Medium- and small-diameter artificial blood vessels composed of core-shell structured nanofibers for preventing intimal hyperplasia

Abstract

The present invention relates to a method for manufacturing an implant material comprising core-shell structured nanofibers for preventing intimal hyperplasia. Nanofibers manufactured by a method for manufacturing core-shell structured nanofibers for preventing intimal hyperplasia and an implant material composition comprising same according to the present invention have the advantages of reducing thrombus formation, which can occur in artificial blood vessels upon initial implantation, and inducing rapid endothelialization. In addition, the implant material according to the present invention can be used for small-diameter peripheral or arterial blood vessel reconstruction and transplantation.

Description

Small- and medium-diameter artificial blood vessels composed of core-shell nanofibers for preventing endometrial hyperplasia

[0001] The present invention relates to a small- to medium-diameter artificial blood vessel composed of core-shell structured nanofibers for preventing endothelial hyperplasia, and more particularly, to a method for manufacturing a small-diameter vascular graft comprising core-shell structured nanofibers for rapid endothelialization and prevention of endothelial hyperplasia.

[0002] Autologous grafts are widely used in many vascular surgeries, such as coronary artery bypass grafting, congenital heart disease, and pulmonary artery disease. However, their application is limited by the structure of the graft, which is not suitable for the vascular site requiring grafting, or the physiological immunoreactivity of the graft material. The development of vascular grafts with antithrombotic and endothelialization properties would be a promising graft for the treatment of vascular diseases. Large-diameter (ID > 6 mm) vascular grafts are already commercially available and in clinical use. However, small-diameter (ID < 4 mm) artificial vascular grafts have been developed, but these grafts suffer from stenosis in the early stage of transplantation and intimal overgrowth later in the transplantation process.

[0003] Unlike conventional electrospinning, coaxial electrospinning simultaneously sprays various polymer solutions or powders from two or more centrally positioned syringes or nozzles to create a coaxial jet. A high voltage is applied to the outer nozzle, inducing electrostatic interactions between the charged polymer solutions to form micro-nanofibers. The inner nozzle contains a core material, which forms core-shell micro-nanofibers during rotation. Coaxial electrospinning offers several advantages over conventional electrospinning and is recognized as a technique for producing core-shell micro-nanofibers. It offers the opportunity to create nanofibers with unique multifunctional properties by loading different materials into the core and shell regions. For example, drugs or bioactive substances can be loaded into the shell region for consistent release. Simultaneously, the core can be designed to provide mechanical properties or enhance biocompatibility. Most research on artificial blood vessels using this process involves loading bioactive substances into the fiber core. However, in principle, this is not suitable for implanting artificial blood vessels.

[0004] According to coaxial electrospinning, both the core and shell of the fiber must degrade to release bioactive substances from the core. Two potential problems arise when applying this system. First, if the shell degrades rapidly, the core will also degrade rapidly, allowing rapid release of bioactive substances. However, this can expose the artificial vessel, making it impossible to implant for a long period of time. Second, if the shell is made of a material that either continuously degrades or does not degrade, bioactive substances cannot be released. The release of bioactive substances from the artificial vessel is essential for initial thrombosis and rapid endothelialization. Therefore, this problem can be solved by simply using a material that continuously degrades when manufacturing the core of the artificial vessel, while loading the shell with a material that enhances or reduces degradability by loading the bioactive substances. This system will allow the shell to continuously degrade while the core maintains its graft form, thereby extending the implantation period.

[0005] PCL has been extensively studied in tissue engineering and is currently an FDA-approved biocompatible material. PCL is used to develop artificial vascular grafts because it slowly degrades over time, enabling tissue remodeling and regeneration. Gelatin is derived from collagen, a naturally occurring protein. Gelatin has a lower potential to induce immune responses or inflammation than synthetic materials. Furthermore, gelatin supports cell adhesion, proliferation, and migration, which are crucial for tissue regeneration and healing. However, if artificial blood vessels are fabricated using only PCL due to its hydrophobic properties, endothelialization within the artificial vessel is difficult. Previous studies have shown that a conventionally electrospun PCL-gelatin inner layer can induce endothelialization within 3 months in a rat aortic model, even without bioactive substances.

[0006] Rapid endothelialization in artificial blood vessels can prevent thrombosis (the development of blood clots) and revascularization (the narrowing of blood vessels). Vascular endothelial growth factor (VEGF) accelerates endothelialization in several ways. First, it stimulates the growth of endothelial cells. Second, VEGF enhances the migration of endothelial cells to the wound site, facilitating rapid regeneration. Third, VEGF induces the formation of new blood vessels from existing blood vessels, facilitating rapid blood flow to injured tissue. Heparin is a natural substance that is highly effective in reducing thrombosis, particularly in relation to the blood clotting process. Its primary mechanism of action is its interaction with anticoagulant III (ATIII), a natural anticoagulant protein in blood.

[0007] Based on the above-described properties, the present inventors developed a core-shell fibrous structured artificial vascular graft. The core was made of PCL, and the shell was made of PCL and gelatin containing VEGF and heparin. The shell, which has excellent degradability, rapidly released VEGF and heparin. In contrast, the PCL core was designed to delay degradability for long-term implantation. The artificial vascular graft was developed by specifying the fiber morphology, hydrophilicity, and bioactive substance release capacity. The elasticity of the developed artificial vascular graft was confirmed through tensile strength evaluation, and stress distribution and blood flow velocity were simulated using COMSOL 5.5 software. The blood compatibility and biocompatibility of the artificial vascular graft were evaluated. Finally, the vascular graft was implanted in a rat aorta model for 2 and 4 months to confirm endothelialization of the artificial vessel and smooth blood flow during the long-term implantation period. Blood flow and velocity within the graft were measured using Doppler ultrasound.

