.63  0.26-fold in 150 mOsm/L). Each bar denotes the mean  standard deviation of the 30 cells tested. Statistically significant differences between columns (shown by solid lines) are indicated by asterisks. *P < .05. (B–D) Examples of transient [Ca2+]i increases in a series of hypotonic test solutions (ie, 300, 250, 200, and 150 mOsm/L) (upper open boxes in B and D) compared with [Ca2+]i in isotonic test solution (upper filled boxes in B and D) in the presence (B, 2.5 mmol/L) or absence (D) of extracellular Ca2+ (white boxes at bottom in B and D). (C) A semilogarithmic plot showing F/F0 values as the function of applied osmolarity (see text). (E and F) Examples of transient [Ca2+]i increases in hypertonic test solutions (320 and 370 mOsm/L) (upper open boxes in E) compared with [Ca2+]i in isotonic test solution (upper filled boxes in E) in the presence of extracellular Ca2+ (white box at bottom). (F) A semilogarithmic plot showing F/F0 values as a function of applied osmolarity. (G) An example of a trace of a transient [Ca2+]i increase after successive 1-minute applications of hypotonic test solution (200 mOsm/L) (upper open boxes) at 2-minute intervals in presence of extracellular Ca2+ (2.5 mmol/L) (white box at bottom). Each point indicates the mean  standard error of separate experiments; the number of tested cells is shown in parentheses.

constant showed a significant increase (gray lines in Fig. 3A–I and Fig. 3J–L).

Hypotonic Stimulation–induced Cell Stretching Activates Inward Currents To further analyze the membrane expression of TRPV1, TRPV2, and TRPV4 channels, hypotonic stimulation–induced 782

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inward currents were recorded using the whole-cell patch-clamp method (Fig. 4). Peak values of inward currents at a holding potential of 60 mV were 337.6  53.5 pA (N = 9) in the hypotonic test solution (200 mOsm/L). Hypotonic test solution–elicited inward currents were significantly inhibited by 1 mmol/L CZP to 167.0  44.5 pA (N = 3), 1 mmol/L tranilast to 91.4  17.5 pA (N = 3), and 1 mmol/L RN-1734 to 226.2  32.3 pA (N = 3) (Fig. 4A and B). JOE — Volume 39, Number 6, June 2013

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Figure 2. The effect of TRPV1, TRPV2, and TRPV4 channel antagonists on hypotonic stimulation–induced Ca2+ entry. (A–H) Examples of transient [Ca2+]i increases after the replacement of isotonic test solution with 200 mOsm/L hypotonic test solution (upper open boxes) in the presence of extracellular Ca2+ (2.5 mmol/L) with or without (A) 1 mmol/L CZP, (B) 10 nmol/L AMG9810, (C) 1 mmol/L tranilast, (D) 1 mmol/L TEA, (E) 1 mmol/L RN-1734, (F) 1 mmol/L 5-BOT, or (G) 10 mmol/L HC030031 channel antagonists or (H) a cocktail of antagonists (CZP, tranilast, and RN-1734, 1 mmol/L each). The upper black bars in A–H indicate times of adding antagonists. (I) A bar graph summarizing an increase in [Ca2+]i by hypotonic (200 mOsm/L) stimulation–induced Ca2+ influx without (open columns) or with 1 mmol/L CZP, 10 nmol/L AMG9810, 1 mmol/L tranilast, 1 mmol/L TEA, 1 mmol/L RN-1734, 1 mmol/L 5-BOT, 10 mmol/L HC030031, or a cocktail of antagonists (CZP, tranilast, and RN-1734, 1 mmol/L each) (black columns). Each point indicates the mean  standard error of the number of separate experiments; the number of tested cells is shown in parentheses. Statistically significant differences between columns (shown by solid lines) are indicated by asterisks. *P < .05.

