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INTRODUCTION
Oral cancer is a pressing global health concern, ranking as the sixth most common cancer type worldwide [1]. Oral squamous cell carcinoma (OSCC) accounts for over 90% of all oral cancer cases and presents a substantial challenge due to its high rates of metastasis and recurrence [2]. Despite relentless efforts to develop oral cancer treatment options, the five-year survival rate for OSCC patients remains below 60%, and this is largely attributed to the limitations of existing treatment modalities [3]. Conventional treatments, such as surgery, radiotherapy, and chemotherapy, often fail to yield the desired outcomes and can impact patients’ aesthetic and functional results [4]. In addition, the significant role that cervical lymph node metastasis plays in the poor prognosis of OSCC patients is of particular importance [5]. Thus, understanding the molecular mechanisms associated with metastasis and developing effective therapeutic strategies that target them are of paramount importance.
Apoptosis is a highly regulated process of programmed cell death that has a pivotal role in human embryonic development and the maintenance of tissue homeostasis in adults [6]. It involves two distinct pathways: the intrinsic pathway, which occurs within cells and is primarily associated with mitochondrial damage, DNA damage, or abnormalities in cellular oxidative processes; and the extrinsic pathway, which is initiated when external signals bind to cell surface receptors. Both pathways include complex mechanisms and lead to nuclear condensation, DNA fragmentation, and apoptosome formation [7,8]. In normal cells, apoptosis and the cell cycle process work harmoniously to ensure tissue equilibrium [9]. However, in cancer cells, genetic mutations and aberrations often disrupt this delicate balance, enabling them to bypass cell cycle checkpoints and proliferate in an uncontrollable manner [10]. Therefore, understanding the intricate interplay between cancer, apoptosis, and the cell cycle is expected to support the development of effective cancer treatment and prevention strategies [11,12].
The biological phenomenon of epithelial-to-mesenchymal transition (EMT), in which epithelial cells acquire mesenchymal characteristics, is a key factor in cancer progression [13]. While it is normal for EMT to occur during embryonic development and organogenesis, its occurrence in tumor cells enhances their motility and significantly contributes to metastasis [14]. Extensive research has established that EMT not only facilitates metastasis but also amplifies the invasive potential of existing cancer cells, which substantially increases the risk of cancer recurrence and diminishes the survival rates associated with various cancer types [15,16].
Tivozanib is a tyrosine kinase inhibitor and promising anticancer agent due to its ability to inhibit the vascular endothelial growth factor receptor (VEGFR), which is critical in the growth of blood vessels that supply nutrients to tumors [17,18]. Tivozanib has primarily been used to treat advanced renal cell carcinoma (RCC), and studies have shown that it effectively impedes tumor blood vessel growth and potentially reduces tumor growth rates [19]. Surprisingly, there is a notable lack of research on its pharmacological effects on OSCC cells; hence, there is a critical need for comprehensive investigations in this area. Consequently, the aim of this study was to evaluate the impact of tivozanib on OSCC cells, with a focus on its capacity to induce apoptosis and cell cycle alterations and influence factors associated with EMT, a crucial process in cancer cell metastasis.
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METHODS
Cell culture
The OSCC cell lines CAL27 and Ca9-22 were acquired from the ATCC and propagated in Dulbecco’s modified Eagle’s medium supplemented with 10% fetal bovine serum and 1% penicillin-streptomycin. The cell cultures were maintained in a 5% CO2 atmosphere at a temperature of 37°C, with routine subculturing performed every two to three days.
MTT assay
To assess cell viability, CAL27 and Ca9-22 cells were transferred to 96-well plates. After a 24 h incubation to allow the cells to adhere, various concentrations of tivozanib (Sigma-Aldrich) were added, and the cells were incubated for a further 24 h. MTT solution (0.5 mg/ml) was then added, and the cells were incubated for 3 h at 37°C. Following the removal of the supernatant, formazan dissolution was achieved with dimethyl sulfoxide (DMSO), and the absorbance at 570 nm was recorded using a SpectraMax iD3 microreader. Three independent experiments were conducted, and the results are presented as mean ± standard deviation (SD).