[0008] An object of the present invention is to provide a method for preparing core-shell structured nanofibers for preventing endometrial hyperplasia, comprising the steps of: preparing a core structure of nanofibers by dissolving polycaprolactone (PCL) in dichloromethane; preparing a shell structure of nanofibers by dissolving polycaprolactone (PCL) and gelatin in 2,2,2-trifluoroethanol (TFE); preparing solutions of each of the core structure and the shell structure of the nanofibers using a magnetic stirrer; adding VEGF to the polycaprolactone (PCL) and gelatin solutions; stirring the solutions of each of the core structure and the shell structure of the nanofibers on a magnetic stirrer; and adding heparin to the polycaprolactone (PCL) and gelatin solutions.

[0009] Another object of the present invention is to provide nanofibers manufactured according to the above manufacturing method.

[0010] Another object of the present invention is to provide a graft composition comprising the nanofibers.

[0011] Other objects and advantages of the present invention will become more apparent from the detailed description, claims and drawings below.

[0012] The present inventors successfully developed a small-diameter artificial vascular graft composed of core-shell micro-nanofibers. The shell, composed of PCL and gelatin, facilitated the controlled release of VEGF and heparin, while the PCL core helped maintain mechanical strength. Tensile strength and coagulation tests confirmed that the graft sufficiently withstood blood pressure. Blood flow simulations also confirmed hemodynamic suitability as a vascular graft. The graft exhibited blood compatibility and biocompatibility, confirming its applicability as an in vivo graft. Animal experiments confirmed complete endothelial cell regeneration within two months after implantation, and demonstrated 30% higher blood permeability compared to the control group at four months. No intimal overgrowth was detected, confirming normal blood flow within the artificial vessel. The graft according to the present invention has the advantage of being applicable to small-diameter peripheral or arterial vascular reconstruction and transplantation.

[0013] Problems such as low mechanical strength, thrombosis, intimal hyperplasia, and difficulty in endothelialization of small-diameter artificial blood vessels used in clinical practice have not yet been resolved. Therefore, the present invention aims to: design and manufacture a core-shell fiber small-diameter vascular graft using coaxiality to enhance mechanical properties, and improve biological properties through an electrospinning process. The graft material is composed of polycaprolactone / gelatin to facilitate vascular grafting. The shell is made of heparin, VEGF, and polycaprolactone micro-nanofibers (core).

[0014] In the present invention, it was hypothesized that this structure would allow the fiber's shell structure to release heparin-VEGF, while the core would enhance long-term mechanical strength and decomposition ability. To verify this hypothesis, physico-mechanical property evaluations, cell compatibility and animal testing, and blood compatibility evaluations were performed.

[0015] In animal transplantation, the artificial blood vessel was implanted into a rat abdominal aorta model for a certain period of time. The graft, with its micro-nano-fiber structure, demonstrated an excellent microenvironment conducive to cell attachment and proliferation. Notably, the graft, harvested four months after animal transplantation, demonstrated smooth muscle cell regeneration, blood flow exceeding 120%, and endothelialization within the graft. Based on these findings, the core-shell graft developed in this invention is expected to play a pivotal role in the treatment of vascular diseases.

[0016] Accordingly, in the present invention, an artificial blood vessel developed with a core-shell structure is manufactured with a core made of a material that is consistently decomposed at a constant rate, and a bioactive material is loaded into the shell portion to induce a reduction in thrombosis that may occur in an initially transplanted artificial blood vessel and rapid endothelialization formation.

[0017]

[0018] Hereinafter, the present invention will be described in more detail.

[0019]

[0020] The present invention provides a method for preparing core-shell structured nanofibers for preventing endometrial hyperplasia, comprising the steps of: preparing a core structure of a nanofiber by dissolving polycaprolactone (PCL) in dichloromethane; preparing a shell structure of a nanofiber by dissolving polycaprolactone (PCL) and gelatin in 2,2,2-trifluoroethanol (TFE); preparing solutions of each of the core structure and the shell structure of the nanofiber using a magnetic stirrer; adding VEGF to the polycaprolactone (PCL) and gelatin solutions; stirring the solutions of each of the core structure and the shell structure of the nanofiber on a magnetic stirrer; and adding heparin to the polycaprolactone (PCL) and gelatin solutions.

[0021] In the present invention, in the stirring step, the mixing ratio of the core structure and the shell structure may be 70:30.

[0022] In the present invention, the shell structure including polycaprolactone (PCL) and gelatin may induce constant release of VEGF and heparin.

[0023] In the present invention, the polycaprolactone (PCL) core may maintain mechanical strength.

[0024] In the present invention, the core-shell structure nanofiber may reduce thrombosis.

[0025] In the present invention, the core-shell structure nanofibers may induce endothelialization formation.

[0026] The present invention provides a nanofiber manufactured according to the above manufacturing method.

[0027] The present invention provides a graft composition comprising the above nanofibers.

[0028] Meanwhile, the corresponding features in the above details can be replaced in the above-described part, so their description is omitted.

[0029] The nanofibers manufactured using the method for manufacturing core-shell structured nanofibers for preventing endothelial hyperplasia of the present invention and the graft composition comprising the same have the advantage of reducing thrombosis that may occur in artificial blood vessels following initial transplantation and inducing rapid endothelialization. In addition, the graft material according to the present invention can be utilized for reconstruction and transplantation of small-diameter peripheral or arterial blood vessels.

