Hypo-osmolarity induces apoptosis resistance via TRPV2-mediated AKT-Bcl-2 pathway

Abstract

In cirrhosis, several molecular alterations such as resistance to apoptosis could accelerate carcinogenesis. Recently, mechanotransduction has been attracting attention as one of the causes of these disturbances. In patients with cirrhosis, the serum sodium levels progressively decrease in the later stage of cirrhosis, and hyponatremia leads to serum hypo-osmolality. Since serum sodium levels in patients with cirrhosis with liver cancer are inversely related to cancer’s number, size, stage, and cumulative survival, we hypothesized that hypo-osmolality-induced mechanotransduction under cirrhotic conditions might contribute to oncogenesis and/or progression of hepatocellular carcinoma (HCC). In this study, we adjusted osmosis of culture medium by changing the sodium chloride concentration and investigated the influence of hypotonic conditions on the apoptosis resistance of an HCC cell line, HepG2, using a serum-deprivation-induced apoptosis model. By culturing the cells in a serum-free medium, the levels of an antiapoptotic protein Bcl-2 were downregulated. In contrast, the hypotonic conditions caused apoptosis resistance by upregulation of Bcl-2. Next, we examined which pathway was involved in the apoptosis resistance. Hypotonic conditions enhanced AKT signaling, and constitutive activation of AKT in HepG2 cells led to upregulation of Bcl-2. Moreover, we revealed that the enhancement of AKT signaling was caused by intracellular calcium influx via a mechanosensor, TRPV2. Our findings suggested that hyponatremia-induced serum hypotonic in patients with cirrhosis promoted the progression of hepatocellular carcinoma.

NEW & NOTEWORTHY Our study first revealed that hypo-osmolarity-induced mechanotransduction enhanced calcium-mediated AKT signaling via TRPV2 activation, resulting in contributing to apoptosis resistance. The finding indicates a possible view that liver cirrhosis-induced hyponatremia promotes hepatocellular carcinogenesis.

INTRODUCTION

Primary liver cancer, hepatocellular carcinoma (HCC), is the sixth most common cancer worldwide and the third most common cause of cancer mortality (1). Major risk factors for HCC include infection with hepatitis B or C virus, alcoholic liver disease, and possibly nonalcoholic steatohepatitis. Most of these pathogeneses lead to the formation and progression of cirrhosis, which is present in 80%–90% of patients with HCC (2). In the cirrhosis stage, several molecular alterations such as loss of cell-cycle checkpoints, activation of oncogenic pathways, and resistance to apoptosis could accelerate carcinogenesis, which is induced by stimulation such as chemicals, cytokines, and growth factors (3). In addition, recently, mechanotransduction, which translates mechanical stimulation including temperature, pH, share stress, and osmotic pressure into biochemical signals, has been attracting attention as one of the causes of these disturbances.

Cirrhosis causes reduction in effective arterial blood volume because of arterial vasodilation secondary to increased production of nitric oxide, endotoxins, and other vasodilators. This leads to activation of the renin-angiotensin-aldosterone axis and impairment of free water excretion, resulting in hyponatremia (4). In the early stage of cirrhosis, the serum sodium levels are within the normal range (>135 mmol/L). However, they progressively decrease in the later stage of cirrhosis. The prevalence of low serum sodium concentration is observed in hospitalized 997 patients with cirrhosis (serum sodium concentration: ≤135 mmol/L, <130 mmol/L, <125 mmol/L, and <120 mmol/L was 49.4%, 21.6%, 5.7%, and 1.2%, respectively) (5, 6). Since sodium is the most important osmosis-regulating substance, hyponatremia leads to serum hypo-osmolality in patients with cirrhosis (5, 7). In hyponatremia-induced hypo-osmotic conditions, extracellular fluid moves into the cells to maintain the osmotic balance, and the resulting cell swelling causes mechano transduction (8). Hypo-osmotic cell swelling causes typical metabolic alterations such as glycolysis, urea synthesis, and ammonia formation from the amino acids (9), whereas its detailed mechanism is still unknown. Mechanotransduction plays crucial roles in cell activation, differentiation, proliferation, apoptosis, and even tumorigenesis (10, 11). Moreover, serum sodium levels in patients with cirrhosis with liver cancer are inversely related to the number, size, stage, and cumulative survival of cancer (12).