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Figure 3. NCX inhibitors prolong Ca2+ extrusion after TRPV channel–mediated Ca2+ entry. (A–I) Examples of transient [Ca2+]i increases with the addition of (A–C) 1 mmol/L capsaicin, (D–F) 0.1 mmol/L probenecid, or (G–I) 1 mmol/L RN-1747. After the application of these TRPV channel agonists, a decay in [Ca2+]i increase to the basal level (60-second period) was examined by fitting to a single exponential function without (black lines in A–I) or with (gray lines in A–I) NCX inhibitors: (A, D, and G) 1 mmol/L KB-R7943, (B, E, and H) 3 mmol/L SN-6, or (C, F, and I) 1 mmol/L SEA0400. Upper black bars indicate times of addition of TRPV channel agonists, and gray bars show times of addition of NCX inhibitors. (J–L) A bar graph summarizing decay time constant after the application of (J) 1 mmol/L capsaicin, (K) 0.1 mmol/L probenecid, or (L) 1 mmol/L RN-1747 without (upper columns in J–L) or with NCX inhibitors: (second upper columns in J–L) 1 mmol/L KB-R7943, (second lower columns in J–L) 3 mmol/L SN-6, or (lower columns in J–L) 1 mmol/L SEA0400. Each bar indicates the mean  standard error of the number of separate experiments; the number of tested cells is shown in parentheses. Statistically significant differences between columns (shown by solid lines) are indicated by asterisks. *P < .05.

Up-regulation of Ca2+ Signal Proteins in Response to Hypotonic Stimulation in Odontoblasts After 24 hours of exposure to the hypotonic medium, messenger RNA expression of TRPV1, TRPV2, and TRPV4 channels and NCX isoforms were measured using real-time RT-PCR and compared with the expression in cells subjected to isotonic medium (Fig. 5). Hypotonic stimulation significantly up-regulated messenger RNA expression of TRPV1 (1.22  0.05-fold), TRPV2 (2.12  0.17-fold), TRPV4 (1.68 784

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 0.15-fold), NCX2 (1.45  0.15-fold), NCX3 (1.66  0.14-fold), and dentin sialophosphoprotein (DSPP; 1.26  0.02-fold) (N = 5) (black columns in Fig. 5). In contrast, no significant change was observed in the messenger RNA expression of adenosine triphosphate (ATP) synthase (H+-transporting ATP synthase), dentin matrix protein-1, or nestin, whereas the expression of NCX1 was downregulated. No up- or down-regulation of this messenger RNA expression was observed after 24 hours of exposure to the hypotonic medium in the JOE — Volume 39, Number 6, June 2013

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Figure 4. Hypotonic stimulation–induced cell stretching activates inward currents. (A) An example of traces of inward currents recorded by using whole-cell patch-clamp recordings after the extracellular application of hypotonic test solution (200 mOsm/L, white boxes) without or with TRPV1 (1 mmol/L CZP), TRPV2 (1 mmol/L tranilast), and TRPV4 (1 mmol/L RN-1734) channel antagonists (black boxes). The holding potential was 60 mV. Extracellular solutions used were isotonic or hypotonic test solutions. (B) A bar graph summarizing peak values of inward currents after hypotonic (200 mOsm/L) stimulation without (open columns) or with 1 mmol/L CZP, 1 mmol/L tranilast, and 1 mmol/L RN-1734 (black columns). Each point indicates the mean  standard deviation of the number of individual experiments. Statistically significant differences between columns (shown by solid lines) are indicated by asterisks. *P < .05.

presence of TRPV channel antagonists CZP, tranilast, and RN-1734 (0.1 mmol/L each) (slashed columns in Fig. 5).

Localization and Expression of TRPV2 and TRPV4 Channels in Rat Odontoblasts In previous studies, we described the localization of TRPV1 channels (7, 15) on the processes inside the dentinal tubules as well as that of NCXs (27) on the distal plasma membrane of rat odontoblasts. Therefore, in this study, we examined the localization and expression of TRPV2 and TRPV4 channels in native rat odontoblasts. Immunoreactivity of TRPV2 channels (Fig. 6A) was observed on the distal membrane, whereas that of TRPV4 channels (Fig. 6B) was distributed on odontoblast cell bodies.

