Between January 2016 and January 2017, 10 cases of single oral buccal and facial VM were reported at Department of Oral and Maxillofacial Surgery, The Affiliated Stomatological Hospital of Nanjing Medical University, including 4 male and 6 female patients, with a mean age of 29.4±13.7 years. The diagnosis of buccal facial vein malformation was confirmed by preoperative pathological examination. The inclusion criteria were as follows: Patients who were diagnosed with single oral buccal and facial venous malformation by pathological examination. The exclusion criteria were as follows: Patients who were diagnosed with other types of venous malformations on the face, hypertension, serious cardiovascular and cerebrovascular diseases, and other patients who were not suitable for treatment (for example cardiopulmonary dysfunction or poor recovery). The present study was approved by the Ethics Committee of The Affiliated Stomatological Hospital of Nanjing Medical University (Nanjing, China) and written informed consent was obtained from each subject. Surgically resected VM tissues were separated under aseptic conditions, and the exposed malformed vascular mass was cut open. Sponge-like sinusoids were then visualized and were sliced into tissue blocks of approximately 3–5 mm². Following immersing in culture medium for minimal humidification, tissue blocks were inoculated in a culture flask that was subsequently plated on 1% gelatin, with the intima facing downward. Tissue blocks were separated by a distance of ~5 mm. The tissue blocks were then incubated in Dulbecco's modified Eagle's medium containing 20% fetal bovine serum (Gibco; Thermo Fisher Scientific, Inc., Waltham, MA, USA) at 37°C with the culture flask inverted. Following 6 h, the culture flask was upended for incubation at 37°C. Tissue blocks and non-adherent cells were removed, and adherent cells were incubation in Endothelial Cell medium (cat. no. 1001; ScienCell Research Laboratories, Inc., San Diego, CA, USA) for 3 days at 37°C. Cellular morphology was observed under an inverted microscope at a magnification of ×400, and cells outside the circle were removed using a cytopipette. Subsequently, tissue blocks were washed using PBS twice to remove other cells, and the culture medium was replaced. When cells had reached 70–90% confluency, 0.25% trypsin supplemented with ethylenediaminetetraacetic acid was used for cell subculture.

Flow cytometry analysis was used to detect endovascular epithelial markers in order to identify endovascular epithelial cells. Separated cells from deformed veins in human VM were detached using 2.5 g/l tryptase and centrifuged at 167 × g and 4°C for 5 min. The cells were washed with cold PBS at 4°C and centrifuged at 167 × g and 4°C for 5 min. The washing and centrifugation were repeated and then the cells were made into a single cell suspension. With 1×10⁶ cells per tube, a 15-min incubation at 37°C was performed with the addition of 20 µl phycoerythrin-conjugated anti-human cluster of differentiation (CD)31 antibody (cat. no. 566177; 1:500; BD Biosciences, San Jose, CA, USA), or a 30-min incubation at 4°C with the addition of 20 µl mouse anti-human von Willebrand factor (vWF) monoclonal antibody (cat. no. 555849; 1:500; BD Biosciences). For determination of CD31/vWF, a simultaneous incubation was performed for 15 min at 37°C. Subsequently, a second incubation for 30 min at 4°C with the addition of 50 µl fluorescein isothiocyanate-labeled rabbit anti-mouse IgG (cat. no. TA130002; 1:1,000; OriGene Technologies, Inc., Beijing, China) was performed. Cells were place in tubes on the ice in the dark for 30 min, washed three times using 2 ml PBS and at 167 × g and 4°C and to remove the supernatant. Then 500 µl PBS (containing 4% paraformaldehyde) was used to fix the cells at 4°C for 30 min prior to performing the flow cytometry. Cell identification was conducted using a FACSCan flow cytometer and the data was analyzed by Cell Quest Software (version 5.1; both BD Biosciences, Franklin Lakes, NJ, USA).

The separated and determined endothelial cells of human VM were assigned into a blank group (cells cultured conventionally, without any treatment); a 1 ng/ml group (cells cultured in medium with rapamycin working solution at a concentration of 1 ng/ml); a 10 ng/ml group (cells cultured in medium with rapamycin working solution at a concentration of 10 ng/ml); a 100 ng/ml group (cells cultured in medium with rapamycin working solution at a concentration of 100 ng/ml), and a 1,000 ng/ml group (cells cultured in medium with rapamycin working solution at a concentration of 1,000 ng/ml). Rapamycin was purchased from Hunan TaiRen Pharmaceutical Co., Ltd. (Changsha, China).

Cell viability of the endothelial cells in each group was measured at 24, 48 and 72 h following incubation. The optimal reaction time point for rapamycin was determined for the observation of cell morphological changes under an inverted microscope and other experiments. A cell suspension for each group was inoculated in 96-well plates at a density of 5×10⁴ cells/well following dilution. A total of 6 reduplicate wells were set up for each group. Upon reaching 80% confluency, endothelial cells were treated as described above. Following reoxygenation, 20 µl MTT solution (Sigma Aldrich; Merck KGaA, Darmstadt, Germany) was added and incubated for 4 h at 37°C, followed by the removal of MTT solution. Subsequently, each well was supplemented with 150 µl dimethyl sulfoxide (Sigma Aldrich; Merck KGaA). Following shaking on a shaking table for 10 min, the optical density (OD) value of each well was measured using a microplate reader at a wavelength of 490 nm. The experiment was repeated 3 times, and the mean OD value was calculated. Cell viability was calculated as follows: (OD value_{experimental group}-OD value_{blank group})/OD value_{blank group}.