[0030] Figure 1 shows SEM surface images. (A) PCL-Gel (PG), (B) PCL-Gel-Hep (PGH), (C) PCL-Gel-Hep-VEGF (PGHV). (D) Core-shell structure of PGHV fibers composed of PCL core and PCL shell. (E), (F), and (G) showing fiber diameter distribution of PG, PGH, and PGHV. (H) Small-diameter vascular graft (ID < 2 mm) with core-shell structure. (I) SEM image of graft containing fibers, showing a smoother structure than the inner diameter (yellow arrow).

[0031] Figure 2 shows (A) a cross-sectional SEM image of a tubular (2 mm diameter) PGHV graft, (B) a high-magnification image of cross-sectional fibers, (C) the luminal surface of the PGHV graft, (D) the luminal surface of the PGHV graft, (E) porosity analysis, (F) heparin release profile (loaded into cores), and (G) VEGF release profile (loaded into cores). Statistical significance was calculated by one-way ANOVA with Tukey's test. *p < 0.05, **p < 0.01, ***p < 0.001. "NS" indicates not significant.

[0032] Figure 3 shows the characterization of PGHV grafts. (A) FTIR analysis, (B) contact angle measurement, (C) heparin, (D) VEGF, and PGHV release profiles, (E) degradation evaluation of vascular grafts, and SEM images of PGHV core-shell fiber grafts after (F) 5, (G) 15, and (H) 25 days of degradation evaluation. Statistical significance was calculated by two-way ANOVA with Tukey's test. *p < 0.05, **p < 0.01, ***p < 0.001. "NS" indicates not significant.

[0033] Figure 4 illustrates the mechanical properties of vascular grafts. (A) Schematic diagram of various tensile tests, (B) stress-strain curves, (C) tensile strength, (D) elongation properties, (E) Young's modulus, and (F) suture retention properties of the vascular graft. (G) Simulation of blood velocity within the graft using COMSOL 5.0 software via blood flow, and (H) Simulation of the stress range within the graft using COMSOL 5.0 software via blood flow. Statistical significance was calculated by two-way ANOVA with Tukey's test. *p < 0.05, **p < 0.01, ***p < 0.001. "NS" indicates not significant.

[0034] Figure 5: Schematic representation of (A) hemolysis rate, (B) coagulation index, (C) platelet counts adhered to the grafts, (D) PT, (E) APTT, (F) activated and inactivated platelets. SEM images of platelet adhesion on (G) PG, (H) PGH, and (I) PGHV grafts. (J) Schematic representation of CPAE cell seeding procedure on fibrous scaffolds, (K) absorbance MTT assay values, (L) cell viability, (M) confocal images of CPAE cell proliferation for 7 days, (N) live (green) and dead (red) cell images of the grafts, (O) image of the percentage of cells in the area where F-actin was observed by confocal microscopy, and (P) evaluation of cell counts by live / dead assay in CPAE cells cultured for 5 days. Statistical significance was calculated by one-way ANOVA using the Tukey test (A, B, C, D, E, L, O) and two-way ANOVA using the Tukey test (K). *p < 0.05, **p < 0.01, ***p < 0.001. "ns" indicates not significant. *p < 0.05, **p < 0.01, ***p < 0.001. "NS" indicates not significant.

[0035] Figure 6 shows (A) SEM images of CPAE cells attached to the graft surface, (B) CPAE cell migration analysis, and (C) wound opening rate after 24 hours calculated from the cell migration analysis.

[0036] Figure 7. Images of (A) PG and (B) PGHV grafts implanted in the rat aorta. (C) Doppler ultrasound analysis of rats implanted for 2 and 4 months, (D) schematic of grafts analyzed by Doppler analysis. (E) Peak velocity and (F) mean velocity of the vascular graft calculated from Doppler analysis. (G) Graphic of laminar flow within the PGHV graft and (H) schematic of the PG graft interior due to stenosis. Histological analysis of the extracted grafts. (I) H&E, (J) Masson's trichrome, (K) Von Kossa, (L) Verhoeff-Van Gieson, (M) Safranin O, (N) Thickness of regenerated tissue, (O) Collagen coverage, (P) Elastin coverage. Statistical significance was calculated by two-way ANOVA with Tukey's test. *p < 0.05, **p < 0.01, ***p < 0.001. "NS" indicates not significant.

[0037] Figure 8 shows (A) SEM images of the lumen wall (distal, middle, and proximal) of the graft, (B) schematic representation of endothelialization after analysis by SEM, (C) evaluation of neointimal regeneration formed throughout the graft using H&E staining, and (D) expression of the endothelial marker CD31 (red) (yellow) throughout the entire cross-section of the vascular graft. Dotted circles indicate neointimal areas. Graphical representation of (i) intimal proliferation and (ii) endothelialization within the graft.

[0038] Figure 9 shows A) a highly vascularized adventitia image of the PGHV graft after 2 months and B) an SEM image of the luminal wall (endothelium) of a rat aorta.

[0039] Figure 10. Expression of endothelial markers (A) CD31 (red) and (B) ICAM1 (red) (nuclei = blue) in vascular grafts 2 and 4 months after transplantation. (C) Endothelial marker vWF (red) (nuclei = blue) in the lumen of vascular grafts 2 and 4 months after transplantation. (D) Total cell counts based on nuclei. (E) Relative fraction area of ​​CD31, (F) ICAM1, and (G) vWF expressed within the grafts. Statistical significance was calculated by two-way ANOVA. Tukey's test. *p < 0.05, **p < 0.01, ***p < 0.001. "NS" indicates not significant.

