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. 2026 Feb 16;30(1):235–249. doi: 10.1080/19768354.2026.2629066

ATP release mediated by TRPM3 enhances invasion in glioblastoma

Kkot Garam Bae a, Chae Won Song a, Jae Hong Yoo a, Myunghoon Lee a, Noah Koo a, Gangsan Lee a, Yongmin Mason Park b, Jihwan Yoo c, In-Young Hwang b, Dong Ho Woo d, C Justin Lee b,✉, Kyung-Seok Han a,CONTACT
PMCID: PMC12912218  PMID: 41710303

ABSTRACT

Glioblastoma (GBM) is the most aggressive and lethal form of primary brain tumor, characterized by uncontrolled proliferation and invasion into surrounding brain tissue. Mechanical stimulation (MS) in the tumor microenvironment (TME) has been correlated to tumor progression, partly via ATP release. However, the underlying molecular mechanisms remain poorly understood. In this study, we found that transient receptor potential melastatin 3 (TRPM3) channel mediates MS-induced ATP release from GBM cells. Genetic knockdown of TRPM3 significantly attenuated ATP release and suppressed GBM cell invasion, indicating its functional relevance in tumor dissemination. Furthermore, TRPM3 regulated ATP release in a Ca2+-independent manner, suggesting a noncanonical mechanism of mechanosensitive signaling. Consequently, targeting TRPM3 may offer a novel target to reduce the invasion of GBM within the TME.

KEYWORDS: Glioblastoma, ATP, mechanical stimulation, TRPM3, invasion

Introduction

Glioblastoma (GBM), the most common and aggressive primary malignant brain tumor classified as WHO grade 4, is associated with a poor prognosis, with a median survival of less than one year following diagnosis (Guo et al. 2023; Qi et al. 2023; Kim HJ et al. 2024; Lee et al. 2024). Even with treatment including surgical tumor resection, chemotherapy with temozolomide, and radiation therapy, overall survival remains poor (Johnson and O’Neill 2012). GBM exhibits rapid proliferation and extensive invasion into surrounding brain tissue (Koh et al. 2018; So et al. 2021b), partially due to responding to neurotransmitters and gliotransmitters from neighboring cells, as well as through autocrine signaling that modulates the tumor microenvironment (TME) (De Silva et al. 2023; Han et al. Sun et al. 2025).

A defining feature of GBM is aggressive invasiveness, which notably differentiates it from most other tumors (So et al. 2021b). Compared to other tumors, GBM grows within the rigid cranial cavity where spatial constraints restrict tumor expansion (Luu et al. 2024). As a result, tumor invasion and proliferation contribute to increased intracranial pressure (ICP) (Purdy and Kirby 2004; Ueda et al. 2022). As GBM proliferates in the limited space, it leads to elevated intracranial (Ueda et al. 2022) and interstitial pressures (Hu et al. 2025). And it impairs normal brain function and lead to clinical symptoms such as severe headaches (Sorribes et al. 2019). The association between GBM invasiveness and ICP elevation is clinically significant, affecting both treatment strategies and patient prognosis (Grossen et al. 2022). Previous studies have implicated elevated intracranial pressure as a contributing factor to key processes driving tumor progression (Sorribes et al. 2019; Ahmed et al. 2021; Shukla et al. 2024).

ATP functions as a signaling molecule in various cell types. In many tumor types, including GBM, ATP has been implicated to promote both metastasis and invasion. Accordingly, extracellular ATP levels in the tumor microenvironment are reported to range between 100 and 500 μM, significantly higher than those in normal tissues (Pellegatti et al. 2008; Di Virgilio and Adinolfi 2017). However, the relationship between elevated ICP and ATP in patients remains largely unknown.

In contrast, the mechanisms of ATP release are generally divided into exocytotic and non-exocytotic pathways. Non-exocytotic ATP release mechanisms include hemichannels, volume-regulated anion channels (VRAC), ATP release during cell death, and P2X7-mediated ATP release. In GBM, Connexin 43 (Cx43), a type of hemichannel, is significantly overexpressed and has been reported to contribute to tumor growth and invasion (Martins et al. 2014; Johnsen et al. 2019; Okada et al. 2020; Zanoni et al. 2022; Fidaleo et al. 2026). VRAC function relies on the leucine-rich repeat-containing protein 8A (LRRC8A) subunit, which is also overexpressed in GBM and known to promote tumor progression (Okada et al. 2020; Fidaleo et al. 2026). ATP also can be released during immunogenic or apoptotic cell death in GBM (Martins et al. 2014). Furthermore, the P2X7 receptor (P2X7R), expressed in tumor cells, can function as a channel when activated. This enabling direct ATP efflux (Johnsen et al. 2019). Additionally, activation of the P2X7R can open Pannexin 1 (Panx1) channels, facilitating ATP release (Johnsen et al. 2019). Once released, ATP binds to P2 purinergic receptors on other cells, particularly P2X7R, mediating intercellular interactions and playing a vital role in cellular signaling (Di Virgilio et al. 2018; Zanoni et al. 2022). The P2X7R is known to promote GBM growth, suggesting that ATP binding to P2X7R may critically contribute to GBM progression. In addition to these pathways, ATP release can also be induced by mechanical stimulation (MS) in a Ca2+-dependent manner in astrocytes (Lee et al. 2015).

