, Hyo-Min Park1 , Jae-Young Park1 , Uijin Kim2, Ha Youn Shin2 , Hee Pom Lee3, Jin Tae Hong3 & Do-Young Yoon1,* 
This is an open-access article distributed under the terms of the Creative Commons Attribution Non-Commercial License (http://creativecommons.org/licenses/by-nc/4.0) which permits unrestricted non-commercial use, distribution, and reproduction in any medium, provided the original work is properly cited.Cancer is a major public health concerns and ranks as the second leading cause of death in the United States (1). During tumor growth, angiogenesis promotes the sprouting of new blood vessels from pre-existing ones (2), which is an important process in cancer development. Tumor tissues rely on establishing capillaries to provide the nutrients and oxygen needed for their proliferation and metastasis (3). Angiogenesis consists of complex sequential steps such as degradation of the extracellular matrix around the vessels, activation of endothelial cells, and proliferation in forming of capillaries (4).
A major factor causing tumor angiogenesis is the release of vascular endothelial growth factor (VEGF) A by cancer cells. VEGFA binds to its receptor, VEGFR2, in endothelial cells, initiating downstream signaling pathways and regulating endothelial cells proliferation, invasion, migration, and angiogenesis (5). The inhibition of the VEGF/VEGFR2 pathway can reduce angiogenesis, suppressing cancer progression. For example, VEGFR2 inhibitors, such as sorafenib, sunitinib, and regorafenib, are effective anti-angiogenesis and anticancer therapies (6).
In a previous study, we identified that (E)-2-methoxy-4-(3-(4-methoxyphenyl) prop-1-en-1-yl) phenol (MMPP) possesses diverse properties, such as anti-inflammatory, anticancer, and anti-diabetic effects (7-9). However, the effects of MMPP in endothelial cells have not yet been elucidated. MMPP is a PPARγ agonist in adipocytes (7). PPARγ agonists have anticancer effects by inhibiting VEGFR2 (10). Based on our hypothesis, we propose that MMPP affects angiogenesis by targeting VEGFR2. MMPP may inhibit VEGFR2, suppressing cell migration, invasion, tube formation, and angiogenesis. In this study, we demonstrate the regulatory role of MMPP in endothelial cells.
We investigated the cell viability of MMPP-treated HUVECs, and observed that MMPP exhibited low cytotoxicity at a concentration of 20 μg/ml. However, MMPP significantly reduced cell viability at a final concentration of 40 μg/ml (Supplementary Fig. 1). All experiments were conducted at a 10 μg/ml non-cytotoxic concentration. In the wound healing assay, MMPP treatment resulted in a wider scratched area in response to VEGFA. These results indicated that MMPP influenced the migratory ability of HUVECs (Fig. 1A). MMPP decreased the invasive capacity of HUVECs in the invasion assay (Fig. 1B). MMPP inhibited the VEGFA-induced migration and invasion of HUVECs. In the tube formation assay, MMPP considerably disrupted the tube formation induced by VEGFA (Fig. 1C). The formation of new vessels was confirmed using an ex vivo aortic ring assay. MMPP inhibited the sprouting of new vessels from the aortic ring, regardless of the presence of VEGFA (Fig. 1D). Quantitative analysis showed that MMPP significantly suppressed VEGFA-induced branching of new vessels. Our results suggest that MMPP inhibits angiogenesis in migration, invasion, tube formation, and aortic ring assay.
Molecular docking studies were performed to investigate MMPP binding to VEGFR2. MMPP exhibited a high binding affinity of −8.2 kcal/mol to VEGFR2 ABD. MMPP formed hydrogen bonds with Cys919 and Arg1051, with bond lengths of 3.19 Å and 2.91 Å (Fig. 2A). The hydrophobic region of MMPP formed strong interactions with the residues Leu1035, Leu840, and Phe1047. These results implied that the interaction between MMPP and VEGFR2 was potentially stable, leading to the potent inhibition of the enzymatic function of VEGFR2. Furthermore, MMPP improved the thermal stability of VEGFR2 in HUVECs (Fig. 2B), indicating that MMPP interacts with VEGFR2. Fig. 2C showed that MMPP significantly suppressed VEGFR2 kinase activity. These results demonstrate that MMPP could be an effective VEGFR2 inhibitor in HUVECs.
