Peripheral blood samples were obtained from patients with DVT (n=5) and healthy subjects (n=5) during physical examination between January and June 2023, with age ranging from 40 to 60 years. The sex distribution in the DVT group is 3 males and 2 females, while in the healthy control group, the sex ratio is 2 females to 3 males. Briefly, a total of 10 ml of peripheral blood was isolated from subjects from both groups, and following centrifugation at 800 g, 4°C for 10 min, the serum was isolated and stored at −80°C. All patients provided written informed consent for their enrollment in the present study, which was approved by the Ethics Committee of the Fengyang County People's Hospital.

Human umbilical vein endothelial cells (HUVECs, cat. no. CTCC-009-493), as immortalized cells, not primary cells, were purchased from the Cell Bank of the Chinese Academy of Sciences and cultured in Endothelial Cell Medium (ECM; cat. no. 1001; ScienCell Research Laboratories, Inc.) supplemented with 10% FBS (cat. no. 10099; Gibco; Thermo Fisher Scientific, Inc.) and 0.5% penicillin/streptomycin solution (cat. no. 15640055; Gibco; Thermo Fisher Scientific, Inc.) at 37°C in an incubator with 5% CO₂. HUVECs were treated with ox-LDL at 37°C for 48 h (40 µg/ml, cat. no. 20605ES10; YEASEN) to establish an in vitro model.

Total RNA was isolated from 5×10⁵ cells or 1.5 ml peripheral blood samples using a TRIzol^® reagent (cat. no. 15596018CN; Invitrogen; Thermo Fisher Scientific, Inc.) according to the protocols. The quality of the isolated RNA was then verified by UV spectrophotometry and formaldehyde denaturing electrophoresis. Subsequently, a total of 1 µg RNA was reverse transcribed into cDNA using AMV reverse transcriptase (cat. no. 10109118001; Roche Diagnostics, Ltd.) according to the manufacturer's protocols. The gene expression levels was amplified by using the SYBR qPCR kit (cat. no. RK21203; ABclonal), and normalized to GAPDH expression, were calculated using the 2^−ΔΔCq method (32). The PCR reaction consisted of pre-denaturation at 94°C for 5 min, 40 cycles of denaturation at 94°C for 30 sec, annealing at 58°C for 30 sec and extension at 72°C for 1 min. PCR was carried out using a real-time amplifier (cat. no. ABI PRISM7700, Thermo Fisher). The assays were performed in triplicate. The primer information was: MDM2: 5′-ATGAAAGCCTGGCTCTGTGT-3 (Forward), 5′-CACTCTCCCCTGCCTGATAC-3′ (Reverse); PTEN: 5′-AGTTCCCTCAGCCGTTACCT-3 (Forward), 5′-AGGTTTCCTCTGGTCCTGGT-3′ (Reverse); and GAPDH: 5′-CAGCCTCAAGATCATCAGCA-3 (Forward), 5′-TGTGGTCATGAGTCCTTCCA-3′ (Reverse).

Prior transfection, cells were seeded into 24-well plates and cultured until they reached ~30% confluence. For short interfering (si)RNA transfection, 0.67 µg (50 pmol) siRNAs (Sangon Biotech Co., Ltd.) were diluted in the appropriate serum-free diluent at a final volume of 25 µl. Subsequently, 1 µl Entranster-R4000 (cat. no. 4000-3; Engreen Biosystem, Co., Ltd.) was mixed with 24 µl serum-free diluent to a final volume of 25 µl. The Entranster-R4000 diluent was left at room temperature for 5 min. Then, the Entranster-R4000 and RNA diluents were thoroughly mixed by oscillating with an oscillator or blowing with a sampler for >10 times. The resulting transfection complex (50 µl) supplemented with 0.45 ml of complete medium was added into each well. The petri dish was gently moved back and forth to ensure even distribution. Cell morphology was observed at 6 h after transfection and when cell viability was unaffected, this indicated that the transfection reagent did not exhibit toxicity to the cells and the medium was not changed. Cells were then cultured for 24–96 h at 37°C to obtain the desired results. The interference sequence were as follows: si-MDM2: SS sequence: GGAACUUGGUAGUAGUCAAUC, AS sequence: UUGACUACUACCAAGUUCCUG; si-PTEN: SS sequence: AGAUGUUAGUGACAAUGAACC, AS sequence: UUCAUUGUCACUAACAUCUGG; si-negative control (NC): SS sequence: UUCUCCGAACGUGUCACGU, AS sequence: ACGUGACACGUUCGGAGAA.