Increased intracranial pressure (ICP) is detected by mechanosensitive ion channels, which respond to mechanical stimuli such as stretch or compression by initiating intracellular signaling through ion flux. Previous studies have suggested that the TRP (Transient Receptor Potential) superfamily exhibits mechanosensitivity (Plant 2014). TRP channels are non-selective ion channels, initiating intracellular cascade reactions (Zhang et al. 2023; Du et al. 2024). This enables cells to sense external stimuli such as temperature, chemical, and mechanical stimuli (Liu et al. 2024). Importantly, the activation of TRP channels has been noted to affect ATP release (Egbuniwe et al. 2014; Rahman et al. 2018), suggesting a potential mechanistic link between TRP activity and extracellular purinergic signaling. Within the TRP superfamily, the TRPM (melastatin) subfamily plays key roles in regulating cell proliferation and survival in response to diverse stimuli (Miller and Cheung 2016). Notably, members of the TRPM subgroup, including TRPM3, TRPM7, and TRPM8, show high expression levels in GBM patients (Alptekin et al. 2015) and are linked to GBM progression and patient survival (Chen et al. 2024). These findings suggest that TRPM channels may contribute to ATP release in response to mechanical stimulation and play a functional role in GBM pathophysiology.

In this study, we demonstrated that MS induces ATP release in GBM cells including U-87 MG, U-373 MG, patient-derived GBM. Our results demonstrate that MS-induced ATP release occurs through a Ca²+-independent mechanism. Pharmacological screening and gene-silencing experiments identified TRPM3 as a key regulator of this process. Furthermore, TRPM3-mediated ATP release was found to enhance GBM cell invasion. These findings suggest that GBM cells dynamically regulate extracellular ATP levels in the TME in response to external mechanical stress, highlighting TRPM3 as a potential therapeutic target for invasion in GBM.

Materials and methods

Cell culture

The human glioblastoma cell line U-87 MG and U-373 MG was obtained from the American Type Culture Collection (ATCC, #HTB-14) and Korean Cell Line Bank (#30017). Cells were maintained in Dulbecco's Modified Eagle's Medium (DMEM; CORNING, #10-013-CVRC) supplemented with 10% fetal bovine serum (FBS; Gibco, #16000-044) and 1% penicillin–streptomycin solution (10,000 units/mL penicillin, 10,000 μg/mL streptomycin; Cytiva, #SV30010). Cultures were incubated at 37 °C in a humidified atmosphere containing 5% CO2.

Patient-derived GBM culture

pdGBM medium was prepared by combining DMEM/F12 (Gibco, NY, USA, 11320-033) and Neurobasal Medium (Gibco, NY, USA, 21103049) in a 1:1 ratio, supplemented with 1X GlutaMAX™ (Thermo Fisher Scientific, 35050061), B27 supplement minus Vitamin A (Gibco, NY, USA, 12587-010), N2 supplement (Gibco, NY, USA, 17502-048), 1X MEM Non-Essential Amino Acids Solution (Thermo Fisher Scientific, 11140050), 1% Penicillin–Streptomycin (Thermo Fisher Scientific, 15140122), and human insulin solution (Sigma, I9278). Tumor fragments obtained from dissection were cultured in this medium using 6-well low-attachment plates. The cultures were maintained at 37 °C with 5% CO₂ under continuous agitation at 120 rpm on an orbital shaker. Every two days, three-quarters of the medium was replaced with fresh pdGBM medium. pdGBMs that developed a round morphology were transferred to an FBS-containing medium to facilitate attachment to the plate before further experimentation.

ATP imaging

GBM cells were transfected with pAAV-GfaABC1D-GRABATP1.0-GFP (Addgene, #167579). Cells were then cultured on 12 mm Ø cover glasses (Paul Marienfeld, Lauda-Königshofen, Germany, #0111520) coated with poly-D-lysine (Sigma-Aldrich, MO, USA, #P6403). Cover glasses were mounted in a chamber for imaging, and perfused with an external solution consisting of 150 mM NaCl (Sigma-Aldrich, #S9888), 10 mM HEPES (Carl Roth, #230-907-9), 2.3 mM KCl (Sigma-Aldrich, #P9541), 2 mM CaCl2 (Sigma-Aldrich, #C3881), 2 mM MgCl2 (Sigma-Aldrich, #M2670), 22 mM sucrose (Carl Roth, #200-334-9), and 5.6 mM glucose (Sigma-Aldrich, #G8270) (pH 7.4). The pharmacological agents, including 200 μM 2-APB (Tocris, #1224), 20 μM SKF-96365 hydrochloride (Tocris, #1147), 40 μM HC-030031 (Tocris, #2896), 100 μM quinine hydrochloride (Tocris, #4114), 1 μM carbenoxolone disodium (Tocris, #3096), and 10 μM capsazepine (Tocris, #0464), were dissolved in the external solution. Before ATP imaging, cells were incubated with these pharmacological agents for 10 min. Fluorescence imaging was conducted using a Nikon TE300 fluorescence microscope equipped with a HAMAMATSU camera (#C11440-36U). MS was applied using a glass pipette (resistance: 4–7 MΩ) filled with an internal solution identical to the external solution. A glass pipette was securely positioned in the pipette holder, and MS was applied using a micromanipulator (Siskiyou, #MC 1000e) to precisely control pressure. Pressure was gradually increased until the pipette resistance reached twice its initial value, ensuring controlled MS. ATP release was quantified using a GFP filter (49054 ET-GFP for 455–465 nm LEDs), and fluorescence intensity was measured. Data acquisition and analysis were performed using ImageWorkBench software (9.0.14.0). To regulate extracellular Ca²+ influx, a Ca²+-free buffer was prepared, consisting of 150 mM NaCl, 10 mM HEPES, 2.3 mM KCl, 2 mM MgCl₂, 22 mM sucrose, 5.5 mM glucose, and 200 mM ethylene glycol-bis(β-aminoethyl ether)-N,N,N′,N′-tetraacetic acid (EGTA, Sigma-Aldrich, #E0396). To evaluate ATP levels in co-expressing cells, shRNA-mCherry was transduced, followed by GRAB sensor expression after 48 h. Cover glasses were subsequently mounted, and ATP imaging was performed using the established imaging setup. Fluorescence intensity was selectively analyzed in mCherry-positive cells.