[0040] Figure 11. Expression of smooth muscle cell markers (A) α-SMA (green) and (B) MYH 11 (green) (nuclei = blue) in blood vessels 2 and 4 months after transplantation. (C) Expression of M1 (CD68) marker (CD68+ cells = green, nuclei = blue) and (D) M2 (CD206) marker (CD206+ cells = green, nuclei = blue) in PG and PGHV grafts 2 and 4 months after transplantation. (E) Total number of α-SMA+ cells calculated by counting nuclei within the green area. (F) Fractional areas of α-SMA, (G) MYH 11 are expressed within the grafts. (H) Total amount of MYH 11+ cells calculated. Graphs of (I) relative area fraction of CD68+ cells and (J) relative area ratio of CD206+ cells 2 and 4 months after transplantation. Statistical significance was calculated by two-way ANOVA with Tukey's test. *p < 0.05, **p < 0.01, ***p < 0.001. "NS" indicates not significant.

[0041] Figure 12 is a schematic diagram of (A) the release mechanism of VEGF and heparin in a core-shell structured vascular graft and (B) endothelialization inside the vascular graft.

[0042] Hereinafter, to aid understanding of the present invention, examples will be given in detail. However, the following examples are intended only to illustrate the scope of the present invention and are not intended to limit its scope. These examples are provided to more fully explain the present invention to those of average skill in the art.

[0043]

[0044] 1. Experimental method

[0045]

[0046] 1.1 Fabrication of vascular graft material

[0047]

[0048] 10% w / v PCL dissolved in dichloromethane was used as the core structure of the fiber. For the shell structure, 10% PCL and 10% gelatin were separately dissolved in 2,2,2-trifluoroethanol (TFE). The mixture was then mixed at a ratio of 70:30 using a magnetic stirrer for 24 h. VEGF dissolved in BSA was added to the PCL-gel solution. The final concentration of VEGF was approximately 40 ng / ml. To increase solubility, a drop of Span-80 was added to the solution and incubated on a magnetic stirrer at 4°C for 24 h. Heparin dissolved in PBS was then added to the solution. The final concentration of heparin was approximately 250 IU / ml. Coaxial nozzles (inner: 27G, outer: 25G) were attached to two separate 12 mL (Luer lock) syringes. The distance between the collector and the nozzle tip was 10 cm. The mandrel rotation speed was adjusted to 500 rpm. The core solution was sprayed at a rate of 1.0 ml / hr, and the shell solution was sprayed at a rate of 2.0 ml / hr, and the applied voltage was adjusted to 20 kV. To compare the release patterns of VEGF and heparin, a different type of graft was fabricated, in which VEGF and heparin were loaded into the core of the fiber instead of the shell. Other formulation parameters were the same.

[0049]

[0050] 1.2 Surface morphology and fiber distribution

[0051]

[0052] A small piece of surface (1X1cm) 2 ) and tubular grafts were cut into cross-sections and mounted on sample containers using double-sided carbon tape. Finally, the samples were sputter-coated with platinum. To maintain the fiber structure of the cross-section, the samples were first frozen in liquid nitrogen before sectioning. The fiber structure was observed using a scanning electron microscope (SEM, JSM-6701F, JEOL, Japan) at 10 kV accelerating voltage. Typically, 100 fibers were randomly selected from various parts of the image, and the average fiber diameter was measured using Image J software. To confirm the core-shell fiber structure, VEGF was mixed with FITC-labeled BSA (Albumin-fluorescein isothiocyanate conjugate, Sigma-Aldrich, USA) and dissolved together with the shell solution. The electrospun fibers were then observed using a confocal fluorescence microscope (Olympus, FV10i-W, Japan).

[0053]

[0054] 1.3 ATR-FTIR analysis

[0055]

[0056] To distinguish the presence of various substances, a Thermo Scientific Nicolet iS10 FTIR spectrometer and OMNIC 7.3 Spectra software were used with an attenuated total reflectance (ATR) sampling approach for spectral interpretation. Approximately 8 cm -1 The functional groups of other substances present in the sample at a resolution of 600–4000 cm -1 It was identified in the wavelength range.

[0057]

[0058] 1.4 Water contact angle measurement

[0059]

[0060] To evaluate the hydrophilicity of various vascular graft materials, a wettability quantification device (drop shape analyzer, DSA 100, KRUSS GmbH, Hamburg, Germany) was used to measure the contact angle using the sessile drop method. Briefly, a 5 μL drop was poured onto a support firmly attached to a flat surface. This event was videotaped using a charge-coupled device (CCD) camera. The recordings were then used to evaluate the droplet shape. The average value was determined after considering three samples from a single group.

[0061]

[0062] 1.5 reduction

[0063]

[0064] Three specimens (1X1cm) for each group of vascular grafts 2 ) were placed in separate Petri dishes and carefully weighed to evaluate the degradation process. Afterwards, 2 mL of phosphate-buffered saline (PBS; pH 7.4) was added to each plate. The samples were placed on a reciprocating shaker (SH30L) at 37°C. Freshly prepared PBS solution was added daily. The PBS was removed, and the samples were thoroughly washed after 1, 3, 7, 14, 21, 28, and 42 days. After freeze-drying, the weight change of the samples was recorded.