Cells were seeded in a 6-well plate at a density of 700 cells per well in complete medium. The cultures were maintained until the majority of single colonies reach a cell count of >50 cells/colony. The medium was changed every three days. Subsequently, cells were washed once with PBS and fixed with 4% paraformaldehyde (1 ml/well; cat. no. P1110; Beijing Solarbio Science & Technology Co., Ltd.) for 30 min at room temperature, followed by staining with crystal violet solution (1 ml/well cat. no. C0121; Beyotime Institute of Biotechnology) for 10 min at room temperature. Following washing with PBS (cat. no. 10010023; Gibco; Thermo Fisher Scientific, Inc.), cells were allowed to air-dry.

Cells were inoculated into a 6-well plate at a density of 5×10⁵ cells/well. The following day, a scratch was made using a sterile pipette tip (200 µl, cat. no. BS-200-T; Biosharp). The cells were then washed three times with PBS and supplemented with serum-free ECM. Finally, cells were cultured in an incubator at 37°C with 5% CO₂ for 0, 6, 12 and 24 h and observed under an optical light microscope (cat. no. CKX41, Olympus). Observe under 200× magnification and randomly select 3 fields of view for photography and counting.

A total of 1×10⁵ cells were re-suspended in serum-free ECM and were then added onto the upper chamber of the Transwell insert (cat. no. 14141; Beijing Labselect). Medium containing 10% FBS was supplemented into the lower chamber and cells were cultured for 48 h at 37°C. The cells were subsequently fixed with 70% methanol for 10 min at room temperature (cat. no. 322415; MilliporeSigma) and stained with 1% crystal violet for 5 min at room temperature. The invasive cells were counted at five randomly selected fields under an optical light microscope (cat. no. CKX41, Olympus). Observe under 100× magnification and randomly select 3 fields of view for photography and counting.

Cell apoptosis was assessed using an Annexin V-FITC/PI Kit (cat. no. 40302ES20; Shanghai Yeasen Biotechnology Co., Ltd.). Cells were collected, mixed with 5 µl Annexin V for 30 min at 37°C, then incubated with propidium iodide in the dark for 20 min at 37°C. Apoptotic cells were counted using flow cytometry (CytoFLEX LX; Beckman Coulter). Analyze the data using Flowjo software (Version 10.8; flowjo.com/; BD Biosciences). Combine the proportion of early apoptotic cells (Annexin V single positive) and late apoptotic cells were considered apoptotic cell proportion.

Angiogenesis was evaluated using an Angiogenesis Assay Kit (In Vitro; cat. no. ab204726; Abcam). A 96-well culture plate was coated with an extracellular matrix solution and incubated at 37°C for 1 h to allow gel formation. Subsequently, 70,000 cells were seeded onto the gel and cultured for an additional 18 h. After the incubation medium was removed, the cells/gel were washed. The formation of blood vessels was then evaluated under a light microscope (cat. no. CKX41, Olympus). Observe under 200× magnification and randomly select 3 fields of view for photography and counting.

5×10⁵ cells in a petri dish were lysed with the corresponding lysis buffer (cat. no. 20-188; Merck Millipore) with a protease inhibitor (cat. no. P8340; MilliporeSigma) on ice for 30 min. After centrifugation at 2,000 g at 4°C in the maximum rotating speed for 30 min, supernatant was collected. A small portion of the lysate was subjected to western blot analysis, while the remaining lysate was mixed with 2 µg of the corresponding antibody and incubated at 4°C overnight. To prepare the protein A agarose beads (cat. no. P1925; MilliporeSigma), 10 µl of the solution/beads was washed three times with cracking buffer (cat. no. 20-188; Merck Millipore), followed by centrifugation at 1,000 g for 3 min each time at 4°C. A total of 10 µl pre-treated protein A agarose beads was added to the cell lysate, which was incubated with 2 µg MDM2 antibody (cat. no. 33-7100; Thermo Fisher) overnight, and the mixture was gently shaken at 4°C for 2–4 h. The aforementioned procedure allowed the coupling between the antibody and protein A agarose beads. Following immunoprecipitation, the agarose beads were centrifuged at 4°C for 3 min at 1,000 × g. After beads were pelleted, the supernatant was carefully aspirated and the agarose beads were washed with 100 µl Elution buffer (cat. no. P2179S; Beyotime Institute of Biotechnology) for 3–4 times. Lastly, the dilution was then supplemented with 100 µl 2X SDS loading buffer (cat. no. 9172; Takara Biotechnology Co., Ltd.) and the mixture was boiled for 5 min prior western blot analysis.