Ca2+ imaging

To assess changes in intracellular Ca²+ levels following MS, GBM cells transfected with GRABATP-GFP were cultured on poly-D-lysine-coated 12 mm Ø cover glasses. Cover glasses were incubated in 1 mL external solution containing 5 μM Fura-2 AM (ThermoFisher, #F1201) and 1 μM Pluronic F-127 (Sigma-Aldrich, #P2443) for 40 min at room temperature in the dark. Cover glasses were transferred to fresh HEPES buffer until experimentation. Fluorescence was detected using a microscope equipped with a 79001 ET-Fura 2 filter (CHROMA, USA, #79001). Thapsigargin (1 μM; TOCRIS Bioscience, #1138) dissolved in Ca2+-free buffer was rapidly perfused into the bath. The pE-340fura light source was set at 36 W, emitting 340 and 380 nm for Ca2+ imaging. Fluorescence intensity for each cell was normalized using the 340/380 ratio, calculated as ΔFF0=(V−AVRBase)AVRBase, where V represents the fluorescence intensity at each time point and Average Base is the average baseline intensity before TG application. Once Ca2+ returned to baseline, MS was applied to compare ATP release between treated and untreated conditions.

Cloning & lentiviral packaging (infection)

Scrambled shRNA and TRPM3, M7, M8 shRNA sequences were designed and synthesized by Cosmo Genetech (Daejeon, Korea). The sequences of shRNA are as follows:

Scrambled shRNA sequences: 5' – TCGCATAGCGTATGCCGTTTT-3’

TRPM3 shRNA sequence: 5'-GCATGCATTCCCACTTCATTC-3’

TRPM7 shRNA sequence: 5'-CCACCAAAGAATCAGAATCAA-3’

TRPM8 shRNA sequence: 5'-GCAGGAGTTCAGAGGAGAATA-3’

The pSicoR-mCh-empty vector (Addgene, #21907) was digested using XhoI (BioLabs, #R0146S) and HpaI (BioLabs, #R0105S) with rCutSmart buffer (BioLabs, #B6004S). DNA purification was performed using the Labopass Gel and PCR Clean-up Kit (Cosmo Genetech, #CMA0112). shRNA oligonucleotides were annealed, phosphorylated, and ligated into the vector using NEB buffer (BioLabs, #B7002S), T4 polynucleotide kinase (Enzynomics, #M005S), ATP (Sigma, #A26209), T4 DNA ligase (Enzynomics, #M001S), and T4 DNA ligase buffer (Enzynomics, #B304). Ligated constructs were transformed into DH5α-competent cells (Enzynomics, #CP011), and plasmid DNA was extracted using the LaboPass Plasmid Mini Kit (Cosmo Genetech, #CMP0112). Sequencing confirmation was performed by Cosmo Genetech. Lentivirus production was conducted using HEK 293 T cells transfected with 0.75 μg psPAX2 (Addgene, MA, USA, #12260), 0.25 μg pMD2.G (Addgene, MA, USA, #12259), and 1 μg pSicoR-shRNA-mCh vector using Lipofectamine 3000 (Thermo Fisher Scientific, MA, USA, #L3000001). Viral supernatants were collected every 24 h after reaching 80% confluence and stored at 4 °C. The collection was repeated for three consecutive days, and polybrene (Merck Millipore, #TR-1003-G) was added at a concentration of 1 μL/mL. Viral supernatants were aliquoted and stored at – 80 °C. GBM cells were infected by replacing 1 mL of media with 1 mL of viral supernatant. After 24 h, the medium was completely replaced with fresh culture medium.

qRT-PCR

Total RNA was isolated 72 h following shRNA lentiviral transduction using the RNeasy Mini Kit (QIAGEN, Hilden, Germany; #74106). For each sample, 1,000 ng/μL of RNA was reverse transcribed into cDNA employing the SuperScript III First-Strand Synthesis System (Invitrogen, MA, USA, #18,080–051) and the MiniAmp Thermal Cycler (Thermo Fisher Scientific, MA, USA, #A37834), resulting in a final cDNA concentration of 50 ng/μL. Quantitative real-time PCR (qRT-PCR) was conducted with PowerSYBR Green PCR Master Mix (Applied Biosystems, MA, USA, #4367659) on a CFX96 Optics Module (Bio-Rad, CA, USA). Gene expression levels were normalized to those of the shSC control group. The primers used in the qRT reaction were as follows.

TRPM3 forward: 5'-CAT GCA CCG TTT TCT CAC CA-3’

TRPM3 reverse: 5'-ATC ACC AGG CCG ATG TCA AT-3’

TRPM7 forward: 5'-CAG AAA CCA AGC GCT TTC CT-3’

TRPM7 reverse: 5'-ACG GCC AAC TGA CCA AAA TC-5’

TRPM8 forward: 5'-CCC TTC CCC TTC ATC GTC TT-3’

TRPM8 reverse: 5'-TTC CTT CAT GAC ACC CTC CC-3’