Cell proliferation assay
CAL27 and Ca9-22 cells were seeded in six-well plates. After incubating for 24 h to allow the cells to stabilize, various tivozanib concentrations (0–10 µM) were added, and the cells were incubated for one week. The cells were then washed with phosphate buffered saline (PBS), fixed in methanol, and stained with 0.5% crystal violet. The stained colonies were dissolved in DMSO. The absorbance was measured at 570 nm with a SpectraMax iD3 microreader; three replicates were used.
Hoechst 33342 and JC-1 staining
For staining with Hoechst 33342, CAL27 and Ca9-22 cells were seeded in Lab-Tek II chamber plates (Nunc; Thermo Fisher Scientific) and treated with tivozanib for 24 h at 37°C in an incubator chamber. The cells were then washed with cold PBS and fixed with 4% paraformaldehyde (PFA) for 10 min at room temperature. Subsequently, the cells were stained with Hoechst 33342 (1 µg/ml in PBS) for 30 min at 37°C in a 5% CO2 incubator chamber. The nuclear morphology was observed using a Lionheart FX automated microscope (BioTek).
For staining with JC-1, CAL27 and Ca9-22 cells were seeded in Lab-Tek II chamber plates and treated with tivozanib for 24 h at 37°C in an incubator chamber. The cells were then washed with cold PBS and stained with JC-1 (1:10,000) for 20 min at room temperature. The cells were observed using a Lionheart FX automated microscope.
Flow cytometry
Cell cycle analysis: Cells were seeded in 60-mm dishes and incubated for 24 h. The cells were then detached using a scraper and washed with PBS containing 1% bovine serum albumin (BSA). After washing, the cells were fixed in 70% ethanol for 24 h. Approximately 10 µM RNase was then added to the cells, and the cells were incubated on ice for 30 min. Afterward, 10 µl of propidium iodide (PI) was added to each sample, and the samples were analyzed using flow cytometry (FACS) with the PI wavelength. Cells in the G1, S, and G2 phases were analyzed, quantified, and graphically represented.
Mitochondrial membrane potential (MMP) assessment: JC-1 staining was used to evaluate the MMP. After treatment with tivozanib, cells were stained with JC-1 according to the manufacturer’s instructions. This dye selectively enters mitochondria and changes color, from green to red, as the membrane potential increases. The stained cells were subjected to flow cytometry analysis. The shift in fluorescence from green (monomers) to red (aggregates) was quantified, and this provided a measure of the change in the MMP in response to tivozanib.
Measurement of the rate of apoptosis: Cells treated with tivozanib were stained with annexin V and PI. Annexin V binds to phosphatidylserine exposed on the outer leaflet of the plasma membrane in early apoptotic cells, while PI stains the DNA of late apoptotic or necrotic cells. Flow cytometry was performed to detect and analyze the live, early apoptotic, and late apoptotic/necrotic cells. This method allowed for the quantification of the apoptosis rates in the cell populations exposed to tivozanib.
Wound healing and invasion assays
Wound healing assay: This assay was performed to assess the ability of cells to migrate in response to tivozanib treatment. Initially, cells were cultured in dishes until they formed a monolayer. Subsequently, a sterile pipette tip was used to create a straight scratch (“wound”) on the cell layer. After this wounding, the cells were washed to remove any detached cells and debris. The medium was then replaced with medium containing 25 µM tivozanib. Images of the wound area were captured after staining with Hoechst and using a Lionheart FX.
Invasion assay: Cells were seeded in Matrigel-coated transwell plates (Corning Costar) and treated with tivozanib for 48 h in serum-free medium. The cells in the upper chamber were washed with PBS and fixed with 100% methanol for 15 min. Hematoxylin and eosin were applied to the upper chamber for 10 min each, and then the cells in the upper chamber were washed with PBS. The membranes from the upper chamber were separated, dehydrated by sequential immersion in 70, 80, 90, and 100% ethanol, and mounted using malinol. Images were captured and analyzed using a Lionheart FX.