Transient receptor potential vanilloid 2 (TRPV2) is a sensor of mechanotransduction and can be activated by hypo-osmolarity-induced cell stretch (13). TRPV2 is widely distributed in human organs and tissue, including brain, vascular smooth muscle cells, gastrointestinal tract, macrophages, urothelial tract, and cardiac intercalated disks (14, 15). TRPV2 was reported to be involved in liver carcinogenesis at the cirrhotic stage, whereas little TRPV2 expression was observed in normal hepatocytes (16). Therefore, we hypothesized that hypo-osmolality-induced mechanotransduction under cirrhotic conditions might contribute to oncogenesis and progression of HCC. Thus, in this study, we examined the influence of hypo-osmolarity on the cell survival of HCC cells and showed that hypo-osmolarity induced the apoptosis resistance via the TRPV2-mediated AKT-Bcl-2 pathway thorough mechano-sensor.

MATERIALS AND METHODS

Preparation of Osmotic Pressure-Adjusted Culture Medium and Apoptosis Induction

Osmotic pressure was modified by adjusting sodium concentration using Dulbecco’s Modified Eagle Medium (Fuji Film Co., Tokyo, Japan), distilled water, and 1.4% sodium chloride solution (Table 1). Osmotic pressure was measured by osmometer (OM802, Vogel, Germany).

Table 1.

Preparation of osmotic pressure-adjusted culture medium

Culture Medium	Mosmol/kg	dDW, mL	DMEM, mL	1.4% NaCl in dDW, mL
Isotonic medium	330	3.5	4	0.5
Middle tonic medium	300	3.0	4	1.0
Hypotonic medium	270	2.5	4	1.5
(reference)
DMEM 10% FBS	336
DMEM FBS free	336

Calcium concentration of all medium is 1.8 mM.

Apoptosis was induced by culturing cells in serum-free (17) and osmolarity-modified culture medium for 24 or 48 h using human HCC cell line HepG2. Since albumin is also reported to prevent apoptosis (18–20), we used bovine serum albumin (BSA) containing medium as control in this experiment.

For calcium influx experiment, PBS (−) buffer (Fuji Film Co.) was used for calcium-free medium and PBS (+) buffer (calcium concentration; 0.90 mM, Nacalai tesque, Kyoto, Japan) for calcium-containing medium.

Cell Viability Assay

Cell viability was examined by the cell counting kit-8 (CCK-8) assay (Dojindo, Kumamoto, Japan) according to the manufacturer’s protocol. A total of ∼5 × 10³ hepatocellular carcinoma cell lines, HepG2, HLF, and Li7 or mouse primary hepatocytes were plated in 96-well plates in triplicate and cultured in 100 μL of osmosis-regulated culture medium containing 10% FBS. After 24 h incubation, culture medium was aspirated, then 50 μL CCK-8 solution was added to each well and incubated for 2 h at 37°C. Absorbance was then recorded at 450 nm using Multifunctional microplate reader SpectraMax M5 (Molecular Devise, San Jose, CA).

As other assays for cell proliferation, the incorporation of radioactively labelled tritiated thymidine was used. After a fixed time period, the cells were washed and lysed, after which the incorporated radioactivity in the DNA of the cells was quantified.

Apoptosis Assay

Apoptotic cells were detected by the terminal deoxyribonucleotidyl transferse (TdT)-mediated biotin-16-dUTP nick-end labelling (TUNEL) stain (In situ Apoptosis Detection Kit, TAKARA BIO, Shiga, Japan) or Annexin stain (Annexin V-FITC kit, BioLegend, San Diego, CA) in accordance with the manufacturer’s instructions. Briefly, apoptosis in HepG2 cells was induced by serum deprivation in parallel with culturing in osmolarity-adjusted medium for 48 h. Then, the cells were fixed with 4% paraformaldehyde. After blocking using H₂O₂ containing methanol for 30 min followed by permeabilization, TUNEL reaction buffer was added and incubated for 1 h. After the reaction with anti-AITC HRP conjugate for 30 min, nuclei were stained with DAPI for 15 min. FITC-positive cells were observed by microscope. For quantification, TUNEL-positive cells were counted in different five fields, respectively. For Annexin V stain, HepG2 cells were stained with Annexin (1 mg/mL) for 30 min. The population of positive stains for Annexin V was analyzed by flow cytometry.