Human GAPDH forward: 5′-GTC TCC TCT GAC TTC AACAGCG-3′

Human GAPDH reverse: 5′-ACC ACC CTG TTG CTG TAG CCAA-3′

Sniffer-patch

Sniffer-patch recordings were performed to measure the full activation current of P2X2-V343Q-IRES-GFP in HEK 293 T cells in response to ATP application and ATP release from mechanically stimulated U-87 MG cells. HEK 293 T cells were transfected with P2X2-V343Q-IRES-GFP using Buffer EC, Effectene, and Enhancer Reagent (Qiagen). After 24 h, the culture medium was replaced, and naïve U-87 MG cells were seeded onto a cover glass in a 24-well plate. On the day of the experiment, transfected HEK 293 T cells were added to the cover glass for 4 h of co-culture. The cover glass was then transferred to the recording chamber, and HEK 293 T cells adhering to U-87 MG cells were identified using an Olympus BX51WI microscope. Expression of P2X2-V343Q-IRES-GFP was confirmed with pE-300 lite (CoolLED), and cellular images were acquired using HCImage Live (Hamamatsu). Patch-clamp recordings were conducted using either an external solution or a Ca²+-free buffer. Borosilicate glass pipettes (4–7 MΩ) were filled with an internal solution containing 135 mM Cs-methanesulfonate, 0.2 mM EGTA, 10 mM HEPES, 10 mM KCl, and 1 mM MgCl₂ (pH 7.3). ATP (100 μM, Sigma-Aldrich, #A9187) was perfused until a steady-state response was reached (Vh = −70 mV). MS was applied to U-87 MG cells using a glass pipette until resistance doubled, following the same protocol described in ATP imaging. Patch-clamp currents were recorded with a Multiclamp 700B amplifier (Axon Instruments) and Clampex 11.2 software (Molecular Devices) and filtered at 2 kHz.

Wound healing assay

U-87 MG cells infected with scrambled or TRPM3 shRNA were cultured to confluence, and a scratch was made using a 200-μL pipette tip. Images were acquired at 100× magnification using a microscope (KOREA LAB TECH, #KCS3 160S, #KI-400) immediately after scratching (0 h) and after 24 h. Cell migration was quantified using ImageJ software by measuring wound closure over time.

Invasion assay

U-87 MG cells infected with scrambled or TRPM3 shRNA were seeded in Transwell plates (SPL, #35224) pre-coated with Vitrogel hydrogel matrix (The Well Bioscience, #VHM01). The upper chamber contained medium with 2% serum, while the lower chamber contained medium with 20% serum to drive cell invasion. After 72 h, cells were detached with 0.05% Trypsin-EDTA (Gibco, #25300-054), fixed with 3.7% formaldehyde (Sigma-Aldrich, #SMB00907), permeabilized with methanol (Samchun Chemicals, #M0588), and stained with DAPI (Thermo Scientific, #62248). Images were acquired at 100× magnification using a fluorescence microscope (Olympus, #CKX53, #KCS3-160S). Three random fields per well were analyzed to quantify invasive capacity.

Statistical analysis

Image analysis was performed using ImageJ (Ver. 1.53k). Statistical analyses, including bar graphs and trace plots, were conducted using GraphPad Prism (Ver. 9.4.1). For comparisons between two groups, a two-tailed unpaired Student’s t-test was used. All data are presented as the standard error of the mean (SEM). Statistical significance was indicated as follows: P < 0.05 (*), P < 0.01 (**), P < 0.001 (***), P < 0.0001 (****), and ‘ns’ denoted non-significant differences.

Results

To investigate whether ATP is released in GBM in response to pressure changes within the tumor microenvironment, we performed ATP imaging using the fluorescent sensor protein GRABATP1.0 (Figure 1A, B) (Wu et al. 2022). To mimic tumor-associated mechanical forces at the single-cell level, we applied MS to the cell (Forsyth and Posner 1993; Park et al. 2018). Notably, MS-induced ATP release was observed in U-87 MG cells (Figure 1C, D). Given that mechanosensitive channels are known to modulate downstream signaling pathways, including those involving G-protein-coupled receptors (GPCRs), we next tested whether GPCR activation contributes to ATP release. To this end, we treated cells with TFLLR-NH₂, a PAR1 (protease-activated receptor 1) agonist. However, activation of GPCRs did not induce ATP release (Figure 1C, D). These results indicate that ATP release in GBM cells is specifically triggered by mechanical stimulation, not by GPCR activation. To further validate ATP release from GBM cells, we employed the sniffer-patch using U-87 MG and HEK293 T cells (Figure 1E). HEK293 T cells were transfected with P2X2-V343Q, a non-selective cation-permeable channel, and subsequently co-cultured with U-87 MG cells. Whole-cell patch-clamp recordings were conducted in HEK293 T cells expressing the P2X2-V343Q, and the corresponding currents were measured upon MS of adjacent U-87 MG cells (Figure 1F). The inward current recorded in HEK cells is mediated by ATP released from nearby U-87 MG cells. (Figure 1F). Following each recording, 100 μM ATP was bath-applied to assess the full extent of P2X2-V343Q expression, and the corresponding values were quantified (Figure 1G). These results provide converging evidence that MS induces ATP release, as demonstrated using two independent experimental approaches.

Figure 1.

Figure 1.