[0065]

[0066] 1.6 Heparin and VEGF release

[0067]

[0068] To quantify the release of heparin and VEGF from vascular grafts, three samples (1X1cm) from each group were collected. 2) were placed in separate petri dishes containing 2 mL of PBS. The samples were then placed on a shaker (60 rpm) at 37°C for 1, 5, 10, 15, 20, and 25 days. After each time point, the PBS was collected and stored at -20°C. Freshly prepared PBS was then added after each time point. To quantify the heparin release, the toluidine blue (TBO) assay, a well-established method for heparin quantification, was used (Cuenca, JP, et al., 2022; Smith, P., A. Mallia, and G. Hermanson, 1980). Briefly, TBO powder was dissolved in 0.01 M HCl / 0.2 wt% NaCl aqueous solution to produce 2 mL of 0.04 wt% TBO solution. The TBO solution was mixed with the collected sample and gently shaken while incubating for 4 hours to form a TBO-heparin complex. The sample was centrifuged at 3,500 rpm for 10 minutes and gently washed twice with an aqueous solution containing 0.01 M HCl and 0.2 wt% NaCl. The precipitate was dissolved in 5 ml of a mixture of 80% ethanol and 0.1 M NaOH (4 / 1 v / v). 150 μl of the supernatant was transferred to a 96-well plate, and the absorbance was measured at 530 nm using a microplate reader. VEGF was quantified using the VEGF Quantikine Kit (R&D Systems, USA) according to the manufacturer's protocol.

[0069]

[0070] 1.7 Determination of tensile strength

[0071]

[0072] A universal testing machine (UTM, R&B UNITECH-T, Korea) was used to measure the tensile strength of various manufactured graft materials. The specimens were 15X4mm in the longitudinal and radial directions, respectively. 2The samples were cut to size and soaked in PBS for 30 minutes before testing. Afterwards, both sides of the samples were fixed to the aluminum clamps of the machine and pulled at a constant speed of 1 cm / min until the sample failed. Stress was calculated by dividing the load value by the cross-sectional area, and strain was calculated by dividing the displacement value by the length. The slope of the stress-strain curve was used to derive the Young's modulus values ​​of various graft materials. The suture retention strength values ​​of various graft materials were obtained by passing a suture loop (5-0 Prolene®) 2 mm below the top of the graft and pulling the graft at a constant speed of 5 mm / min until failure. The maximum strength was considered the suture retention strength value.

[0073]

[0074] 1.8 Simulation of blood flow within the lumen

[0075]

[0076] Blood flow across the lumen was simulated using COMSOL 5.5 software using a fluid-solid coupling approach. Initially, a basic geometric model of the vascular graft was constructed. Then, the simulation modules for solid mechanics and laminar flow were selected. The parameters were an inner diameter of 2 mm, a length of 6 mm, a wall thickness of 0.2 mm, and a material density of 1255 kg / m. 3 , Young's modulus was fixed at 20 MPa. The material density was calculated by dividing the mass of the graft by its volume. The Young's modulus value was obtained through a tensile strength test. Since the Young's modulus value is around 20 to 30 MPa, the minimum value was selected. The fluid flow was performed with a fluid density of 1060 kg / m 3 It was constructed as a single-phase laminar flow model based on typical human hemodynamic parameters, with a dynamic viscosity of 0.005 Pa / s and inlet and outlet pressures of 126.0 and 125.5 mmHg, respectively.

[0077]

[0078] 1.9 Sterilization of graft materials

[0079]

[0080] The grafts were sterilized by soaking them in 70% ethanol for 30 minutes. They were then exposed to UV light for 90 minutes. After sterilization, the remaining ethanol was thoroughly washed with PBS. The grafts were then soaked in cell culture medium before cell seeding to enhance biocompatibility.

[0081]

[0082] 1.10 Hemolysis Test

[0083]

[0084] The hemolysis rate was used to measure the proportional amount of hemoglobin released from red blood cells into the test sample during the solution phase. Initially, all grafts were incubated in PBS for 48 hours, and the PBS extracts were used for this experiment. Blood was collected from Sprague-Dawley rats and placed in test tubes containing the anticoagulant acid citrate dextrose (ACD) for storage. The blood was then centrifuged at 3000 rpm for 10 minutes. After centrifugation of the whole blood sample, the cells separated from the plasma were washed twice with PBS and centrifuged again to separate the red blood cell pellet. Subsequently, 10 mL of PBS extracts from various samples were mixed with 0.2 mL of red blood cell solution (2% in PBS), and the mixture was centrifuged at 3000 rpm for 10 minutes. The absorbance at 540 nm was measured using a microplate reader (Infinite F50, Tecan, Austria). The following equation was used to calculate the hemolysis rate (HR):

[0085]

[0086]

[0087] Here, AS represents the absorbance of the sample, and AP and AN represent the absorbance of the positive control and negative control, respectively.

[0088]

[0089] 1.11 Blood clotting index

[0090]

[0091] Typically, 100 μL of fresh rabbit blood containing a 10% 0.1 M calcium chloride solution was added to a centrifuge tube containing 5 mg of accurately weighed graft material. Each tube was incubated for 30 minutes and then 3 mL of purified water was added. Then, 200 μL of the supernatant from each sample was collected in a 96-well plate, and the absorbance was measured at 540 nm. The percentage of blood coagulation index was calculated using the following equation:

[0092]

[0093]

[0094] Here, AC represents the absorbance of the control (100 μL of fresh rabbit blood containing 3 mL of water and 10% 0.1 M calcium chloride solution), and AS represents the absorbance of the sample.

[0095]

[0096] 1.12 PT and APTT tests

[0097]

[0098] Initially, 500 μL of human plasma was filled into a tapered tube containing the electrospun graft. A control tube containing only human plasma served as a tapered tube. Each tapered tube was incubated at 37°C for 1 hour. The graft was then removed from the tube, and the activated partial thromboplastin time (APTT) and prothrombin time (PT) of the remaining plasma were measured using an automated coagulation analyzer (SYSMEX CA-5100). Data were calculated by averaging the measurements of three samples.