Transfection of siRNA

HepG2 cells were seeded in 12-well plates 1 day before transfecting with 100 nM TRPV2 siRNA using Lipofectamine RNAiMAX (Invitrogen), according to the manufacturer’s protocol. At 48 h after transfection, the cells were used in indicated experiments.

Quantitative PCR of Analysis

RNA was extracted from cells using TRIzol reagent (Thermo Fisher Scientific, Waltham, MA) and Direct-zol RNA Miniprep (Zymo Research, Irvine, CA). Quantitative PCR (qPCR) was performed using cDNA generated from RNA and the SuperScript III Reverse Transcriptase kit (Thermo Fisher Scientific). qPCR reaction was carried out using SYBR green PCR master mix (Thermo Fisher Scientific) in the Thermal Cycler Dice Real Time System 2 (TAKARA BIO). The values were quantified using the comparative CT method and were normalized to 18S ribosomal RNA. The data were expressed as the ratio to the average of control group. The primers used in this study are human 18S forward 5′- CAGCCACCCGAGATTGAGCA-3′, reverse 5′- TAGTAGCGACGGGCGGTGTG-3′, human TRPV2 forward 5′- TGTAGCCCTGGTGAGCCT-3′, reverse 5′- CCAACGGTCAGCATCACA-3′, mouse 18S forward 5′- ATTGGAGCTGGAATTACCGC-3′, and reverse; 5′- CGGCTACCACATCCAAGGAA-3′. Data are expressed as means ± SD of three different samples.

Western Blot Analysis

After starvation with serum-free iso-osmotic culture medium, the culture medium was changed to iso-osmotic or hypo-osmotic solution. Then the cells were harvested at the indicated points. Cells were homogenized with radioimmunoprecipitation assay (RIPA) buffer (50 mM Tris·HCl at pH 7.5, 150 mM NaCl, 1% Triton X-100, 1% SDS) containing protease inhibitor cocktail cOmplete Mini (Roche, Basel, Switzerland) and phosphatase inhibitors (1 mM sodium fluoride, 1 mM β-glycerol phosphate, and 1 mM sodium vanadate). Protein samples were subjected to 8%–15% SDS-polyacrylamide gel electrophoresis and were transferred to polyvinylidene difluoride membranes using standard Western blot techniques. After being blocked with 5% skim milk, the membranes were probed with primary antibodies diluted at 1:1,000 to 5,000 and horseradish peroxidase-conjugated secondary antibodies diluted at 1:5,000. Immunoreactive bands were visualized using the ImmunoStar Zeta or ImmunoStar LD system and were detected using the LAS3000 or LAS4000 device (GE Healthcare, Chicago, IL). WB Stripping Solution (Nacalai tesque, Kyoto, Japan) was used to remove the antibodies from the Western blot membrane for reprobing. The quantification of Western Blot bands was performed by using ImageJ software version 1.52 [National Institutes of Health (NIH), Bethesda, MD] and normalized to GAPDH or control. Data are expressed as means ± SD of three different experiments.

Primary Antibodies

The following antibodies were used for Western blot analysis and immunofluorescence: anti-Bcl-2 (1:2,000, No. 2870, 50E3), anti-caspase 3 (1:2,000, No. 9665, G10), anti-AKT (1:3,000, No. 9272), anti-phosphorylated AKT (1:3,000, No. 4060, D9E), anti-Bax (1:2,000, No. 5023, D2E11), and anti-p53 (1:2,000, No. 2524, 1C12), were from Cell Signaling Technology (Danvers, MA). Anti-TRPV2 (1:1,000, No. NBP1-92576) was from Novus Biologicals (Littleton, CO), and anti-GAPDH (1:5,000, No. AB2302) was from Millipore (Billerica, MA).

AKT Constitutive Activation Plasmid

Myristoylated AKT known as AKT constitutive active form was purchased from Addgene (No. 10841, Cambridge, MA). The myristoylated AKT and control vectors were transfected into HepG2 cells using Lipofectamine 3000 (Invitrogen, Carlsbad, CA). The transfected cells were cultured with or without FBS and sampled with RIPA buffer for Western blot analysis.