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ATP is released in U-87MG cells by mechanical stimulation (MS), but not by GPCR activation (A) Schematic illustration of ATP release induced by MS in U-87 MG cells expressing the GRABATP sensor. (B) Fluorescence images of U-87 MG cells expressing GRABATP before (top) and after (bottom) MS. Scale bar = 100 μm. (C) Representative ATP traces in U-87 MG cells. The top trace shows ATP release induced by TFLLR puffing (triangle), while the bottom trace shows ATP release induced by MS (diamond). (D) Bar graph comparing ATP responses (ΔF/F₀) between TFLLR puffing and MS (n = 8 per condition). (E) Schematic illustration of the sniffer-patch technique using co-cultured U-87 MG cells and HEK 293 T cells expressing P2X2-V343Q-IRES-GFP. (F) Representative current traces recorded using the sniffer-patch. Inward currents induced by ATP release from U-87 MG cells (Right). Full activation current recorded from a HEK 293 T cell after bath application of 100 μM ATP (Left). (G) Bar graph showing ATP responses (% of full activation). (H) Scatter plot showing ATP response (ΔF/F0) versus pipette resistance (n = 20). The red line indicates the linear fit (R2 = 0.0007). (I) Scatter plot of ATP response (ΔF/F0) as a function of relative pipette resistance change (%) before and after MS. The red line represents the linear regression fit (R2 = 0.01615). Data are presented as mean ± SEM. (D) Two-tailed unpaired t-test. (H, I) Simple linear regression. Each dot represents ATP response of an individual cell. ns > 0.05, *p < 0.05.

To elucidate whether MS-induced ATP release is mediated by intracellular calcium signaling, we examined whether the source of Ca²+ was extracellular influx or release from the endoplasmic reticulum (ER). First, to block extracellular Ca²+, cells were treated with a Ca²+-free buffer containing EDTA for 10 min prior to mechanical stimulation. ATP release was still observed under these conditions, indicating that extracellular Ca²+ is not required for MS-induced ATP release (Figure 2A, B). We next tested whether ER-derived Ca²+ contributes to this process, based on the prediction that mechanical stimulation might affect Ca²+ from intracellular stores. To deplete intracellular Ca²+ from ER, cells were pretreated with 1 μM TG in a Ca²+-free buffer for 10 min before MS was applied (Figure 2C) (Sehgal et al. 2017). ATP release persisted even under ER Ca²+-depleted conditions (Figure 2D), further suggesting a Ca²+-independent mechanism. To further validate this finding, we performed sniffer-patch recordings. Co-cultured cells were incubated in a Ca²+-free buffer for 10 min, and whole-cell patch-clamp recordings were conducted in HEK293 T cells (Figure 2E). Consistent with imaging results, MS-induced ATP release elicited a measurable current, with the full activation current reaching approximately 60% (Figure 2F, G). This value did not significantly differ from that observed under normal condition (Figure 1G). Together, these results demonstrate that MS-induced ATP release in GBM cells occurs independently of both extracellular and intracellular Ca²+ sources. Additionally, to assess whether the MS intensity affects ATP release, we compared ATP release across different pipette resistances as well as changes in pipette resistance (Figure 1H, I). Our results showed that neither absolute pipette resistance nor changes in resistance correlated with the amount of ATP released, suggesting that ATP release occurs independently of the intensity of mechanical stimulation.

Figure 2.

Figure 2.

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ATP release induced by MS is independent of Ca2+ in U-87 MG cells (A) Representative ATP traces in normal Ca²+ condition (Left) and extracellular Ca²+-free condition (right). (B) Bar graph showing ATP responses (ΔF/F₀) between control and extracellular Ca2+-free condition (n = 6 per condition). (C) Representative ATP trace in extracellular Ca²+-free conditions (Left). Representative intracellular Ca²+ trace following treatment with 1 μM TG (middle). Representative ATP trace after TG treatment in extracellular Ca²+-free conditions (Right). (D) Bar graph showing ATP responses (ΔF/F₀) between control and TG-treated cells in Ca2+-free condition (n = 10 per condition). (E) Fluorescence image of HEK 293 T cells expressing P2X2-V343Q (left) and DIC image showing U-87 MG cells (white arrow) adjacent to HEK 293 T cells (red arrow) (right). (F) Representative current of extracellular Ca2+-free traces recorded using the sniffer-patch (Right). Full activation current recorded from a HEK 293 T cell after bath application of 100 μM ATP (Left). (G) Bar graph showing ATP responses of Ca²+-free condition (% of full activation). Each dot represents ATP response of an individual cell. Data are shown as mean ± SEM. (B, D) Two-tailed unpaired t-test. ns > 0.05.

TRP channels are implicated in cellular survival and invasion and are known to be activated by various stimuli, including mechanical force (Shapovalov et al. 2011). Since MS induces mechanical force, we investigated its involvement in ATP release by pharmacologically inhibiting specific TRP channels. ATP release was assessed following treatment with 2-aminoethoxydiphenyl borate (2-APB), an inhibitor of IP3 receptors and several TRP channels, including TRPC1, TRPC3, TRPC5, TRPC6, TRPV6, TRPM3, TRPM7, TRPM8, and TRPP2 (Figure 3A, purple). Preincubation with 2-APB for 10 min completely abolished MS-induced ATP release, whereas treatment with SKF-96365, a TRPC channel blocker, had no additional effect (Figure 3A, orange).

Figure 3.

Figure 3.