[0099]

[0100] 1.13 Platelet adhesion

[0101]

[0102] The surface of the vascular graft was treated with platelet-rich plasma (PRP), and SEM images of platelet activation were taken. Anticoagulated whole blood was centrifuged at 2,000 rpm for 10 minutes to separate the PRP, which was then gently removed. The separated PRP (200 μL) was applied to a sterile sample and incubated at 37°C for 1 hour. The sample was washed, treated with 3% glutaraldehyde, dehydrated with a high-concentration ethanol solution (50-100% concentration), and dried for 1 day before SEM analysis.

[0103]

[0104] 1.14 Cell culture

[0105]

[0106] The biocompatibility of fibrous vascular grafts was investigated by seeding bovine pulmonary endothelial cells (CPAE) onto the grafts. CPAE cells were cultured in Eagle's minimal essential medium (EMEM) containing 20% ​​fetal bovine serum (FBS) and 1% penicillin-streptomycin (PS). Cells were cultured in a conductive humidified incubator supplemented with 5% CO2 at 37°C. Fresh medium was added to the culture flasks three times a week.

[0107]

[0108] 1.15 MTT analysis

[0109]

[0110] After 24 hours of culture, remove the medium from each well and add approximately 1X10 4300 μL of CPAE cell suspension containing cells was inoculated. The plates were incubated at 37°C with 5% CO2 for 1, 3, and 7 days. After each incubation period, 30 μL of a 5 mg / mL 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT) solution was added to each well and incubated at 37°C for 4 h. After 4 h, the solution was removed, and 400 μL of dimethyl sulfoxide (DMSO) was added to each well and incubated for another 1 h to dissolve the formazan crystals. Finally, the absorbance was measured at 595 nm.

[0111]

[0112] 1.16 Life / Death Analysis

[0113]

[0114] Cell viability was measured using a Live / Dead viability kit (Invitrogen, US) according to the manufacturer's instructions. The first step was to measure 1X10 5 CPAE cells were seeded and cultured for 1 and 5 days. Each sample was incubated with 200 μL of assay solution and left at room temperature for 45 minutes. The assay solution contained both calcein-AM (green fluorescence), which stains live cells, and ethidium homodimer-1 (EthD-1) (red fluorescence), which marks dead cells. After the incubation period, the stained cells were rinsed with PBS. A confocal fluorescence microscope (Olympus, FV10i-W, Japan) was used for photography. The number of live and dead cells was counted using Image J software.

[0115]

[0116] 1.17 In vivo transplantation

[0117]

[0118] Twelve healthy male Sprague-Dawley rats (weighing approximately 250 g) were used for aortic transplantation. The Animal Ethics Committee of Soonchunhyang University approved the experiment (SCH22-0058). The rats were sedated with isoflurane. First, a midline laparotomy incision was made to isolate the rat abdominal aorta. The tubular graft (5-7 mm in length, 2 mm in internal diameter) was then sutured end-to-end with 8-10 ruptured threads using 9-0 monofilament nylon suture (ETHILON™, ETHICON, USA). The wound was then closed with 4-0 monofilament nylon suture (SURGIFIT, Ailee Company Ltd, South Korea). Anticoagulation was avoided postoperatively. The graft was removed at predetermined intervals (2 and 4 months). Before extraction, the patency of the graft was assessed using Doppler ultrasound (Liu, J., et al., 2022). After 2 and 4 months, the mice were sacrificed, and the grafts were extracted. The extracted grafts were washed with PBS to remove excess blood. The grafts were then embedded in optimal cutting temperature (OCT) compound and frozen at -80°C. The grafts were then sectioned into 10-μm-thick slices using a cryotome. For SEM observation, the grafts were cut longitudinally into two pieces. The lumenal wall was then fixed with 2.5% glutaraldehyde and dehydrated using a sequence of ethanol solutions. The samples were dried overnight under a fume hood and sputter-coated with platinum. The proximal, middle, and distal lumenal walls of the grafts were observed using a scanning electron microscope (SEM, JSM-6701F, JEOL, Japan) at 10 kV accelerating voltage.

[0119]

[0120] 1.18 Histological analysis

[0121]

[0122] First, histological staining with Hematoxylin and Eosin (H&E), Masson's trichrome, Von Kossa, Verhoeff-Van Gieson (VVG), and safranin-O was performed on frozen sections of the extracted grafts.

[0123] For immunofluorescence staining, frozen slices were washed up to three times with PBS. Cryosection samples were then permeabilized with 0.5% Triton X-100. After thorough rinsing, the samples were blocked with 10% goat serum for 50 minutes at room temperature. Subsequently, the sections were incubated overnight at 4°C with primary antibodies such as anti-CD31 (Abcam, USA, 1:200), anti-ICAM1 (Abcam, USA, 1:100), anti-vWF (Abcam, USA, 1:100), anti-α-SMA (Abcam, USA, 1:100), anti-MYH11 (Santacruz, USA, 1:100), anti-CD68 (Bio-Rad, USA, 1:100), and anti-CD206 (Santacruz, USA, 1:200) to distinguish endothelial cells, smooth muscle cells, inflammatory cells, and macrophages (M2 type), respectively. The sections were then washed four times with PBS and incubated with appropriate secondary antibodies (1:200) for 2 h at room temperature. Hoechst was used to stain the nuclei. Visualization and imaging were performed using a fluorescence microscope (Nikon Eclipse Ti2, Japan). The captured images were quantified using Image-J software. Neo-tissue thickness, collagen coverage, elastin coverage, and cell number (mm 2 ), while calculating the relative fraction area ratio of MYH11 and the relative fraction area ratio of αSMA, different parts (middle part and anastomotic part) were considered for quantification. Three images of the middle part of the graft were considered to calculate the relative area fraction (%) of vWF, CD31, and ICAM1.