To elucidate whether MMPP influenced the VEGFR2 pathway, we performed immunoblotting using antibodies against p-VEGFR2, p-AKT, and p-ERK. As shown in Fig. 2D, MMPP reduced the phosphorylation of VEGFR2, AKT, and ERK. Furthermore, we examined the transcription factor NF-κB. The level of NF-κB (p65) in the nuclear fraction was significantly reduced by MMPP (Fig. 2E), indicating the inhibition of NF-κB nuclear translocation. This was evaluated using an immunofluorescence assay. These results demonstrated that MMPP attenuated NF-κB translocation (Fig. 2F). Thus, MMPP inhibited angiogenesis via the VEGFR2/AKT/ERK/NF-κB pathway. To investigate the downstream genes of the VEGFR2 signaling pathway that MMPP inhibited, we focused on the transcription of NF-κB target genes. MMPP significantly suppressed the mRNA expressions of NF-κB target genes including VEGFA, VEGFR2, MMP2, and MMP9 (Fig. 3).
To determine the potential roles of MMPP in the interaction between breast cancer cells and endothelial cells, breast cancer cells-conditioned medium (BCCM) was used to assess its effect on angiogenesis in HUVECs. In breast cancer MDA-MB-231 cells, MMPP suppressed the mRNA expressions and secretion of VEGFA (Supplementary Fig. 2). We hypothesized that conditioned medium from MMPP-treated breast cancer cells (MMPP + BCCM) would induce anti-angiogenic properties in HUVECs. This effect was attributed to decreased expression of VEGFA and the inhibitory ability of MMPP. Compared to the control-conditioned medium (CCM), the BCCM promoted the migratory and angiogenic abilities of endothelial cells (Fig. 4A, B). However, MMPP + BCCM inhibited migration and tube formation in HUVECs, and the formation of new vessels in the mouse aortic ring (Fig. 4C). This suggests that breast cancer cell-derived factors can induce angiogenesis, and that MMPP inhibits the interaction between breast cancer cells and endothelial cells.
Angiogenesis is a critical process in tumorigenesis and metastasis, and the inhibition of angiogenesis has emerged as a therapeutic strategy for cancer. VEGF antibodies and VEGFR2 inhibitors have been investigated as promising anti-angiogenic agents (11). Our study evaluated the modulatory effects of MMPP on anti-angiogenic properties, such as inhibition of migration, invasion, tube formation, and sprouting of new vessels. These effects were mediated via the VEGFR2/AKT/ERK/NF-κB pathway (Fig. 4D).
VEGFR2 is a well-known regulator for angiogenesis (5). VEGFR2 inhibitors are classified into type I (DFG in, e.g., sunitinib) and type II (DFG out, e.g., sorafenib) based on their binding sites on the receptor. The active sites of VEGFR2 includes the hydrophobic region (HYD)-I, HYD-II, and the linker (12, 13). Type I inhibitors competitively inhibit ATP, binding to HYD-I (14). Our findings suggest that MMPP functions similarly to type I inhibitors by binding to the HYD-I region (Leu840, Phe918, and Gly922), and forming hydrogen bonds with Cys919.
Upon binding to VEGFR2, VEGFR2 initiates its kinase activity, leading to receptor autophosphorylation (15). Phosphorylation of Tyr1175 triggers signal transduction to the MAPK and PI3K/AKT pathways, stimulating the survival and migration of endothelial cells (16). We confirmed that MMPP suppressed VEGFR2 kinase activity in HUVECs. Furthermore, MMPP downregulated the phosphorylation levels of VEGFR2, AKT, and ERK. Consequently, our results showed that MMPP effectively inhibited migration, invasion, and angiogenesis by suppressing the VEGFR2/AKT/ERK pathway.