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MS-induced ATP release is mediated by TRPM3 in GBM (A) Representative ATP traces recorded from naïve cells and cells pretreated with 200 μM 2-APB, 20 μM SKF-96365, 40 μM HC-030031, 100 μM quinine, 1 μM CBX, or 10 μM capsazepine for 10 min before MS. (B) Bar graph showing ATP responses (ΔF/F₀) under each condition (2-APB, n = 10; SKF-96365, n = 6; HC-030031, n = 16; Quinine, n = 4; CBX, n = 4; capsazepine, n = 8). (C) Schematic of the vector constructs. (D) Relative mRNA expression levels of TRPM3, TRPM7, and TRPM8 in U-87 MG cells after shRNA-mediated knockdown, normalized to shSC. (E) Fluorescence images showing mCherry, GFP, and merged signals in U-87 MG cells infected with shTRPM3-mCherry. White arrows indicate representative cells. Scale bar = 50 μm. (F) Representative ATP traces recorded from U-87 MG (upper left), U-373 MG (lower left), and patient-derived GBM (lower right) cells infected with scrambled shRNA (shSC) or shTRPM3 #1. Representative ATP traces recorded from U-87 MG (upper, right) cells infected with shSC or shTRPM3 #2. (G) Bar graph comparing ATP responses (ΔF/F₀) in each cell line (U-87 MG infected shTRPM3 #1, n = 28; U-87 MG infected shTRPM3 #2, n = 14; U-373 MG infected shTRPM3 #1, n = 10; patient-derived GBM infected shTRPM3 #1, n = 14). (H) Representative ATP traces recorded from U-87 MG cells infected with shSC or shTRPM7 (left) and corresponding bar graph comparing ATP responses (ΔF/F₀) between shSC – and shTRPM7-infected cells (right) (n = 20 per condition). (I) Representative ATP traces recorded from U-87 MG cells infected with shSC or shTRPM8 (left) and corresponding bar graph comparing ATP responses (ΔF/F₀) between shSC – and shTRPM8-infected cells (right) (n = 10 per condition). Each dot represents ATP responses of an individual cell. Data are presented as mean ± SEM. (B, D) Ordinary one-way ANOVA (Multiple comparisons). (G) Multiple Mann-Whitney tests. (H, I) Two-tailed unpaired t-test. *p < 0.05, **p < 0.01, ****p < 0.0001.

Previous studies have demonstrated that ATP is released through TRPA1 in a Ca²+-dependent manner in odontoblasts (Egbuniwe et al. 2014). However, in GBM, ATP release was not entirely abolished by the selective TRPA1 antagonist HC-030031, despite a significant reduction (Figure 3B). Treatment with quinine, a Ca²+-activated K+ channel blocker, and capsazepine, a TRPV channel antagonist had no significant effect on ATP release. In contrast, partial inhibition was observed with carbenoxolone, a gap junction blocker, suggesting the involvement of additional pathways. Given that ATP release was suppressed by 2-APB-sensitive channels, we hypothesized that specific TRPM isoforms contribute to MS-induced ATP release. Previous studies have implicated TRPM channels in GBM pathophysiology, leading us to identify TRPM3, TRPM7, and TRPM8 as candidate channels potentially involved in ATP release.

To investigate the involvement of TRPM channels in MS-induced ATP release, we designed short-hairpin RNAs (shRNAs) targeting TRPM3, TRPM7, and TRPM8 (Figure 3C). The relative mRNA levels of shTRPM3, shTRPM7, and shTRPM8 were determined by qRT-PCR. When normalized to shSC, the knockdown efficiencies were 38% for shTRPM3, 91% for shTRPM3 #2, 82% for shTRPM7, and 72% for shTRPM8, respectively (Figure 3D). We co-expressed GRABATP sensors with each shRNA to evaluate ATP release following knockdown of TRPM3, TRPM7, and TRPM8 and measured MS-induced ATP only in cells co-expressing mCherry and GFP (Figure 3E). Knockdown of TRPM3 using two independent shRNA constructs resulted in a significant reduction of ATP release across all three GBM cell lines examined – U-87 MG, U-373 MG, and patient-derived GBM (Figure 3F, G). These indicate that MS-induced ATP release from GBM is mediated by TRPM3, whereas silencing TRPM7 or TRPM8 does not significantly affect ATP release (Figure 3H, I).

To examine the functional role of TRPM3 in MS-induced ATP release in GBM, we assessed cell migration and invasion. GBM cells were transfected with TRPM3-targeting shRNA, and migration and invasion assay were conducted five days post-infection (Figure 4A). The infection efficiency in U-87 MG cells was assessed by comparing DIC images with mCherry fluorescence images following shRNA infection (Figure 4B). After inducing artificial scratches, the migration rate was assessed 24 h later. However, TRPM3 knockdown did not result in a significant change in cell migration (Figure 4C). To assess the role of TRPM3 in invasion, a Vitrogel-based invasion assay was conducted to evaluate the ability of cells to traverse confined spaces. Knockdown of TRPM3 using shTRPM3 #1 and shTRPM3 #2 significantly decreased the number of invading cells (Figure 4D, E). To explore the downstream mechanisms underlying TRPM3-mediated invasion and migration, we examined gene expression in TRPM3 knockdown cells using qRT-PCR. Previous studies have implicated MMPs in invasive behavior and integrin α5, β1 in cell migration. Changes in RNA levels induced by shRNA did not correlate with differences in migration or invasion (Supplementary Fig. 1A). These findings suggest that TRPM3-driven invasion likely occurs through an alternative pathway independent of MMP activity (Sato and Takino 2010). In parallel, we determined whether exogenous ATP influences GBM invasion by performing invasion assays with ATP treatment. ATP treatment significantly increased the number of invading cells compared with the control group (Figure 4F). These findings suggest that TRPM3 contribute to ATP release and increases the invasive properties of GBM cells.

Figure 4.

Figure 4.