Measurement of DNA Synthesis

Wells-attached HepG2 cells were incubated in 0.5 mL/well of DMEM-10% dialyzed FBS containing 37 KBq/mL (1 μCi/mL) of methyl-[³H]thymidine for 90 min at 37°C in a moisturized incubator gassed with 95% air and 5% CO₂. On the termination of incubation, 0.5 mL of 10% trichloroacetic acid (TCA) was added, left for 2 h at 4^°C, washed three times with 0.5 mL of cold PBS (−), and dissolved with 0.5 mL of 1 N NaOH at 37°C overnight. Aliquots (0.4 mL) of the NaOH lysates were mixed with 3.6 mL of a scintillation cocktail, Hionic-Fluor (PerkinElmer Japan) in counting vials, which were subjected to determining the radioisotope intensity using a radioisotope liquid scintillator (Aloka, Japan).

Chemicals

Tranilast, a TRPV2 antagonist, was purchased by Fuji Film Company. Probenecid, a TRVP2 agonist (0.5 mM) (21), was from Sigma Aldrich (St. Louis, MO) and triciribine, an AKT inhibitor (5 µM) (22) were purchased from WAKO. Methyl-[³H]thymidine (740 GBq/mmol) was a product (NET027E) of PerkinElmer, Inc. (Waltham, MA).

Chronic Hepatitis Induction

Chronic hepatitis was induced by 12 times intraperitoneal injections of carbon tetrachloride (CCl4, twice a week) to 8- to 10-wk-old C57BL/c mice (0.5 µL/g, dissolved in corn oil at a ratio of 1:3). This experiment was approved by the institutional animal care and use committee of Osaka Metropolitan University and was conducted in compliance with the guidelines.

Statistical Analysis

All data are expressed as the means ± standard deviation. All data were analyzed by using ANOVA with a post hoc Dunnet’s test or unpaired Student’s t test.

RESULTS

Hypo-Osmotic Stimulation Suppressed Serum-Deprivation-Induced Apoptosis

The cell number was reduced by serum depletion. In contrast, hypo-osmotic stimulation suppressed the decrease in cell number (Fig. 1A). We then quantitatively evaluated the effect of hypo-osmotic stimulation on cell viability and DNA synthetic activity as shown Fig. 1, B and C, respectively. The hypo-osmotic stimulation showed a significant increase in these activities. The effect was also observed in other hepatocellular carcinoma cell lines HLF and Li-7 (Fig. 1D). In addition, TUNEL and Annexin V staining assay indicated that hypo-osmotic stimulation significantly inhibited the serum-deprivation-induced cell death (Fig. 1, E and F). These results strongly suggested that hypo-osmic stimulation suppresses serum-deprivation-induced apoptosis.

Figure 1.

Influence of hypo-osmotic condition on HepG2 cells survival. A: apoptosis was induced in HepG2 cells by serum deprivation and culturing in different osmotic medium for 48 h. Representative microscopic image of cells showing the influence of medium osmolarity is shown. BSA+: The culture medium contained 5% bovine serum albumin. Scale bar: 200 µm. Cell viability assay. Cell viability was examined by cell counting kit (B) or incorporation of tritiated thymidine (C). Data are expressed as means ± SD (experiments were performed in triplicate). **P < 0.01, *P < 0.05. D: cell viability assay using other hepatocellular carcinoma cell lines HLF and Li7. Data are expressed as means ± SD (experiments were performed in triplicate). **P < 0.01, *P < 0.05. E: image of TUNEL assay stain of HepG2 cells in isotonic or hypotonic culture medium. TUNEL (green), DAPI (blue), and merged images of the representative from three independent experiment are shown. Scale bar: 100 µm. The TUNEL-positive cells were counted in the five independent fields, respectively. E, right: quantification data is shown. F: flow cytometry analysis of apoptotic cells. Apoptosis-induced HepG2 cells under different osmotic culture conditions were stained with annexin V, a marker of apoptosis and the rate of apoptotic cells were counted by flow cytometry (left). The quantification data is shown on the right and expressed as means ± SD (experiments were performed in triplicate). TUNEL, terminal deoxyribonucleotidyl transferse (TdT)-mediated biotin-16-dUTP nick-end labelling.