NF-κB regulates various biological processes including inflammation, oncogenesis, and angiogenesis (17). In VEGFR2 signaling, AKT and ERK activate NF-κB transcriptional activity (18). Once activated, NF-κB induces nuclear translocation and transcription of its target genes. Inhibition of the VEGFR2 signaling pathway downregulates NF-κB target genes such as VEGFA, VEGFR2, MMP2, and MMP9 (19-21). These factors are associated with angiogenesis. VEGFA can activate the VEGFR2 pathway through paracrine, autocrine, and intracrine functions, promoting angiogenesis (22). Upregulation of VEGFR2 leads to increased signaling and subsequent angiogenic processes (20). The MMP family is a key component of cell migration and angiogenesis (23). Our findings suggest that MMPP targets the VEGFR2/AKT/ERK/NF-κB signaling pathways, which is involved in the angiogenesis-related genes including VEGFA, VEGFR2, MMP2, and MMP9.
Additionally, we examined angiogenic effects by collecting aortic rings from mice to account for the influence of tissues surrounding the endothelial cells. Furthermore, we evaluated the impact of BCCM on endothelial cells. These approaches were adopted to overcome in vitro assays and provide a deeper understanding of the potential anticancer properties of MMPP. This study provides an understanding of the roles of VEGFR2 in endothelial cells, which is crucial for developing effective therapeutic strategies.
Our findings provide the novel evidence that MMPP effectively suppresses angiogenesis including cell migration, invasion, and tube formation in HUVECs. Notably, MMPP binds to VEGFR2 with a high affinity, leading to the inhibition of VEGFR2 enzymatic function in HUVECs. Finally, MMPP downregulate the mRNA expression of VEGFA, VEGFR2, MMP2, and MMP9 via the VEGFR2/AKT/ERK/NF-κB pathways. Thus, MMPP is a promising VEGFR2 inhibitor for treating various solid cancers, including breast cancer.
MMPP was synthesized and provided by Dr. Jin Tae Hong (Chungbuk National University, Cheongju, Republic of Korea) (24).
Human umbilical vein endothelial cells (HUVECs) were obtained from the American Type Culture Collection (ATCC, Manassas, VA, USA). HUVECs were cultured in Dulbecco’s Modified Eagle Medium (DMEM) supplemented with 10% (v/v) thermally inactivated fetal bovine serum (Hyclone Laboratories, Logan, UT, USA). The cells were incubated at 37°C and 5% CO2.
A molecular docking study of MMPP with the VEGFR2 ATP-binding domain (ABD) was performed using AutoDock VINA v1.2.0 (25). The crystal structure of the VEGFR ABD (PDB code: 4asd) was used in the docking experiments (26). The best binding model was visualized using Biovia Discovery Studio Visualizer v21.1.0 (27).
In vitro, VEGFR2 tyrosine kinase activity was assessed using a cell-based enzyme-linked immunoassay (ELISA) method (28). Cells were seeded in 96-well plates and starved in a serum-free medium for 24 h. Cells were treated with 30 min in a serum-free medium. After fixing with 10% formalin, cells were incubated with an anti-pVEGFR (Tyr 1175) antibody (1:1,000) for 1 h. The cells were washed twice with PBST and incubated with horseradish peroxidase (HRP)-condugated anti-rabbit mouse igG (1:5,000) for 1 h. After washing, the cells were treated with TMB substrate (Thermo Fisher Scientific, Waltham, MA, USA). The reaction was stopped by adding 2 N H2SO4 and the absorbance was measured at 450 nm.
The cells were starved in a serum-free medium for 24 h, and then treated with MMPP and/or VEGFA for 24 h. Total RNA was isolated using an easy-BLUE Total RNA Extraction Kit (iNtRON, Seoul, Korea). First-strand cDNA was synthesized with M-MulV reverse transcriptase (New England Biolabs, Ipswich, MA, USA). The synthesized cDNA was used for real-time polymerase chain reaction (RT-PCR) amplification of specific genes. The primers sequences used are listed in Supplementary Table 1.