[0124]

[0125] 1.19. Statistical Analysis

[0126]

[0127] Statistical analyses were performed using GraphPad Prism Software v5.0 (San Diego, CA, USA). One-way analysis of variance (Tukey's test) was used for multiple comparisons of one variable, and two-way analysis of variance (Tukey's test) was used for multiple comparisons of two variables. Statistical significance was considered p < 0.05. Data are presented as the mean ± standard error of the mean (SEM).

[0128]

[0129] 2. Experimental Results

[0130]

[0131] 2.1 Characteristic Analysis

[0132]

[0133] Surface SEM images of core-shell structured PCL-gelatin (PG), PCL-gelatin-heparin (PGH), and PCL-gelatin-heparin-VEGF (PGHV) vascular grafts show smooth, bead-free fiber morphology, as shown in Figures 1A, 1B, and 1C, respectively. The distribution of VEGF within the shell of the fibers was traced using a confocal fluorescence microscope (Olympus, FV10i-W, Japan). The fluorescence images of core-shell fibers containing VEGF and FITC-BSA are shown in Figure 1D. The emitted green fluorescence indicates the presence of proteins within the shell of the fibers. The average fiber diameters of PG, PGH, and PGHV are 0.19±0.01 μm (Figure 1E), 0.30±0.014 μm (Figure 1F), and 0.42±0.04 μm (Figure 1G), respectively. A cross-sectional SEM image of the PGHV graft is shown in Figure 1I. The cross-sectional image shows the porous structure of the graft, but the fibers near the lumen (indicated by the double yellow arrows) were almost completely dissolved, creating a slightly smoother surface (Figures 2C and 2D) that was conducive to endothelial cell adhesion. The inner diameter of the graft was approximately 2 mm. A higher-magnification cross-sectional SEM image of the PGHV graft is shown in Figure 2. Loading VEGF and heparin into the fiber shell enabled faster release and ensured long-term stability of the graft. Conventional electrospinning, when loading biologically active molecules into the fiber, leads to rapid degradation, making it unsuitable for long-term applications. In contrast, in core-shell fibers, the shell can degrade, releasing the bioactive molecules. However, the mechanically strong core prevents graft failure during long-term application. Figure 12A depicts the release mechanism of bioactive molecules from core-shell fibers.

[0134] FTIR analysis confirmed that all materials were successfully incorporated into the core-shell structured graft (Fig. 3A). The graft was characterized by -COO vibration (1400 cm -1 ), -C=O stretching (1727cm) -1 ) and -CH2 stretching (2865 cm-1 ) showed prominent PCL characteristic peaks such as (Gomes, SR, et al., 2015).

[0135] The presence of gelatin was confirmed by the peaks of amide A (3300 cm-1), amide I (1650 cm-1), and amide II (1540 cm-1) (Al Fahad, MA, et al., 2023). FTIR spectra showed that PGHV nanofibers had a characteristic heparin absorption peak amide I (1640 cm -1 ) was shown to have.

[0136] Water contact angle tests showed that adding gelatin significantly reduced the contact angle (Fig. 3B). The contact angle of PCL was approximately 128°, indicating that it was inherently hydrophobic.

[0137] The addition of gelatin significantly reduced the contact angle of each graft to less than 90°, further enhancing hydrophilicity. It has been well-documented that hydrophilic grafts can promote cell adhesion and proliferation.

[0138] Heparin release curves (Figure 3C) show that the cumulative percentages of heparin released from the PGH and PGHV grafts after 25 days were 82.47±4.43 and 79.47±1.54, respectively. No statistically significant differences in heparin release behavior were found between the two groups. It is important to note that nearly 50% of the heparin was released in both groups after 5 days. Therefore, it is thought that impregnating the fiber shell structure with heparin can induce explosive heparin release. This explosive release of heparin is desirable for vascular grafts because it eliminates concerns about impending thrombosis (Bhakta, G., et al., 2012). VEGF release curves (Figure 3D) show that the cumulative percentage of VEGF released after 25 days was 86.25±1.19. Nearly 40% of the VEGF was released within 10 days. VEGF levels range from 119 to 238 pg / mL in healthy individuals, but are much higher in various cancers (Kut, C., F. Mac Gabhann, and A. Popel, 2007). In the present example, the VEGF concentration in PBS was 208.38 pg / mL after 5 days. The release patterns of both VEGF and heparin showed that controlled release of these molecules was achieved within a 4-week period, at which point the degradation profile of the implant was rapid (Figure 3E). We also measured the release patterns of heparin and VEGF when loaded into the fiber core (Figure 2). After 25 days, only approximately 55% of heparin and 29% of VEGF were released from the core, and the release pattern was found to be very slow compared to the release pattern of the shell, which achieved burst release.

[0139] To establish the stability and lifespan of the manufactured grafts, their degradation behavior was evaluated (Fig. 3E). After 6 weeks, the residual mass ratios of the PG, PGH, and PGHV vascular grafts were 83.22±0.99, 81.14±0.55, and 84.30±0.31, respectively. No statistically significant differences were found among the three groups. It is important to note that after 4 weeks, the residual mass ratios of the PG, PGH, and PGHV vascular grafts were 84.28±1.13, 82.05±0.60, and 85.65±2.08, respectively. The values ​​at the last two time points were almost similar. The degradation values ​​at 4 and 6 weeks were almost identical, likely because all gelatin degraded within 4 weeks, whereas PCL, a long-term degrading material, maintained a similar degradation rate even after 6 weeks. One of the major challenges when using small-diameter vascular grafts is their degradation over time, which can lead to graft failure. This phenomenon may be due to the fact that small-diameter vascular grafts experience high shear stress and low blood flow rates, which can lead to thrombosis or inflammation (Cecchi, E., et al., 2011). SEM images of degraded fibers, presented in Figures 3F-2H, show that the fiber shell began to degrade after 5 days. After 25 days, the exposed, clean PCL core and a portion of the remaining shell can be observed. It is well known that PCL fibers are suitable for long-term implantation due to their slow degradation and high mechanical properties (Vieira, A., et al., 2011; Shamsah, AH, et al., 2020; Pitt, C., et al., 1981). Therefore, the exposed PCL core is expected to provide sufficient mechanical support to the graft and maintain its integrity for long-term implantation.