Hypo-Osmotic Condition Upregulated Bcl-2 Protein Level

Since it was reported that the protein level of Bcl-2, known as antiapoptotic protein, was reduced during serum deprivation-induced apoptosis (23, 24), we next analyzed the influence of hypo-osmotic condition on Bcl-2 protein levels during apoptosis. As shown in Fig. 2A, serum deprivation reduced Bcl-2 protein levels, in concomitant with the activation of proapoptotic caspase 3, a downstream molecule of Bcl-2 pathway. In contrast, hypo-osmotic condition inhibited the downregulation of Bcl-2, resulting in the suppression of caspase 3 activation. To confirm whether this effect is dependent on low sodium concentration or hypo-osmolarity, a similar experiment was conducted using other osmolarity regulators such as potassium chloride, maltose, and sucrose (25, 26). These hypo-osmotic stimulations upregulated Bcl-2 protein level inhibiting the apoptosis (Fig. 2, B and C). These results suggested that the apoptotic resistance was caused by osmotic pressure but not by low sodium condition.

Figure 2.

Hypo-osmotic condition inhibits serum deprivation-induced apoptosis. Influence of hypo-osmotic condition on the apoptosis pathway in HepG2 cells. A: apoptosis was induced in HepG2 cells by serum deprivation culture medium with the indicated osmosis for 48 h (isotonic: 330 mosmol/kg, hypotonic: 270 mosmol/kg). The expression of Bcl-2, an antiapoptotic molecule, and proapoptotic protein, caspase 3 were determined by Western blot. B and C: effect of osmotic substances other than sodium chloride on HepG2 cells survival. HepG2 cells were cultured with potassium chloride (KCl), maltose, or sucrose-adjusted cultured medium for 48 h. B: expression of Bcl-2 was investigated by Western blot. C: cell viability was analyzed using cell counting kit. The quantification of Western blot bands was normalized to GAPDH. Data are expressed as means ± SD (experiment was performed in triplicate). **P < 0.01, *P < 0.05. Isotonic: 330 mosmol/kg, hypotonic: 300 or 270 mosmol/kg.

Hypo-Osmotic Condition Activated AKT Signaling, Leading to the Upregulation of Bcl-2

Next, to verify which signal causes apoptosis resistance, we investigated the upstream signaling of Bcl-2. As shown in Fig. 3A, hypo-osmotic stimulation enhanced only AKT phosphorylation in the tested signal pathway. We next investigated whether the enhanced AKT phosphorylation is related to an increase in the Bcl-2 protein level. The addition of triciribine, an AKT inhibitor, attenuated hypo-osmolarity-induced upregulation of Bcl-2 protein level and diminished the apoptosis resistance (Fig. 3, B and C). Similar results were also observed in the culture with maltose-adjusted hypo-osmotic medium (Fig. 3D).

Figure 3.

Hypo-osmotic condition activates AKT signaling, leading to the upregulation of Bcl-2. A: HepG2 cells were starved in serum-free isotonic medium (330 mosmol/kg) for 6 h. Then culture medium was changed to different osmolarity medium. Cell survival-related pathway such as AKT, NF-κβ, p38, and ERK1/2 at the indicated time points were investigated by Western blot. B and C: influence of AKT inhibitor on hypotonic-induced apoptosis resistance. Apoptosis was induced in HepG2 cells by serum deprivation and culturing in different osmolarity medium in the presence or absence of AKT inhibitor triciribine (1 µM). The cell viability and the expression of Bcl-2 is shown in B and C, respectively. *P < 0.05. Data are expressed as means ± SD (experiments were performed in triplicate). Quantification of Western blot bands of pAKT was normalized to total AKT and Bcl-2 density was normalized to GAPDH. D: confirmation of hypotonic-induced upregulation of Bcl-2 expression through AKT pathway using other osmolarity-regulating substance, maltose. HepG2 cells were cultured in serum-free isotonic or hypotonic medium for 48 h in the presence or absence of AKT inhibitor. The phosphorylation of AKT and the expression of Bcl-2 were examined by Western blot. The quantification of Western blot bands of pAKT was normalized to total AKT and Bcl-2 density was normalized to GAPDH. E: analysis of the influence of AKT activation on Bcl-2 expression. HepG2 cells that was transfected with constitutively activate AKT plasmid were cultured in the presence or absence of bovine serum albumin (BSA) for 48 h. The expression of apoptosis-related protein such as Bcl-2, Bax, and p53 was analyzed by Western blot. Iso, isotonic medium (330 mosmol/kg); Hypo, hypotonic medium (270 mosmol/kg).