Harvested cells were lysed in RIPA buffer to obtain whole protein extract. The cells were fractionated for nuclear and cytoplasmic fractionation using NE-PERTM Nuclear and Cytoplasmic Extraction Reagents (Thermo Fisher Scientific Inc., Waltham, MA, USA). Equal amounts of quantified proteins were loaded onto an appropriate percentage of an SDS-PAGE gel and transferred onto a polyvinylidene fluoride (PVDF) blotting membrane (Cytiva, Malborough, MA, USA). The membranes were blocked with 5% skim milk for 1 h. Specific primary antibodies (1:1,000) were incubated for 1 h. The primary antibodies were used against p-VEGFR2 (#3770s), VEGFR2 (#2472s), and p-ERK (#9101s) (Cell Signaling Technology, MA, United States), p65 (#4764s), p-AKT1/2/3 (#sc-7985), AKT (#sc-1619), ERK (#sc-94), and GAPDH (#sc-47724) (Santa Cruz Biotechnology, CA, United States). Membranes were probed with HRP-conjugated anti-rabbit or anti-mouse IgG antibodies. Visualization was performed using an ECL reagents (Advansta, San Jose, CA, USA). Membranes were stripped using Easter-Blot Stripping Buffer (BioMax, Seoul, South Korea).
The cells were seeded into 24-well plates, and allowed to grow until confluent. A 200 μl tip was used to scratch the wound area. The cells were then treated with MMPP in serum-free medium for 24 h, and images were captured using an inverted phase-contrast microscope (magnification, ×4). The wound areas were calculated and normalized to those at 0 h using Image J (29).
The upper chamber was coated with 0.1% gelatin and 7% basement membrane extract (R&D Systems, Minneapolis, MN, USA). Cells were seeded in the upper chamber in a serum-free medium. In contrast, the lower chamber was filled with a medium containing 10% FBS (Sysmex, Hyogo, Japan). MMPP was added to the upper chamber. The invading cells were stained using Diff-Quick (Sysmex, Hyogo, Japan). Images were obtained using a microscope (magnification at ×4).
To analyze the capillary formation ability of HUVECs, a tube formation assay was performed as previously described (30). BME was coated into 96-well plates, and HUVECs were seeded with MMPP and/or VEGFA in a serum-free medium. Tubular networks were captured and analyzed using the Angiogenesis Analyzer tool in ImageJ (31).
The aortic ring assay has been used as a physiologically relevant ex-vivo model of angiogenesis (32). Eight-week-old female C/57BL/6 mice were purchased from Orient Bio (Seongnam, South Korea). The aortic arch was dissected from 10–12-week-old C57BL/6 mice. The arches were then cut, and embedded in BME. The aortic rings were incubated in an opti-MEM medium supplemented with 2.5% FBS and treated with MMPP or VEGFA. After six days, images were captured using a microscope (magnification at ×4). The total tube length was analyzed using the Angiogenesis Analyzer tool in Image J (31). Animal experiments were approved by the Institutional Animal Care and Use Committee (IACUC) of Konkuk University (Seoul, Korea).
The cells fixed and permeabilized with paraformaldehyde (4%) and methanol, respectively. For blocking, BSA in PBS (1%) was added for 1 h. The cells were incubated with p65 primary antibodies (#4764s, 1:200 dilution) for 1 h, followed by incubation with goat anti-rabbit IgG secondary antibodies labeled with FITC (#AP124F, 1:400 dilution) for 1 h. Cells were labeled with DAPI (Sigma, Missouri, USA, 1:1,000 dilution). A confocal fluorescence microscope (EVOSTM M7000 Imaging System, Thermo Fisher Scientific Inc., Waltham, MA, USA) was used to acquire fluorescence images.
CETSA is used to identify interactions between a ligand and a target protein in cells, because ligand binding can improve the thermal stability of protein (33). HUVECs were treated with MMPP for 2 h, and harvested using trypsin. The cells were heated for 3 min at sequentially increased temperatures using a thermal cycler (40, 45, 50, 55, and 60°C). The heated cells were freeze-thawed tree times with liquid nitrogen, and the supernatants were assessed by immunoblot analyses.
To prepare breast cancer cells-conditioned medium (BCCM), MDA-MB-231 cells were cultivated to 70% confluence in DMEM medium. The cells were starved in a serum-free medium for 48 h. To obtain MMPP-stimulated BCCM (MMPP + BCCM), MDA-MB-231 were starved with serum-free medium for 24 h. In serum-free medium, cells were treated with MMPP (5 and 10 μg/ml) for 24 h. Control-conditioned medium (CCM) was made by incubating DMEM without serum for 24 h. All of those were collected and stored at −20°C.