Discussion Our results showed the hypotonic stimulation–induced activation of TRPV1, TRPV2, and TRPV4 (ie, membrane stretchingactivated TRPV) channels in odontoblasts. An increase in [Ca2+]i via TRPV channel activation was extruded by NCXs, indicating functional coupling between TRPV channels and NCXs. These results indicate that odontoblasts functionally detect plasma membrane stretching to drive cellular function(s). The present results also showed that TRPV1, TRPV2, and TRPV4 channels had no desensitizing effect on Ca2+ entry after membrane stretch. This implies that in odontoblasts these channels may contribute to sensory transduction in pain because it is widely known that pain sensation does not generally show adaptation. Both TRPM8 and TRPA1 channel antagonists did not affect hypotonic stimulation–induced Ca2+ influx in OLCs, which are mouse OLCs. It has been reported that TRPM8 and TRPA1 channels are expressed in human cells (14) but not in mouse odontoblasts (13). Our results agree with the results from mouse odontoblasts; however, further studies are necessary to clarify their expression and functional roles. TRPM8 and TRPA1 channels are expressed in acutely isolated rat odontoblasts, and TRPA1 channels show sensitivity to membrane stretch (personal communication, Y Shibukawa, 2013). To measure [Ca2+]i and inward currents and perform PCR analysis, cellular stimulation was applied using a series of extracellular hypotonic solutions (150–300 mOsm/L). A recent study on the JOE — Volume 39, Number 6, June 2013

Figure 5. The change in expression of TRPV channels, NCXs, ATP synthase, and markers specific for odontoblasts in response to hypotonic stimulation. The expression of each messenger RNA was determined using semiquantitative real-time RT-PCR analysis relative to GAPDH messenger RNA. Relative upregulation of TRPV1, TRPV2, TRPV4, NCX2, NCX3, and DSPP was observed in OLCs cultured for 24 hours in 170 mOsm/L hypotonic alpha-minimum essential culture medium (by adding sterilized distilled water) (filled columns) compared with values in isotonic alpha-minimum essential medium (open columns) and in hypotonic alpha-minimum essential culture medium in the presence of CZP, tranilast, and RN-1734 (slashed column) (0.1 mmol/L each). Each bar indicates the mean  standard error of 5 experiments. Note that the relative messenger RNA expression level does not indicate the absolute level of expression. Statistically significant differences in relative messenger RNA expression between cells exposed to hypotonic (filled or slashed columns) or isotonic culture medium (open columns) are indicated by asterisks. *P < .05. Note that we did not observe any significant changes between the percentage of living cells after 24 hours of exposure to hypotonic medium without (94.4%  2.0%) or with (95.9%  1.0%) TRPV channel antagonists or isotonic medium (94.2%  1.0%) (N = 5 each). Cell viability was separately examined using 0.4% trypan blue (Invitrogen) and hemocytometer (Digital Bio, Seoul, Korea).

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Basic Research—Biology (Fig. 1A), this was sufficient to activate TRPV1, TRPV2, or TRPV4 channels. The plasma membrane NCX is a bidirectional transporter that catalyzes electrogenic exchange of Na+ for Ca2+ depending on the electrical and chemical gradients of substrate ions (29, 30), which plays a critical role in regulating [Ca2+]i to primarily pump Ca2+ outside of the cell. Mammalian NCX comprises 3 NCX isoforms: NCX1, NCX2, and NCX3. In the present study, NCX antagonists KB-R7943, SN-6, and SEA0400 increased the decay time constant after TRPV1-, TRPV2-, and TRPV4-mediated Ca2+ entry, which indicates that Ca2+ extrusion efficiency was lowered after the inhibition of NCXs. These results agree with those of our previous study showing the expression of NCX1 and NCX3 (but not NCX2) at the distal membrane in rat odontoblasts, with potencies of NCX3 > NCX1 (27). However, in the present study, up-regulation of NCX2 messenger RNA and down-regulation of NCX1 were observed after 24 hours of hypotonic stimulation. Further studies are needed to confirm the modular effect of expressing these NCXs during plasma membrane stretching. However, the results of the present study clearly showed that NCXs are functionally coupled with membrane stretch–sensitive TRPV channel activation in odontoblasts to regulate [Ca2+]i according to Ca2+ extrusion results. Transmembrane Ca2+ signals may play an important role in forming dentin as a response to cellular disturbances caused by dentinal fluid movement after external stimuli applied to the dentin surface (7). Additionally, 24 hours of exposure of odontoblasts to hypotonic stimulation resulted in the up-regulation of not only TRPV channels NCX2 and NCX3 but also DSPP. This hypotonic stimulation–induced up-regulation of messenger RNA was not observed in the presence of a mixture of TRPV1, TRPV2, and TRPV4 channel antagonists, which indicates that plasma membrane stretching specifically accelerates the expression of TRPV channels, NCXs, and DSPP.