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Gene-silencing of TRPM3 reduces cell invasion in U-87 MG cells (A) Schematic timeline of the experimental design. (B) DIC image showing U-87 MG cells (top) and fluorescence image showing mCherry (bottom). Scale bar = 250 μm. Infection efficiency quantified by counting mCherry+/DIC+ cells (83.9%, 125/149 cells). (C) Representative images of the wound healing assay in shSC – and shTRPM3 #1-infected cells (right). Red lines indicate the cell boundary. Migration rate quantified as the ratio of the cell boundary at 24 h relative to the initial boundary at 0 h, measured using ImageJ (n = 7 per condition). (D) Representative images from the invasion assay in shSC – and shTRPM3 #1-infected U-87 MG cells, which were stained with DAPI (n = 12 per condition). Number of invaded cells quantified by counting DAPI-stained nuclei (n = 12 per condition). (E) Representative images from the invasion assay in shSC – and shTRPM3 #2-infected U-87 MG cells, which were stained with DAPI (left, shSC, n = 17; shTRPM3 #2, n = 6). Bar graph showing number of invaded cells quantified by counting DAPI-stained nuclei (right) (shSC, n = 17; shTRPM3 #2, n = 6). (F) Representative images from the invasion assay in shTRPM3 #2-infected U-87 MG cells with D.W and shTRPM3 #2-infected U-87 MG cells 10μM ATP, which were stained with DAPI (left). Bar graph showing number of invaded cells quantified by counting DAPI-stained nuclei (right) (shTRPM3 #2 with D.W, n = 6; shTRPM3 #2 with 10 μM ATP, n = 6). Data are presented as mean ± SEM. (C-F) Two-tailed unpaired t-test. ns > 0.05, *p < 0.05, **p < 0.01.

To validate the involvement of TRPM3 in glioma invasion, we analyzed its spatial expression pattern across tumor regions using the human IVY-GAP database (Figure 5A). TRPM3 expression was markedly elevated in the leading edge and infiltrating tumor areas, which are representative of highly invasive regions. In contrast, TRPM7 and TRPM8 did not exhibit increased transcript levels in these invasion-associated zones. Furthermore, there was no change in the relative expression level of TRPM3 between high-grade and low-grade gliomas, suggesting the possibility of MS-induced ATP release regardless of cancer grade (Figure 5B). Furthermore, to assess the clinical relevance of TRPM3, we evaluated patient survival based on TRPM3 alterations using multiple GBM cohorts (Figure 5C, D, E, F). Kaplan – Meier survival analyses consistently demonstrated a rightward shift in the TRPM3-altered group across four independent datasets. These findings align with our in vitro results and suggest that TRPM3-mediated invasion may be linked to ATP signaling, ultimately influencing patient prognosis (Figure 6).

Figure 5.

Figure 5.

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Spatial expression mapping and clinical significance of TRPM3 in glioblastoma revealed by IVY-GAP and TCGA datasets (A) Heatmap showing spatial expression patterns of TRPM3, TRPM7, and TRPM8 across glioblastoma microenvironments obtained from the IVY-GAP dataset. TRPM3 expression is enriched in the leading-edge and infiltrating-tumor regions. (B) Relative TRPM3 transcript levels (TPM) between high-grade and low-grade gliomas derived from GSE7554447. (C-F) Kaplan – Meier survival analyses of glioblastoma patients stratified by TRPM3 alteration status across multiple TCGA and publicly available datasets.

Figure 6.

Figure 6.

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Schematic image of MS-induced ATP release through TRPM3 (A) Schematic of TRPM3-mediated ATP release in response to MS in GBM cells. It demonstrates that TRPM3 mediates MS-induced ATP signaling independently of both intracellular and extracellular Ca2+.

Discussion

In this study, we demonstrated that MS induces ATP release in GBM. Notably, MS-induced ATP release occurred independently of Ca2+ signaling. Using pharmacological screening and genetic knockdown, we found that TRPM3 mediates ATP release upon MS. Our results not only elucidate the mechanism of MS-induced ATP release but also implicate TRPM3 in promoting invasion in GBM. While TRPM3 is generally known as a Ca2+-permeable channel, this study reveals its additional role in mediating MS-inducted ATP release in GBM. Overall, this novel mechanism provides new insights into how mechanical forces regulate GBM invasion via TRPM3-mediated ATP release, highlighting TRPM3 as a potential therapeutic target.

Through pharmacological screening, we identified multiple ion channels involved in MS-induced ATP release. Treatment with 2-APB resulted in the full blockade of MS-induced ATP release. However, since 2-APB targets multiple ion channels, it was not possible to determine which specific channel was responsible for MS-induced ATP release. To address this limitation, we used selective blockers to identify specific channels. Given that 2-APB also inhibits gap junctions, we applied carbenoxolone (CBX), a gap junction-specific blocker, and observed a similar reduction in ATP release as seen with 2-APB treatment. These findings suggest that gap junctions, composed of connexin and pannexin channels, mediate intercellular interactions and play a role in MS-induced ATP release (Anselmi et al. 2008). However, since CBX treatment did not fully block ATP release to the same extent as 2-APB, we hypothesized that additional 2-APB-sensitive channels are involved in MS-induced ATP release. Among the channels inhibited by 2-APB, we investigated TRPA1 using the specific inhibitor HC-030031, followed by ATP imaging. The results demonstrated a decrease in ATP release, indicating that TRPA1 is indirectly involved in MS-induced ATP release (Figure 3A, Blue). Since TRPA1 plays a key role in pain sensation, it is likely to interact with other mechanosensitive channels to detect diverse stimuli (Zhang et al. 2022). However, since TRPA1 inhibition only partially reduced ATP release compared to 2-APB, we hypothesized that additional channels contribute to this mechanism. Furthermore, we did not target TRPC or TRPV channels, as ATP release was still detected upon their inhibition. Given that our results under Ca²+-free conditions indicated a Ca²+-independent process, we excluded IP3 receptors from further investigation (Mikoshiba 2015). Instead, we focused on the TRPM subfamily and decided to generate shRNA constructs to specifically knock down TRPM channels.