Immunofluorescence staining
CAL27 and Ca9-22 cells were seeded in Lab-Tek II chamber plates and incubated for 24 h to allow attachment and stabilization. After the cells were treated with tivozanib, immunofluorescence staining was performed to detect the expression of cytochrome c and E-cadherin (Cell Signaling Technology). The cells were fixed with 4% PFA for 10 min and then permeabilized with 0.1% Triton X-100 solution (in PBS) for another 10 min to enhance antibody penetration. Primary antibodies specific for cytochrome c and E-cadherin were diluted (1:200) in PBS containing 1% BSA. The primary antibodies were applied to the cells, and the cells were incubated for 24 h at 4°C. The cells were then washed three times with PBS, and an FITC-conjugated goat anti-rabbit IgG Fc secondary antibody (Thermo Fisher Scientific) (diluted at 1:200) was then applied for 2 h at room temperature. After three additional washes with PBS, the cytoskeleton was stained with rhodamine phalloidin, a red fluorescent dye, and the nuclei were stained with DAPI (Thermo Fisher Scientific). Each stained sample was observed, photographed, and analyzed using a Zeiss LSM 750 laser scanning confocal microscope.
Western blot analysis
The proteins present in tivozanib-treated cells were extracted and quantified. The cells were lysed at 4°C for 2 h using RIPA buffer, which contained 50 mM Tris-Cl, 300 mM NaCl, 0.5% Triton X-100, 2 µl/ml aprotinin, 2 mM PMSF, and 2 µl/ml leupeptin and had a pH of 7.6. The protein content of the cell lysates was determined using the Bradford assay (Bio-Rad), and each sample was adjusted to contain 20 µg of protein. The samples were loaded onto a 10% SDS-PAGE gel and subjected to electrophoresis. The separated proteins were then transferred to a PVDF membrane (Millipore). The membranes were blocked with 5% nonfat dry milk for 1 h. Primary antibodies (specific for cyclin B1, cyclin E1, Cdk2, Cdc2, p21, caspase 7, caspase 3, PARP, Bcl-2, apoptosis-inducing factor [AIF], E-cadherin, N-cadherin, Slug, Snail, Twist, matrix metalloproteinase 9 [MMP9], vascular endothelial growth factor receptor 2 [VEGFR2], and β-actin; Cell Signaling Technology) were diluted 1:1,000 and applied, and the membranes were placed in a refrigerator overnight. After washing the membranes six times with PBS for 10 min each time, a secondary antibody (diluted 1:5,000) was applied for 1 h at room temperature. The membranes were washed a further six times with PBS for 10 min each time, and the proteins were detected using a Super Signal West Femto-enhanced chemiluminescent substrate. The protein signals were captured using an Image Quant LAS 500 chemiluminescence system (GE Healthcare).
RNA isolation and RT-qPCR
Tivozanib-treated cells were collected using a scraper, and the total RNA (30 ng) was isolated from each sample using the RNeasy mini kit (Qiagen Inc.). The isolated RNA from each sample was then converted into complementary DNA (cDNA) using an M-MLV cDNA synthesis kit (Enzynomics). The genes of interest were labeled using the SYBR Green kit (Applied Biosystems), and gene expression was quantified by performing 40 amplification cycles using the ABI 7500 real-time PCR detection system (Applied Biosystems). Table 1 shows the gene type and sequence details.
Table 1
Sequences of primers
Statistical analysis
Data are presented as mean ± SD, and the figures shown were generated from the results of at least three independent experiments. The statistical analysis was conducted using one-way analysis of variance, followed by Dunnett’s comparison. Differences with a probability value (p-value) of less than 0.05 were considered statistically significant.
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RESULTS
Tivozanib reduced the viability and proliferation of OSCC cells from two different cell lines
To evaluate the impact of tivozanib on the viability and proliferation of OSCC cells, we exposed Ca9-22 and CAL-27 cells to various concentrations of tivozanib (0–200 µM). We employed the MTT assay to assess cell viability, and the results indicated that there was a dose-dependent reduction in cell viability in both cell lines, as shown in Fig. 1A. These results guided our selection of the range of tivozanib concentrations to utilize in subsequent experiments, which was 0–75 µM. We investigated the effect of tivozanib on cell proliferation by subjecting OSCC cells to lower concentrations of the drug. As shown in Fig. 1C, tivozanib exerted a dose-dependent inhibitory effect on cell proliferation. The results of a quantitative analysis of cell colonies stained with crystal violet provided statistical confirmation of these findings (Fig. 1B). Thus, it was found that tivozanib could effectively reduce the viability and proliferative ability of OSCC cells.