Proteins were extracted from cells using RIPA lysis buffer (cat. no. P0013B; Beyotime Institute of Biotechnology) and protein concentration was quantified using a BCA kit (cat. no. P0011; Beyotime Institute of Biotechnology). A total of 30 µg total protein extracts were separated by 10% SDS-PAGE and were then transferred to PVDF membranes (cat. no. IPVH00010; MilliporeSigma). Subsequently, the membranes were blocked with 5% skimmed milk (cat. no. P0216; Beyotime Institute of Biotechnology) at 4°C for 2 h and then with primary antibodies at 4°C overnight. Following washing with PBS-1% Tween-20 (PBST; cat. no. BL314B; Biosharp Life Sciences) for three times, the membrane was incubated for 2 h with the corresponding secondary antibody. After washing with PBST for three times, the protein bands were visualized using ECL Chemiluminescent buffer (cat. no. 36208ES76; YEASEN) in a dark room. The grayscale values were calculated using ImageJ software (Version 1.8.0; National Institutes of Health). The following primary antibodies were used: Anti-MDM2 (cat. no. ab16895), anti-PTEN (cat. no. ab32199), anti-phosphorylated (p)-JAK1 (cat. no. ab138005), anti-JAK1 (cat. no. ab133666), anti-p-STAT3 (cat. no. ab76315), anti GAPDH (cat. no. ab8245) and anti-STAT (cat. no. ab68153; all from Abcam). In addition, the HRP anti-rabbit IgG antibody (cat. no. ab288151; Abcam) served as a secondary antibody.

Search Tool for the Retrieval of Interacting Genes/Proteins (STRING) database (string-db.org/) was used to verify the association between MDM2 and PTEN.

All statistical analyses were performed using GraphPad prism 6.0 (Dotmatics). The experiments were independently repeated at least three times. The results are expressed as the mean ± standard error of the mean. The differences between two groups were compared using unpaired Student's t-test, while those among multiple groups by ANOVA followed by Tukey's post hoc test. P<0.05 was considered to indicate a statistically significant difference.

To investigate the involvement of MDM2 and PTEN in DVT, RT-qPCR analysis was performed to detect their expression levels in venous blood samples obtained from patients with DVT. The participants were allocated into two groups, namely the control and DVT groups. The results revealed that MDM2 and PTEN were markedly upregulated in the intravenous blood samples of patients with DVT compared with the control group (Fig. 1A and B). These results strongly supported the potential significance of MDM2 and PTEN in the pathogenesis of DVT.

To investigate the role of MDM2 in the development of DVT, an in vitro model was established. The results demonstrated that HUVECs treatment with ox-LDL significantly upregulated MDM2 (Fig. 2A). To further elucidate the functional significance of MDM2 in different cellular behaviors, the siRNA-mediated knockdown approach was employed in HUVECs (Fig. 2B). Notably, MDM2 knockdown not only promoted the proliferation of ox-LDL-treated HUVECs (Fig. 2C), but also markedly enhanced their migration and invasion abilities (Fig. 2D and E). The aforementioned findings strongly suggested that MDM2 could play a crucial role in facilitating cell metastasis and invasion during DVT progression. In addition, the results showed that MDM2 silencing suppressed the apoptosis of ox-LDL-treated HUVECs. More particularly, cell apoptosis was significantly reduced in the si-MDM2 group compared with the control group (Fig. 2F). This finding indicated that MDM2 could modulate the delicate balance between cell survival and death throughout DVT development. Furthermore, MDM2 silencing also significantly augmented the angiogenic capacity of ox-LDL-treated HUVECs. Therefore, tube formation assays demonstrated that cells in the si-MDM2 group displayed a more pronounced vascular lumen structure (Fig. 2G). Collectively, these compelling findings underscored the critical involvement of MDM2 in DVT pathogenesis and highlighted its potential therapeutic value in this context.