Conclusions Figure 6. Localization of TRPV2 and TRPV4 channels in rat dental pulp. Tall, columnar, and polarized odontoblasts were observed at dental pulp section. These cells were positive for (A) TRPV2 and (B) TRPV4 channel immunoreactions (green). Nuclei are shown in blue. D, dentin; O, odontoblast layer; P, dental pulp. Scale bars = 50 mm (A) and 25 mm (B). No fluorescence was detected in negative control cells (not shown).

relationship between dentinal pain intensity and dentinal fluid flow rates induced by stimulation of the exposed dentin surface showed that outward flow induced greater pain than inward flow (28). Outward flow of dentinal fluid in the tubules (ie, away from pulp ends into the dentin surface) elicits stretching of the plasma membrane inside the dentinal tubules (ie, odontoblast processes and/or free nerve endings) (4). In the present study, the hypotonic solution elicited cellular membrane stretching in odontoblasts (Fig. 1). The peripheral length of the cells increased as extracellular osmolarity decreased, and hypotonic stimulation–induced Ca2+ influx showed a dependence on osmolarity. However, only a small increase was observed in [Ca2+]i in response to hypertonic compared with the increase observed in the response to hypotonic solution, thereby inducing shrinkage of the plasma membrane. Further studies are needed to clarify whether these results reflect the odontoblast response to the outward and inward dentinal flow to elicit stretching and shrinking of the odontoblast membrane, respectively (18). The value of half-maximal activation osmolarity (301.6 mOsm/L) for the hypotonic stimuli–induced [Ca2+]i response showed a negative change to 8.4 mOsm/L from isotonic 310 mOsm/L (Fig. 1). Although only a small change in peripheral length of cells was observed 786

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The present results indicate that cell membrane stretching activates Ca2+ influx via TRPV1, TRPV2, or TRPV4 channels, with subsequent increases in [Ca2+]i extruded by NCXs. Known as the hydrodynamic mechanism, all stimuli to the exposed dentin surface elicit fluid movement in the dentinal tubules (1–3, 6). This, in turn, disturbs odontoblastic processes within the dentinal tubules and activates their TRPV channels. This indicates that odontoblasts can act as sensors (31), detecting stimuli applied to exposed dentin and driving cellular functions such as reactionary dentin formation after stimulation and sensory transduction.

Acknowledgments We would like to thank Associate Professors Jeremy Williams (for assistance with the English of the manuscript) and Kenji Ikegami (for advice on the physics involved). We would also like to thank Professors Akemichi Baba and Toshio Matsuda of Osaka University for gifting SEA0400 as well as Masayuki Tokuda of Kagoshima University for providing OLCs. The authors deny any conflicts of interest related to this study.

Supplementary Material Supplementary material associated with this article can be found in the online version at www.jendodon.com (http://dx.doi. org/10.1016/j.joen.2013.01.012).