In addition, as shown in Fig. 3E, the constitutive activation of AKT increased the Bcl-2 protein level, whereas other cell survival-related protein levels such as p53 and Bax were not affected. Under apoptotic conditions, the Bcl-2 protein level was decreased in control cells but was maintained by constitutive AKT activation. These results suggested that the activation of AKT signaling directly increased the Bcl-2 protein level, followed by the prolongation of cell survival in HCC cells.

The Hypo-Osmotic Condition Induced Apoptosis Resistance via TRPV2

TRPV2 deletion leads to downregulation of AKT signaling (15, 27). Therefore, we hypothesized that the elevation of TRPV2 expression level causes the resistance to apoptosis of HCC cells in hypo-osmotic conditions. We investigated the TRPV2 expression using CCl4-induced chronic hepatitis mouse model. The TRPV2 expression increased in liver by CCl4 administration (Fig. 4, A and B). In addition, as shown in Fig. 4C, hepatocytes around the central vein expressed TRPV2 in CCl4 group, whereas little TRPV2 expression was observed in hepatocytes of vehicle group. As shown in Figs. 4, D–F, TRPV2 silencing or the addition of TRPV2 antagonist, tranilast weakened hypo-osmotic stimulation-induced apoptotic resistance in HepG2 cells. Moreover, in mouse primary hepatocytes, which have little TRPV2 expression, the hypo-osmotic condition rather promoted apoptosis (Fig. 4G). However, TRPV2 overexpression in the mouse primary hepatocytes inhibited apoptosis (Figs. 4H). These results suggested that hypo-osmotic stimulation suppressed apoptosis via TRPV2.

Figure 4.

TRPV2 regulates the apoptosis resistance under hypo-osmolality. A–C: expression of TRPV2 in liver in CCl4-induced chronic liver inflammation model. Expression of TRPV2 mRNA (A) and protein (B) in liver was analyzed by qPCR and Western blot, respectively. A: data are expressed as means ± SD (experiment was performed in triplicate). **P < 0.01. B, right: quantification of three samples. Data are expressed as means ± SD. **P < 0.01. C: immunohistochemical staining for TRPV2 is shown. Scale bar: 100 µm. D and E: influence of TRPV2 silencing on hypotonic-induced apoptosis resistance in HepG2 cells. Apoptosis was induced in control or TRPV2 knockdown HepG2 cells by serum deprivation for 48 h. Downregulation of TRPV2 was confirmed by Western blot (D, top). Representative image is shown in D, bottom, and cell viability assay is shown in E. Data are expressed as means ± SD (experiment was performed in triplicate). *P < 0.05. Scale bar: 200 µm. F: blocking effect of TRPV2 on apoptosis resistance. Apoptosis was induced in HepG2 in the presence or absence of tranilast, a TRPV2 antagonist for 48 h. Then, the cell viability was examined. Data are expressed as means ± SD (experiment was performed in triplicate). **P < 0.01. G and H: influence of TRPV2 expression on apoptosis resistance in normal mouse hepatocytes. Primary hepatocytes that have no TRPV2 expression were isolated from C57BL/6J mice and cultured with indicated osmolarity medium for 48 h (G). Apoptosis was induced by serum deprivation in control or TRPV2 overexpressed mouse hepatocytes cultured in hypotonic medium (300 mosmol/kg) for 48 h. Cell viabilities were determined are expressed as means ± SD (H, experiment was performed in triplicate). **P < 0.01, *P < 0.05. CV, central vein; PV, portal vein; TRPV2, transient receptor potential vanilloid 2.

Hypotonic-Induced AKT Signal Activation Enhanced the Bcl-2 Protein Level via TRPV2