Statistical analysis was conducted using one-way ANOVA with Tukey’s honest significant difference test. Differences were considered significant at P < 0.05. Results were obtained from three independent experiments and are expressed as the mean ± standard deviation (SD).
The authors have no conflicting interests.
Fig. 1. Effects on migration, invasion, and tube formation in HUVECs and new vessel sprouting in mouse aortic ring. (A) A wound healing assay was conducted to identify migratory ability. The area of the scratch was captured at 0 h (top) and 24 h (bottom) in every group. Quantification of wound area normalized to the scramble control. (B) Transwell invasion assay. The cells penetrating the BME-coated membrane were fixed, and stained, and images were captured using a microscope. (C) Tube formation assay. The results of tube formation were capture by microscopy. Quantification of capillary network formation for 16 h using Image J. (D) Ex-vivo mouse aortic ring assay. Aortic ring was incubated in BME gel with MMPP and/or VEGFA for 6 days. Quantification of branching length using Image J. Data are represented as the mean ± SD (n = 3). *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001 by one-way ANOVA.
Fig. 2. Docking studies with MMPP and VEGFR2 and effects of VEGFR2 downstream signaling. (A) 3D diagram showing the docking complex of MMPP (CID: 122517441) with the VEGFR2 ABD (PBD code: 4asd). (B) CETSA analyzed the thermal stabilization of VEGFR2 protein at different temperatures (40, 45, 50, 55, and 60°C). HUVECs were starved in serum-free medium for 24 h, and then treated with MMPP (10 μg/ml) for 2 h. (C) Cell-based ELISA assessed the VEGFR2 inhibitory effect to measure VEGFR2 kinase activity. HUVECs were treated with MMPP for 30 min, and phosphorylation levels of VEGFR2 (Tyr 1175) were measured at 450 nm. (D) Immunoblotting analyses of phosphorylation levels of VEGFR2, AKT, and ERK. HUVECs were starved in serum-free medium for 24 h. MMPP (10 μg/ml) was treated for 1 h, subsequently VEGFA (40 ng/ml) for 30 min. Bar graphs were represented the band intensities of phosphorylated form normalized with normal form. (E) p65 was examined using immunoblotting of the nucleic and cytosolic fraction. p65 was normalized with PARP in nucleic fraction. MMPP (10 μg/ml) was treated for 1 h, subsequently VEGFA (40 ng/ml) for 1 h. (F) Translocation of p65 was observed using immunofluorescence staining. MMPP (10 μg/ml) was treated for 1 h, subsequently VEGFA (40 ng/ml) for 1 h. Data are represented as the mean ± SD (n = 3). *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001 by one-way ANOVA.
Fig. 3. Effects on mRNA levels of angiogenesis-related factor. mRNA levels of (A) VEGFA, (B) VEGFR2, (C) MMP2, and (D) MMP9 were determined using RT-PCR. HUVECs were starved in serum-free medium for 24 h. The cells were treated with MMPP (10 μg/ml) for 1 h, subsequently VEGFA (40 ng/ml) for 24 h. Data are represented as the mean ± SD (n = 3). *P < 0.05, **P < 0.01, ***P < 0.001 by one-way ANOVA.
Fig. 4. Effects on tube formation, migration, and aortic ring in BCCM. (A) Wound healing assay was performed on HUVECs treated with CCM, BCCM and MMPP + BCCM to analyze their migratory ability. (B) The images of HUVECs tubes formation in CCM, BCCM and MMPP + BCCM were captured, and then the number of junctions was quantified by Image J. (C) The microvessels sprouts of mouse aortic rings were captured in the treatment of CCM, BCCM and MMPP + BCCM. Data are represented as the mean ± SD (n = 3). *P < 0.05, **P < 0.01, ***P < 0.001 by one-way ANOVA. (D) Schematic diagram of anti-angiogenic effects of MMPP in endothelial cells. The figure was created using the Motfolio PPT Drawing Toolkits (motfolio.com).
Author ORCID Information
Na-Yeon Kim Hyo-Min Park Jae-Young Park Ha Youn Shin Jin Tae Hong Do-Young Yoon |
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