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Basic Research—Biology 2. Br€annstr€om M. A hydrodynamic mechanism in the transmission of pain-producing stimuli through the dentine. In: Anderson DJ, ed. Sensory Mechanisms in Dentine. Oxford, England: Pergamon Press; 1963:73–9. 3. Andrew D, Matthews B. Displacement of the contents of dentinal tubules and sensory transduction in intradental nerves of the cat. J Physiol 2000;529: 791–802. 4. Lin M, Luo ZY, Bai BF, et al. Fluid mechanics in dentinal microtubules provides mechanistic insights into the difference between hot and cold dental pain. PLoS One 2011;6:e18068. 5. Goldberg M, Smith AJ. Cells and extracellular matrices of dentin and pulp: a biological basis for repair and tissue engineering. Crit Rev Oral Biol Med 2004;15: 13–27. 6. Magloire H, Maurin JC, Couble ML, et al. Topical review. Dental pain and odontoblasts: facts and hypotheses. J Orofac Pain 2010;24:335–49. 7. Tsumura M, Sobhan U, Muramatsu T, et al. TRPV1-mediated calcium signal couples with cannabinoid receptors and sodium-calcium exchangers in rat odontoblasts. Cell Calcium 2012;52:124–36. 8. Liu X, Yu L, Wang Q, et al. Expression of Ecto-ATPase NTPDase2 in human dental pulp. J Dent Res 2012;91:261–7. 9. Minke B, Cook B. TRP channel proteins and signal transduction. Physiol Rev 2002; 82:429–72. 10. Clapham DE. TRP channels as cellular sensors. Nature 2003;426:517–24. 11. Minke B. The history of the Drosophila TRP channel: the birth of a new channel superfamily. J Neurogenet 2010;24:216–33. 12. Wu LJ, Sweet TB, Clapham DE. International Union of Basic and Clinical Pharmacology. LXXVI. Current progress in the mammalian TRP ion channel family. Pharmacol Rev 2010;62:381–404. 13. Son AR, Yang YM, Hong JH, et al. Odontoblast TRP channels and thermo/mechanical transmission. J Dent Res 2009;88:1014–9. 14. El Karim IA, Linden GJ, Curtis TM, et al. Human odontoblasts express functional thermo-sensitive TRP channels: implications for dentin sensitivity. Pain 2011;152: 2211–23. 15. Okumura R, Shima K, Muramatsu T, et al. The odontoblast as a sensory receptor cell? The expression of TRPV1 (VR-1) channels. Arch Histol Cytol 2005;68:251–7. 16. Liedtke WB, Heller S. TRP Ion Channel Function in Sensory Transduction and Cellular Signaling Cascades. Boca Raton, FL: CRC Press; 2007.

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17. Arany S, Kawagoe M, Sugiyama T. Application of spontaneously immortalized odontoblast cells in tooth regeneration. Biochem Biophys Res Commun 2009; 381:84–9. 18. Fujisawa M, Tokuda M, Morimoto-Yamashita Y, et al. Hyperosmotic stress induces cell death in an odontoblast-lineage cell line. J Endod 2012;38:931–5. 19. Livak KJ, Schmittgen TD. Analysis of relative gene expression data using real-time quantitative PCR and the 2(-Delta Delta C(T)) method. Methods 2001;25:402–8. 20. Schmittgen TD, Livak KJ. Analyzing real-time PCR data by the comparative C(T) method. Nat Protoc 2008;3:1101–8. 21. Mohapatra DP, Nau C. Regulation of Ca2+-dependent desensitization in the vanilloid receptor TRPV1 by calcineurin and cAMP-dependent protein kinase. J Biol Chem 2005;280:13424–32. 22. Vriens J, Appendino G, Nilius B. Pharmacology of vanilloid transient receptor potential cation channels. Mol Pharmacol 2009;75:1262–79. 23. Mihara H, Boudaka A, Shibasaki K, et al. Involvement of TRPV2 activation in intestinal movement through nitric oxide production in mice. J Neurosci 2010;30: 16536–44. 24. Juvin V, Penna A, Chemin J, et al. Pharmacological characterization and molecular determinants of the activation of transient receptor potential V2 channel orthologs by 2-aminoethoxydiphenyl borate. Mol Pharmacol 2007;72:1258–68. 25. Vincent F, Acevedo A, Nguyen MT, et al. Identification and characterization of novel TRPV4 modulators. Biochem Biophys Res Commun 2009;389:490–4. 26. DeFalco J, Steiger D, Dourado M, et al. 5-benzyloxytryptamine as an antagonist of TRPM8. Bioorg Med Chem Lett 2010;20:7076–9. 27. Tsumura M, Okumura R, Tatsuyama S, et al. Ca2+ extrusion via Na+-Ca2+ exchangers in rat odontoblasts. J Endod 2010;36:668–74. 28. Charoenlarp P, Wanachantararak S, Vongsavan N, et al. Pain and the rate of dentinal fluid flow produced by hydrostatic pressure stimulation of exposed dentine in man. Arch Oral Biol 2007;52:625–31. 29. Matsuda T, Takuma K, Baba A. Na(+)-Ca2+ exchanger: physiology and pharmacology. Jpn J Pharmacol 1997;74:1–20. 30. Blaustein MP, Lederer WJ. Sodium/calcium exchange: its physiological implications. Physiol Rev 1999;79:763–854. 31. Ichikawa H, Kim HJ, Shuprisha A, et al. Voltage-dependent sodium channels and calcium-activated potassium channels in human odontoblasts in vitro. J Endod 2012;38:1355–62.

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