A previous study revealed that MDM2 interacts with PTEN (33). In the present study, bioinformatics analysis verified the association between MDM2 and PTEN (Fig. 3A). To further validate this finding, a co-immunoprecipitation assay was performed, which verified the binding capacity between MDM2 and PTEN (Fig. 3B). Subsequently, to determine the expression levels of PTEN in different treatment groups, RT-qPCR was employed. Therefore, the results demonstrated that PTEN was notably upregulated in ox-LDL-treated HUVECs compared with the control group (Fig. 3C). To further investigate the effect of MDM2 knockdown on the mRNA and protein expression levels of PTEN, RT-qPCR and western blot analyses were performed, respectively. The results demonstrated a significant reduction in the mRNA expression levels of PTEN in the si-MDM2 group (Fig. 3D). Consistently, western blot analysis further verified that the protein expression levels of PTEN were markedly decreased following MDM2 silencing (Fig. 3E). In summary, the aforementioned findings provided compelling evidence supporting the interaction between MDM2 and PTEN and highlighted the effect of MDM2 knockdown on the mRNA and protein expression levels of PTEN.

Subsequently, the current study aimed to investigate the role of PTEN in DVT. Therefore, PTEN was knocked down in HUVECs (Fig. 4A). Clone formation assay revealed that PTEN silencing promoted the proliferation of ox-LDL-treated HUVECs (Fig. 4B). Furthermore, the wound healing and Transwell assays further substantiated the promoting effect of PTEN knockdown on HUVECs migration and invasion (Fig. 4C and D). More particularly, the migration and invasion abilities of HUVECs were markedly enhanced in the PTEN silencing group compared with the control group. Additionally, flow cytometry demonstrated that PTEN silencing suppressed the apoptosis of ox-LDL-induced HUVECs (Fig. 4E). Lastly, tube formation assays showed that the angiogenic capability of ox-LDL-treated HUVECs was increased in PTEN-depleted cells (Fig. 4F). Overall, the aforementioned results supported the pleiotropic effects of PTEN silencing on the behavior of HUVECs, including cell proliferation, migration, invasion, apoptosis and angiogenesis. These findings, combined with the effect of MDM2 silencing on PTEN expression, further supported the strong association between PTEN and MDM2.

Based on previous studies, Kyoto Encyclopedia of Genes and Genomes (KEGG) analysis was performed to identify the signaling pathways associated with PTEN expression (34). The analysis showed that PTEN could regulate several crucial signaling pathways, including the JAK/STAT signaling pathway (Fig. 5A). This finding was further supported by STRING analysis, which further revealed the interactions between PTEN and proteins involved in the JAK/STAT signaling pathway (Fig. 5B). Furthermore, to elucidate the functional involvement of PTEN in the JAK/STAT pathway, western blot analysis was conducted to detect the expression levels of key proteins (p-JAK1, JAK1, p-STAT3, STAT3) in different treatment groups. The results unveiled that HUVECs treatment with ox-LDL significantly attenuated the phosphorylation levels of JAK1 and STAT (Fig. 5C), thus suppressing the activity of JAK/STAT signaling. Intriguingly, the JAK/STAT signaling pathway was activated following PTEN silencing (Fig. 5C). To further validate the aforementioned finding, cells were also treated with AG-490, a specific inhibitor of the JAK/STAT pathway. As expected, HUVECs treatment with AG-490 effectively inhibited the activation of JAK/STAT signaling (Fig. 5C). Collectively, these findings suggested that PTEN could serve a significant role in DVT via modulating the JAK/STAT signaling pathway.

The aforementioned findings indicated that PTEN could inhibit the JAK/STAT signaling pathway, thus affecting HUVECs proliferation, migration, invasion, apoptosis and angiogenesis. The results obtained from the clone formation assays revealed that PTEN silencing could promote proliferation of ox-LDL-treated HUVECs. However, the stimulatory effect of PTEN silencing was reversed following cell treatment with AG-490 (Fig. 6A). Furthermore, the results demonstrated that PTEN silencing enhanced the migratory and invasive capabilities of HUVECs, as evidenced by the wound healing and Transwell assays, respectively. Conversely, these effects were notably diminished in AG-490-treated HUVECs (Fig. 6B and C). Additionally, flow cytometry analysis further elucidated that PTEN knockdown reduced the apoptosis of ox-LDL-treated HUVECs, which was also reversed by AG-490 (Fig. 6D). In addition, the tube formation assays revealed that PTEN silencing promoted the vasculogenesis of HUVECs, which was counteracted by the presence of AG-490 (Fig. 6E). These findings further supported the role of PTEN in modulating the JAK/STAT signaling pathway and its subsequent effects on the behavior of HUVECs.