 | Fig. 1Tivozanib exhibits cytotoxic and antiproliferative effects and induces cell cycle arrest in OSCC cells.(A) Ca9-22 and CAL27 cells were treated with varying concentrations of tivozanib (0–200 μM) for 24 h. (B, C) Cells were treated with 1 or 10 μM tivozanib for 7 days. Cell proliferation was assessed by staining with 1% crystal violet. Cell viability and proliferation values are presented as percentages (compared to control cells with 0 μM tivozanib). (D) Cells were treated with 0–75 μM tivozanib for 24 h. After staining with PI, the cells were analyzed using FACS. The proportion of cells in the G2 phase after tivozanib treatment is presented as a histogram. (E) The protein levels of cell cycle-related factors were confirmed via Western blot. β-actin was used as a loading control. The results are presented as mean ± SD and were derived from three independent experiments (Ca9-22: *p < 0.05, **p < 0.01, ***p < 0.001; CAL27: #p < 0.05, ##p < 0.01, ###p < 0.001). OSCC, oral squamous cell carcinoma; PI, propidium iodide.
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Tivozanib arrested the cell cycle in OSCC cells from two different cell lines
To evaluate the capacity of tivozanib to alter the cell cycle in OSCC cell lines, we examined the proportions of Ca9-22 and CAL-27 cells that were within specific phases of the cell cycle using FACS. Upon tivozanib treatment, both Ca9-22 and CAL-27 cells displayed an increasing trend in the proportion of cells arrested in the G2 phase, and this was found to be dose-dependent (Fig. 1D).
In accordance with these observations, we conducted a western blot analysis to investigate whether proteins known to be involved in the regulation of the cell cycle were altered. It is well known that the Cdk2/cyclin B1 and Cdc2/cyclin E1 complexes play pivotal roles in the mechanisms that control progression through the G2 phase of the cell cycle; their activation and regulation are key factors [20]. Therefore, we examined whether there were any changes in the levels of these proteins and found that there was a dose-dependent decrease in the levels of the Cdk2/cyclin B1 and Cdc2/cyclin E1 complexes after tivozanib treatment. Additionally, p21 showed a dose-dependent increase in response to tivozanib treatment (Fig. 1E). These findings indicated that tivozanib had an effect on the cell cycle in OSCC cells and provided further support for the notion that tivozanib inhibits the proliferation of cancer cells.
Tivozanib induced apoptosis through the intrinsic pathway in OSCC cells from two different cell lines
The morphological changes induced by tivozanib treatment in OSCC cells were assessed by staining the cell nuclei with Hoechst 33342 and observing them under a fluorescence microscope. Following tivozanib treatment, significant morphological alterations were observed in both Ca9-22 and CAL27 cells. Specifically, there was an increase in the number of condensed nuclei, and strong fluorescence was detected in both condensed and fragmented nuclei in cells from both OSCC cell lines (Fig. 2A). FACS was used to quantify the number of cells that underwent apoptosis after treatment with tivozanib (Fig. 2B), and the results revealed a dose-dependent increase in the rate of apoptosis. In the Ca9-22 cells, the late apoptosis/death rate was 6.57% in the control cells and 32.0% in the cells treated with 75 µM tivozanib; a significant increase. Similarly, in the CAL27 cells, the rate was 7.27% in the control cells and 24.73% in the cells treated with 75 µM tivozanib. We also conducted a western blot analysis to assess the expression of apoptosis-related factors, including caspase 7, caspase 3, and PARP. Following tivozanib treatment, a decrease in caspase 7 expression and cleavage of caspase 3 and PARP were observed in both Ca9-22 and CAL27 cells (Fig. 2C). To determine whether tivozanib induces apoptosis through the intrinsic pathway, we first evaluated the MMP in Ca9-22 and CAL-27 cells treated with 50 µM tivozanib using JC-1 staining and FACS. Treatment with tivozanib resulted in the conversion of JC-1 aggregates (red) to monomers (green) (Fig. 3A), and the FACS analysis showed that there was a significant increase in green fluorescence in both cell types following tivozanib treatment (Fig. 3B; Ca9-22 cells: control cells = 25.67%, tivozanib-treated cells = 93.64%; CAL-27 cells: control cells = 24.57%, tivozanib-treated cells = 79.92%). We then assessed the levels of Bcl-2, AIF, and cytochrome c. In the intrinsic pathway of apoptosis, Bcl-2 is the pivotal protein; when the level of this antiapoptotic factor decreases, there is a reduction in the MMP. Consequently, the pores in the mitochondrial inner membrane open, which enables the release of cytochrome c, and AIF is also released from the mitochondria into the cytoplasm [21]. Treating Ca9-22 and CAL-27 cells with tivozanib clearly decreased the Bcl-2 level and increased the AIF level (Fig. 3D). Furthermore, there was a distinct release of cytochrome c (Fig. 3C). Based on these results, it can be inferred that tivozanib induces apoptosis in OSCC cells through the intrinsic pathway.