After discovering that TRPM3 regulates MS-induced ATP release, we conducted an invasion assay to investigate the effects of TRPM3-knockdown in U-87 MG. The invasion assay showed a reduction in invasion in TRPM3-knockdown U-87 MG cells. Intracellular ATP can be released to act in an autocrine manner, affecting the same cell, or in a paracrine manner, influencing neighboring cells (Corriden and Insel 2010). Therefore, in TRPM3-knockdown U-87 MG cells, the reduction in MS-induced ATP release may have led to a decrease in ATP signaling for both the same cell and neighboring cells. Given that TRPM3 functions as a Ca²+-permeable channel, the observed reduction in GBM cell invasion could be attributed not only to diminished ATP release but also to decreased Ca²+ influx. While Ca2+ influx is known to play a critical role in invasion, it has been reported that in GBM, P2X7R is activated upon binding to extracellular ATP, leading to intracellular Ca²+ influx through ligand-gated cation channels (So et al. 2021a). Therefore, the reduction in MS-induced ATP release may have decreased Ca2+ influx not only through TRPM3 but also through other ATP-sensitive Ca²+ channels, ultimately contributing to decline in invasion. Furthermore, previous studies have shown that tumor-associated microglia and macrophages (TAMs) enhance extracellular ATP production, which in turn promotes glioma growth and invasion in the GBM microenvironment through the activation of P2X7R (Wu et al. 2024). These findings support that ATP-mediated signaling within the GBM plays a pivotal role in regulating tumor invasion. Collectively, our study elucidates the relationship between MS-induced ATP release and TRPM3, proposing that extracellular ATP levels play a crucial role in regulating GBM invasion.

Within the highly complex and heterogeneous TME of GBM, intricate interactions between tumor cells and a diverse cells intricately regulate tumor growth and invasion (Giraldo et al. 2019). The GBM TME consists of GBM cells, glioma stem cells, neurons, glial cells, immune cells, the brain vascular system, and the extracellular matrix layer (Sharma et al. 2023). Previous studies have demonstrated that all cell types within the GBM TME express ATP receptors, and their activation upon ATP binding initiates a purinergic signaling cascade (Illes and Nörenberg 1993; Hidetoshi et al. 2012; Illes et al. 2021; Kan et al. 2023). Notably, we have observed that MS induces ATP release in GBM, leading to a transient increase in extracellular ATP concentration. This ATP release into the TME activates ATP receptors on various TME components, thereby promoting GBM progression. Activation of purinergic receptors has been directly linked to cytoskeletal remodeling and invasive-protrusion formation, as demonstrated by the identification of a P2X7-associated signaling complex that physically interacts with actin-binding proteins and induces membrane blebbing upon ATP stimulation (Kim et al. 2001). Moreover, purinergic signaling broadly regulates focal adhesion turnover and membrane dynamics via PLC/PI3K-dependent signaling downstream of P2Y receptors, thereby facilitating cell traction and invasion (Huang et al. 2021).

Consequently, numerous studies have identified ATP receptors as potential therapeutic targets for GBM (Monif et al. 2014; Di Virgilio et al. 2021). Furthermore, GBM exhibits upregulated P2X7R, and prior research indicates that P2X7R activation, modulated by ATP concentrations within the TME, enhances GBM invasion (Kan et al. 2023; Wu et al. 2024). Furthermore, it is possible that TME-derived ATP promotes GBM progression by activating non-GBM cells (tumor-associated cells). GBM conditioned medium (CM) plays a role in promoting extracellular matrix invasion by microglia (Chang et al. 2025). Furthermore, exogenous ATP promotes macrophage growth (Kawamura et al. 2012), which, as previously reported, may also induce the release of exogenous ATP from tumor-associated macrophages (TAMs), thereby boosting GBM growth (Wu et al. 2024). Further studies are needed to elucidate the effects of MS-induced ATP release on other cells within the TME. Given that our current experimental design may not fully recapitulate the in vivo complexity of the TME, future studies should aim to develop more physiologically relevant models. While P2X7 receptors generally contribute to cancer cell survival, studies suggest that they may fail to form functional pores under conditions of high ATP concentrations within the TME (Gilbert et al. 2019). Considering this characteristic, further research is imperative to delineate how MS-induced ATP release from GBM influences ATP concentration dynamics within the TME.

Author contributions

K.G.B., C.W.S., J.H.Y., and K-S.H. designed the study and analysis. Experiments were performed by K.G.B., C.W.S., J.H.Y., M.L., N.K., G.S.L., and Y.M.P. Data analysis was performed by K.G.B., C.W.S., J.H.Y., and G.S.L. The study was supervised by C.J.L. and K-S.H. Glioblastoma specimens and derived cell lines were kindly provided by co-author I-Y.H. and J.Y., obtained through a protocol approved by the Institutional Review Board of Gangnam Severance Hospital (IRB No. 3-2024-0129). This manuscript was prepared by K.G.B., C.W.S., J.H.Y., D.H.W., C.J.L., and K-S.H. K.G.B. and C.W.S. can be interchanged in the author list because they contributed equally, with the order decided by ladder-climbing.

Supplementary Material

Supplemental Material
TACS_A_2629066_SM7871.tiff (19.3MB, tiff)

Funding Statement

This work was supported by Regional Innovation System & Education (RISE) program through the Daejeon RISE Center, funded by the Ministry of Education (MOE) and the Daejeon Metropolitan City, Republic of Korea: [Grant Number No. 2025-RISE-06-012].

Disclosure statement

No potential conflict of interest was reported by the author(s).

Supplemental Material

Supplemental data for this article can be accessed online at https://doi.org/10.1080/19768354.2026.2629066.

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