We further investigated the relationship between Bcl-2 expression and TRPV2-mediated AKT signal activation. TRPV2 silencing diminished the hypo-osmotic condition-mediated AKT phosphorylation (Fig. 5A). In addition, TRPV2 agonist probenecid (28) also induced the activation of AKT signaling (Fig. 5B). Because TRPV2 activation leads to calcium influx (29), we next confirmed whether the AKT phosphorylation in HepG2 cells is mediated by calcium influx. Hypo-osmotic stimulation enhanced the AKT phosphorylation in the presence of calcium. On the other hand, the upregulation of AKT phosphorylation was attenuated in the absence of calcium even if under hypo-osmotic conditions (Fig. 5C). As shown in Fig. 5D, osmotic pressure inversely increased Bcl-2 protein levels, although its effect was diminished by the downregulation of TRPV2. In contrast, probenecid increased Bcl-2 protein levels and prolonged the cell survival under apoptotic conditions (Fig. 5, E and F). Moreover, overexpression of TRPV2 in Huh7 cells, which have lower TRPV2 expression than in HepG2 cells, led to the upregulation of Bcl-2 and increased resistance to apoptosis (Figs. 5, G–I). Supposed mechanism of apoptosis resistance by hyponatremia-induced hypo-osmotic condition is shown in Fig. 6.

Figure 5.

TRPV2 signal protects from the serum-deprivation-triggered suppression of AKT phosphorylation. A: control or TRPV2 silenced HepG2 cells were starved in isotonic serum-free medium (330 mosmol/kg) for 6 h. Subsequently, cultured medium was changed to isotonic (Iso:330 mosmol/kg) or hypotonic (Hypo: 270 mosmol/kg) medium. Phosphorylation of AKT at the indicated time points were examined. B: HepG2 cells were stimulated by probenecid, a TRPV2 agonist, and the phosphorylation of AKT was determined by Western blot. C: analysis of calcium dependence on AKT phosphorylation induced by hypotonic condition. HepG2 cells were starved in isotonic serum-free medium (330 mosmol/kg) for 6 h. Then, culture medium was changed to different osmolarity medium adjusted by PBS (+; calcium containing) or PBS (−), which have no calcium. phosphorylation of AKT was detected by Western blot. D: effect of TRPV2 silencing on Bcl-2 expression. Control or TRPV2 knockdown HepG2 cells were cultured in serum-free and osmolarity-adjusted medium for 48 h. Expression of Bcl-2 was examined by Western blot. Quantification of Western blot bands was normalized to GAPDH. E and F: correlation between TRPV2 activation and the expression of Bcl-2. Apoptosis was induced in HepG2 cells in the presence or absence of 0.5 or 2 µM probenecid, a TRPV2 activator for 48 h in serum-free isotonic medium (330 mosmol/kg). Western blot for Bcl-2 is shown in E and the cell viability is shown in F. G, H, and I: validation of the relationship between TRPV2 and Bcl-2 expression using other HCC cell line Huh7. G: expression of TRPV2 mRNA in Huh7 and HepG2 was examined by qPCR. Control or TRPV2-overexpressed Huh7 cells (TRPV2 O/E) was cultured in isotonic serum-free medium for 48 h. Then, the phosphorylation of AKT and Bcl-2 expression were investigated by Western blot. The quantification of Western blot bands was normalized to control. H: the result of cell viability assay is shown in I. Data are expressed as means ± SD (experiments were performed in triplicate). **P < 0.01, *P < 0.05. TRPV2, transient receptor potential vanilloid 2.

Figure 6.

Proposed mechanism of apoptosis resistance in cirrhosis-induced hyponatremia. In cirrhotic liver, hyponatremia-induced hypo-osmotic condition causes TRPV2 activation and calcium influx into hepatocellular carcinoma cells. By the enhancement of AKT signaling, the Bcl-2 expression was upregulated, leading to apoptosis resistance. TRPV2, transient receptor potential vanilloid 2.

DISCUSSION

We first revealed that hypo-osmolality induced apoptosis resistance via osmosis receptor, TRPV2-mediated AKT-Bcl-2 pathway. Serum deprivation has been used for the research of apoptosis induction. In addition to serum, BSA inhibited cell apoptosis (18, 20). In the current study, the BSA-containing buffer was used as a control to eliminate several factors derived from serum in cell survival signal.

Several reports have shown that AKT signaling activation in HCC enhances Bcl-2 expression and leads to tumor growth and is also associated with drug resistance (30–32). The cancer microenvironment is altered by mechanotransduction factors such as temperature, stiffness, and shear stress (33, 34). Our findings suggested that these mechanotransduction factors also activate AKT and increase Bcl-2 expression, thereby causing apoptosis resistance of cancer cells.