 | Fig. 2Apoptotic responses in OSCC cells induced by tivozanib.After treating cells with 0–75 μM tivozanib for 24 h, (A) nuclear condensation and formation were observed. The number of condensed nuclei was quantified and is displayed in the graph. (B) Apoptosis in OSCC cells following tivozanib treatment was quantified using annexin V/PI staining and FACS. (C) The levels of apoptosis-related proteins were analyzed via Western blot. β-actin was used as a loading control. The results are presented as mean ± SD and were derived from three independent experiments (Ca9-22: *p < 0.05, **p < 0.01; CAL27: #p < 0.05, ##p < 0.01). OSCC, oral squamous cell carcinoma; PI, propidium iodide.
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 | Fig. 3Intrinsic pathway-mediated apoptosis in OSCC cells triggered by tivozanib.Cells were treated with 0.75 μM tivozanib for 24 h. After staining with JC-1, the cells were observed using fluorescence microscopy (A) and FACS (B). The green fluorescence signal in the cells treated with tivozanib was compared with that in the control cells. (C) An increase in the release of cytochrome c from the mitochondria suggested the activation of apoptotic pathways. (D) The levels of proteins involved in intrinsic pathway-mediated apoptosis were analyzed via Western blot. β-actin was used as a loading control. OSCC, oral squamous cell carcinoma; AIF, apoptosis-inducing factor.
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Tivozanib inhibited EMT in cells from two different OSCC cell lines
To investigate the impact of tivozanib on the migratory and invasive characteristics of Ca9-22 and CAL-27 cells, we conducted wound healing and invasion assays. The microscopy results unequivocally demonstrated that tivozanib treatment significantly reduced the ability of both Ca9-22 and CAL-27 cells to migrate and invade (Fig. 4A, B). These findings strongly indicated that tivozanib exerted a pronounced inhibitory effect on the motility of the OSCC cells.
 | Fig. 4Tivozanib inhibited EMT in OSCC cells.(A, B) The wound area and number of invaded cells both significantly decreased with tivozanib treatment. (C) Tivozanib regulated the expression of EMT-related proteins in OSCC cells. (D) qPCR analysis of the mRNA levels of EMT-related markers. (E) E-cadherin expression increased with tivozanib treatment, as confirmed by confocal microscopy. The results are expressed as mean ± SD. Data were derived from three independent experiments (Ca9-22: *p < 0.05, **p < 0.01; CAL27: #p < 0.05). EMT, epithelial-to-mesenchymal transition; OSCC, oral squamous cell carcinoma; MMP9, matrix metalloproteinase 9; VEGFR2, vascular endothelial growth factor receptor 2.
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Additionally, we assessed the expression of E-cadherin, a protein involved in maintaining cell–cell adhesion, using confocal microscopy. The results revealed a conspicuous increase in green fluorescence, which signified enhanced E-cadherin expression in the cytoplasm of both Ca9-22 and CAL-27 cells (Fig. 4E). The results of a western blot analysis of EMT-related proteins corroborated these observations, showing an upregulation of E-cadherin. Conversely, the levels of N-cadherin, Snail, Slug, and Twist, which are associated with increased cell motility, were significantly reduced. Furthermore, the level of MMP9, a factor directly involved in cell motility, was markedly reduced, as was the level of VEGFR2, which is involved in cancer cell angiogenesis (Fig. 4C). These findings were supported by the qPCR results, which showed that there was an increase in the E-cadherin mRNA level and decreases in the mRNAs that encode N-cadherin, Slug, and other related proteins (Fig. 4D). These findings suggest that tivozanib has considerable potential as an anticancer agent that suppresses metastasis and invasion in OSCC.