Slides with attached cells were fixed in 4% (w/v) paraformaldehyde for 30 min at room temperature, rinsed in PBS, blocked with 0.3% H₂O₂ in methanol for 15 min at room temperature, and finally rinsed in PBS. Subsequently, slides were maintained in 0.1% Triton X-100 at 4°C for 20 min. The TUNEL assay (cat. no. C1098; Beyotime Institute of Biotechnology, Haimen, China) was performed according to the manufacturer's instructions. Slides were blocked with 3% bovine serum albumin (cat. no. C0225; Beyotime Institute of Biotechnology) at 37°C for 20 min, followed by 3 washes with PBS (5 min per wash). Subsequently, peroxidase solution was added, and the slides were incubated in a humidity chamber at 37°C for 30 min, followed by PBS washing 3 times (5 min/wash). Diaminobenzidine (0.3 ml) as a substrate was added as a chromogen, and the slides were incubated in a humidity chamber at 15–20°C for 10 min, followed by 3 washes with PBS (5 min per wash). Counterstaining was performed conventionally with hematoxylin at 37°C for 10 sec. Subsequently, ethanol-hydrochloric acid was used for differentiation, and ammonia was used to color the cells blue. The cells were mounted with 4% Paraformaldehyde Fix Solution (cat. no. P0099; Beyotime Institute of Biotechnology) and photographed. Normal endothelial cells were stained blue, and apoptotic cells were yellow and brown. Five fields of vision were randomly selected. The apoptosis rate for a fixed area was calculated (apoptosis rate=apoptotic cells/total cells in a field).

On the back of a 6-well plate, uniform horizontal lines at intervals of approximately 0.8 cm were drawn across the wells using a marker. Each well was crossed by at least 5 lines, and ~5×10⁵ cells were inserted into each well and grown to 100% confluency. The following day, a sterile 10-µl pipette tip, perpendicular to the horizontal lines on the back, was used to scratch wounds along a ruler. Importantly, the pipette tip was upright, rather than slanted. Following wound scratching, the cells were rinsed gently with PBS 3 times. Then, PBS was added along the wall, and the scratched cells were rinsed and removed. The endothelial cells were incubated with culture medium in a 5% CO₂ incubator at 37°C. Samples were taken at 48 h and then photographed. The rate of wound closure was calculated by measuring the distance between the two wound edges at 0 h and at 48 h. The experiment was repeated 3 times, and the mean value was calculated.

SPSS 21.0 software (IBM Corp., Armonk, NY, USA) was used to analyze data in the present study. Measurement data are presented as the mean ± standard deviation. Comparisons among multiple groups were performed using one-way or two-way analysis of variance followed by Tukey's post hoc test. P<0.05 was considered to indicate a statistically significant difference.

The cellular morphology of separated human VM endothelial cells was observed under an inverted microscope. When the separated endothelial cells grew adherent to the wall, they were enlarged to different degrees and grew in clusters, resembling paving stones, with clear, homogeneous cytoplasm and a round or oval nucleus (Fig. 1A). For accurate identification of endovascular epithelial cells separated from the deformed vein of a human VM, flow cytometry was performed to evaluate a more specific marker for endovascular epithelial cells, CD31 and vWF, and it was demonstrated that there were 92.5% CD31-positive cells, 88.2% vWF-positive cells, and 76.3% CD31- and vWF-positive cells (Fig. 1B).

An MTT assay was performed to detect the cell viability of human VM endothelial cells exposed to rapamycin (Fig. 2). Rapamycin significantly inhibited the proliferation of human VM endothelial cells at 48 and 72 h. Higher concentration of rapamycin induced greater inhibitory effects. However, only rapamycin at a concentration of 1,000 ng/ml significantly inhibited the viability of human VM endothelial cells at 24 h, suggesting that the inhibitory effects of rapamycin on human VM endothelial cells were concentration- and time-dependent. Therefore, 48 h following incubation was determined to be the optimal reaction time.

An inverted microscope was used to observe the morphological changes of human VM endothelial cells exposed to rapamycin at different concentrations (Fig. 3). At 48 h, when compared with the blank group, treatment with rapamycin at a concentration of 1 ng/ml produced a few shrunken cells and a reduced cell number; however, the majority of cells exhibited a clear boundary and a regular shape. Treatment with rapamycin at 10 and 100 ng/ml reduced the number of cells even further and caused some cells to become round and desquamated; treatment with rapamycin at 1,000 ng/ml produced more obvious desquamation, round-shaped cells and necrosis.

A TUNEL assay was conducted to determine the apoptosis conditions of human VM endothelial cells exposed to rapamycin at different concentrations (Fig. 4). In comparison with the blank group, apoptosis of human VM endothelial cells was significantly elevated when treated with rapamycin at 1, 10, 100 and 1,000 ng/ml. Higher concentrations yielded greater numbers of apoptotic endothelial cells, indicating that rapamycin treatment promoted the apoptosis of human VM endothelial cells.

A wound-healing assay further verified the inhibitory effects of rapamycin on the migration capacity of human VM endothelial cells (Fig. 5). Compared with the blank group, rapamycin significantly suppressed the migration capacity of human VM endothelial cells at different concentrations. Higher concentrations yielded greater inhibitory effects.