In contrast, hypo-osmotic conditions promoted apoptosis in normal hepatocytes with little expression of TRPV2. Since mice with hepatocyte-specific attenuation of apoptosis resistance show hepatic fibrosis at 3 mo old and have a high incidence of liver carcinogenesis within a year, apoptosis in normal hepatocytes is considered to cause liver oncogenesis (35). Considering our findings, a hypotonic environment in sinusoids contributes to hepatocarcinogenesis through the promotion of apoptosis of normal hepatocytes.

The stimulant that induces TRPV2 expression remains undetermined. Another group reported that H₂O₂-induced oxidative stress increased TRPV2 expression in HepG2 cells (36). Chronic liver inflammation leads to oxidative stress, which has been demonstrated as an important factor for liver carcinogenesis (37). In addition, we confirmed the elevation of TRPV2 expression in hepatocytes by the administration of CCl4, which cause liver damage due to oxidative stress (38). Thus, oxidative stress might contribute to hepatocarcinogenesis by inducing TRPV2 expression in patients with chronic hepatitis.

Higher expression of TRPV2 was associated with poor prognosis in patients with esophageal squamous cell carcinoma, urothelial carcinoma, and gastric cancer (39–41). In addition, overexpression of TRPV2 in bladder cancer cells leads to enhancement of migration and invasion (42). Therefore, the inhibition of TRPV2 activation is a potential target of cancer treatment. Actually, the effect of tranilast for cancer therapy has been demonstrated using in vitro or mouse models. Tranilast inhibited several cancer cell proliferation, such as glioma, gastric cancer, lung carcinoma, prostate cancer, pancreatic cancer, and breast cancer (43). However, with respect to the treatment of liver diseases, the therapy using tranilast is still controversial. Tranilast had hepatoprotective property in the CCl4-treated acute liver injury model (44) and ameliorates a dietary rat model of nonalcoholic steatohepatitis (45). In contrast, tranilast treatment for HepG2-xenografted SCID mice promoted tumor growth, whereas the treatment of probenecid, a TRPV2 activator, suppressed the size of tumor (46). TRPV2 is highly expressed in cirrhotic liver and well-differentiated liver cancer, but is low in normal liver, chronic hepatitis, and undifferentiated liver cancer, suggesting that it is difficult to determine when to use tranilast for the treatment of liver diseases. In addition, since TRPV2 plays important roles in cardiac, muscular, neuronal physiology, thermogenesis, and immune response (15, 47), anticancer drugs targeting TRPV2 may have strong side effects. Thus, there might be promising candidate molecules in the downstream of TRPV2 pathway.

Patients with hyponatremia have higher rates of hospitalization with spontaneous bacterial peritonitis, hepatic encephalopathy, and hepatorenal syndrome (5). In addition, hyponatremia in cirrhosis has been clearly described as an independent risk factor for mortality (48). Patients whose serum levels of sodium chloride are less than 130 mmol/L should be considered for treatment (5). Whereas several treatments such as correction of hypokalemia (49, 50), albumin infusion (51, 52), and vasopressin receptor antagonist (53) have been used to improve serum levels of sodium chloride, management of hyponatremia is still challenging. Further studies focusing on the suppression of TRPV2-mediated mechanotransduction pathway can be expected to help solve hyponatremia-related clinical problems in addition to liver carcinogenesis.

DATA AVAILABILITY

Data will be made available upon reasonable request.

GRANTS

This work was supported by the Japan Society for the Promotion of Science (JSPS) KAKENHI Grant Nos. JP17K15561 (to H.U.) and JP20K21907 (to T.M.). Osaka City University “Think Globally, Act Locally” Research Grant for Young Scientists (to H.U.), and “Glocal Hub of Wisdom and Wellness filled with Smile” (to T.M.) and Grant for Research Program on Hepatitis from the Japan Agency for Medical Research and Development (AMED) Grant No. 16fk0210104h0001 (to K.I.).

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the authors.

AUTHOR CONTRIBUTIONS

H.U., T.M., and K.I. conceived and designed research; H.U., M.M., S.K., H.Y., and K.Y. performed experiments; H.U., T.M., S.K., and K.Y. analyzed data; H.U., T.M., K.Y., and K.I. interpreted results of experiments; H.U. prepared figures; H.U., T.M., M.M., S.K., K.Y., and K.I. drafted manuscript; H.U., T.M., S.K., and K.I. edited and revised manuscript; H.U., T.M., M.M., S.K., H.Y., K.Y., and K.I. approved final version of manuscript.