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DISCUSSION
OSCC is the most prevalent form of oral cancer and is closely associated with risk factors such as tobacco and alcohol consumption, as well as human papillomavirus infection [22,23]. OSCC frequently metastasizes to lymph nodes and blood vessels, which increases the risk of the disease spreading to other parts of the body [24]. Unfortunately, the effectiveness of existing anticancer drug treatments against OSCC is limited; thus, the overall survival rates associated with OSCC are low [24,25]. Therefore, it is crucial to research and develop effective anticancer agents that specifically target oral cancer cells. Tivozanib is a VEGFR inhibitor known for its ability to suppress angiogenesis and inhibit metastasis in RCC [26]. However, the potential anticancer effects of tivozanib in oral cancer cells have not been fully elucidated. To address this gap, we have investigated the anticancer activity of tivozanib in OSCC cells.
Previous studies have explored the effects of tivozanib on cell viability and proliferation in ovarian cancer cells and glioblastoma cells. In ovarian cancer cells, treatment with 10 µM tivozanib led to an approximately 50% reduction in cell viability, while in glioblastoma cells, a significant decrease in cell viability was observed after treatment with 20 µM tivozanib. In both cases, treatment with 5 µM tivozanib resulted in at least 50% inhibition of cell proliferation [17,27]. In the current study, we evaluated cell viability by exposing cells to varying concentrations of tivozanib, up to 200 µM. The results revealed that there was a gradual dose-dependent decrease in cell viability that started when the concentration of tivozanib reached 10 µM. Additionally, there was a significant decrease in the cell proliferation rate that began at 2.5 µM tivozanib. Based on these experimental findings, we established that tivozanib effectively reduces cell viability and proliferation rates in OSCC cell lines. We also used flow cytometry to analyze the proportions of OSCC cells that were within specific phases of the cell cycle following tivozanib treatment. In the Ca9-22 cells, the percentage of cells in the G2 phase rose from 18.9% to 67.19% after tivozanib (50 µM), treatment. Similarly, in the CAL-27 cells, the percentage of cells in the G2 phase significantly increased from 28.75% to 67.87% in response to tivozanib treatment. Cell cycle arrest allows cells to undergo necessary preparations and checks before cell division [28], and G2 arrest can occur in response to various cellular signals, such as DNA damage or incomplete replication [29].
One of the most characteristic changes that occurs in cancer cells undergoing apoptosis is the condensation of the nucleus, which leads to DNA fragmentation [30]. Our experimental results showed that the cells from both the OSCC cell lines exhibited typical morphological changes associated with apoptosis after tivozanib treatment. Nuclear condensation was observed in both cell lines, along with a significant increase in the number of fragmented nuclei. It was also found that treatment with tivozanib significantly increased the rate of late apoptosis/death. In the Ca9-22 cell line, the apoptosis rate in the control cells was 6.57%, whereas it increased to 32.0% in the cells treated with 75 µM tivozanib. Similarly, in the CAL-27 cell line, the late apoptosis/death rate increased from 7.27% in the control cells to 24.73% in the tivozanib-treated cells. These results suggest that tivozanib induces morphological changes and enhances apoptosis in OSCC cells.
Bcl-2 is a key regulator of apoptosis, promotes cell survival, and is located in the outer mitochondrial membrane [31]. During apoptosis, Bcl-2 is inhibited, which leads to a decrease in the MMP and the depolarization of the mitochondria [32]. This change can be detected via JC-1 staining—JC-1 aggregates exhibit red fluorescence in healthy polarized mitochondria, whereas JC-1 monomers exhibit green fluorescence in depolarized mitochondria [33]. The release of cytochrome c into the cytosol and the formation of apoptosomes are also considered key apoptotic events; they result in the activation of caspases and lead to cell death [34]. Additionally, AIF, which is normally localized in the mitochondria, is released into the cytosol during apoptosis and translocates into the nucleus where it promotes DNA fragmentation [35]. Our experimental results showed that treatment with tivozanib led to a decrease in Bcl-2 and an increase in green-fluorescent JC-1, indicating a reduction in the MMP in both OSCC cell lines. There was also an increase in the release of cytochrome c in both cell lines. These findings suggest that tivozanib treatment induces cell death in OSCC cells through the intrinsic pathway of apoptosis.
Given that a cancer cell’s capacity to metastasize is heavily reliant on its inherent motility and invasiveness [36], we conducted wound healing and invasion assays to investigate the effects of tivozanib on the motility and invasiveness of the tested cell lines. Our findings indicated that tivozanib exerted an inhibitory effect on the motility of the OSCC cells.
As the main focus of this study, we examined the effect of tivozanib on EMT, a biological process that plays a crucial role in embryonic development, tissue repair, and cancer metastasis [37]. EMT is regulated by various proteins that modulate the transition of epithelial cells to a mesenchymal phenotype [38]. In cancer progression, E-cadherin plays a crucial role in suppressing tumor progression and metastasis [39], and the loss or downregulation of E-cadherin is frequently observed in several types of cancer, including OSCC [40]. The expression of E-cadherin was observed using confocal microscopy, and the findings indicated that treatment with tivozanib promoted E-cadherin expression and maintained the integrity of cell junctions. The increase in E-cadherin expression was confirmed by western blot analysis. In contrast, the expression of N-cadherin, Snail, Slug, and Twist, which are factors that increase cell motility, decreased in a dose-dependent manner as the tivozanib concentration increased.
MMP9 and VEGFR2 are also involved in cancer progression, including OSCC progression [41]. MMP9 has been shown to degrade the extracellular matrix surrounding tumor cells and thus facilitate their invasion into surrounding tissues, as well as to promote angiogenesis to support tumor growth [42]. The receptor for vascular endothelial growth factor (VEGF), VEGFR, when bound to VEGF, activates intracellular signaling pathways controlling angiogenesis, cell growth, and survival. Notably, VEGFR1 and VEGFR2 are well-known; VEGFR2, in particular, plays a key role in angiogenesis and is a significant target in cancer research [43].
Hypoxia-inducible factor-1 alpha (HIF-1α) is activated under hypoxic conditions and increases in various solid tumors, including lung cancer. HIF-1α affects the expression of genes related to cancer progression such as cell division, angiogenesis, energy metabolism, and EMT, thus promoting metastasis in lung cancer. This finding suggests that VEGF is also a crucial gene regulated by HIF-1, potentially offering a new strategy for cancer treatment [44]. In this context, our study observed a significant decrease in VEGFR2 at concentrations over 25 µM. Similarly, research by Momeny et al. [27] indicated that tivozanib effectively inhibits VEGFR at concentrations above 5 µM, with a marked reduction at 20 µM in ovarian cancer cells. These parallels suggest a consistent dose-response relationship in the inhibition of the VEGFR pathway by tivozanib. In our study, we observed a decrease in the expression levels of MMP9 and VEGFR2, suggesting that tivozanib may hold promise in curtailing cancer metastasis and invasion. These findings provide valuable insights into the development of therapeutic strategies for OSCC. Following treatment of OSCC cells with tivozanib, we conducted qPCR analysis to measure the mRNA levels of EMT-related factors. The results demonstrated an upregulation in E-cadherin expression and a downregulation of N-cadherin and slug expressions. These results further support tivozanib's role as a VEGFR inhibitor in regulating EMT-related processes and inhibiting the invasive potential of OSCC cells.
In this study, we evaluated the diverse anticancer effects of tivozanib. Our findings revealed that tivozanib effectively induces cell cycle arrest, which leads to the inhibition of cell proliferation. Furthermore, apoptosis was observed in the treated cells. Therefore, we have demonstrated that tivozanib has potential as an anticancer agent that suppresses the motility and invasiveness of cancer cells and thus the metastasis of OSCC cells. Based on these findings, it can be concluded that tivozanib holds promise as an effective therapeutic agent for the treatment of OSCC.
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