[0140]

[0141] 2.2 Mechanical strength of vascular grafts

[0142]

[0143] The tensile properties of the grafts were evaluated in both the longitudinal (L) and circumferential (C) directions (Fig. 4A). The longitudinal maximum tensile strengths of the PG, PGH, and PGHV grafts were 8.83±0.89 MPa, 7.36±0.59 MPa, and 5.28±0.60 MPa, respectively (Fig. 4C).

[0144] The ultimate tensile strength values ​​in the circumferential direction of the PG, PGH, and PGHV grafts were 8.78±0.50 MPa, 6.587±0.44 MPa, and 6.59±0.41 MPa, respectively (Fig. 4C). The percentage of elongation values ​​in the longitudinal direction of the PG, PGH, and PGHV grafts were 158.0±16.26, 105.0±8.72, and 145.7±12.99, respectively (Fig. 4D). The percentage of elongation in the radial direction of the PG, PGH, and PGHV grafts was 104.3±17.14, 145.3±7.446, and 125.7±15.68, respectively (Fig. 4D). The Young's modulus in the longitudinal direction of the PG, PGH, and PGHV grafts was 20.27±1.19 MPa, 25.98±1.56 MPa, and 18.87±2.16 MPa, respectively (Fig. 4E). The Young's modulus in the radial direction of the PG, PGH, and PGHV grafts was 28.13±1.97 MPa, 19.19±1.75 MPa, and 32.70±2.31 MPa, respectively (Fig. 3I). The suture retention forces of the PG and PGHV grafts were 6.07±0.47 N and 5.75±0.66 N, respectively (Fig. 4F). Fiber diameter is hypothesized to influence the mechanical properties of the graft, such as tensile strength, compliance, and burst pressure. Smaller fiber diameters generally result in greater tensile strength and flexibility, properties required for successful vascular grafting (Pai, ​​C.-L., MC Boyce, and GC Rutledge, 2011).

[0145] The data also proved consistent with the reported theory. PG had the smallest average fiber diameter and higher tensile strength and elongation values ​​compared to PGH and PGHV grafts. The tensile strength decreased with increasing fiber diameter of PGH and PGHV. In addition, the ultimate tensile strength of the PGHV graft was 5.28 ± 0.60 MPa (L) and 6.59 ± 0.41 MPa (R); these values ​​are very similar to those of the native human artery (4.3 ± 1.8 MPa) (Stekelenburg, M., et al., 2009). Notably, the tensile properties of the vascular grafts in the longitudinal and radial directions were different due to the random orientation of the fibers (Al Fahad, MA, et al., 2023).

[0146]

[0147] 2.3 Blood flow simulation

[0148]

[0149] Blood flow simulation analysis (Figure 4G) shows that the blood velocity near the lumen wall can be approximately 0.13–0.44 m / s. The blood velocity can increase radially in a gradual manner toward the center. The velocity within the center can be approximately 0.6–0.7 m / s. This is consistent with the human coronary and cerebral artery blood flow velocity, which ranges from 0.2 to 0.72 m / s (Klarhφfer, M., et al., 2001; Coverdale, NS, et al., 2014). The stress range within the graft wall is approximately 1.65–1.7X10 4 Pa (Fig. 4H), which was much lower than the tensile stress values ​​obtained from the tensile strength test (5.28±0.60 MPa (L) and 6.59±0.41 MPa (R)). Therefore, it can be assumed that the graft material is sufficiently robust to handle hemodynamic stress.

[0150]

[0151] 2.4 In vitro hemocompatibility

[0152]

[0153] Heparin-loaded vascular grafts (PGH and PGHV) showed significantly reduced hemolysis rates compared to pure PG grafts (p<0.05) (Fig. 5A). The hemolysis rate of PG was 2.81±0.30%, which was less than 5%, and pure PG grafts themselves were hemocompatible according to ISO 10993-4 guidelines (Zhou, Y., et al., 2023). The hemolysis rates of PGH and PGHV were 1.57±0.30% and 1.65±0.16%, respectively. Thus, all grafts were confirmed to have excellent hemocompatibility. The BCI value reflects the proportion of uncoagulated RBCs in the sample solution (Teng, L., et al., 2021). A higher BCI value indicates a slower blood clotting rate (Du, X., et al., 2020). After 30 minutes, the BCI of the heparin-loaded grafts was significantly higher than that of the pure PG system (p<0.001). The BCIs of PG, PGH, and PGHV were 61.04±2.91, 82.96±2.36, and 78.14±1.95, respectively (Fig. 3B). The data demonstrate that the heparin-loaded grafts inhibited thrombosis. The BCI of PGHV was slightly smaller than that of PGH, but the difference was not statistically significant. The prothrombin time (PT) (Fig. 5D) and activated partial thromboplastin time (APTT) (Fig. 5E) were measured after 1 hour of co-incubation with the electrospun grafts and human plasma. Both heparin-loaded grafts (PGH and PGHV) exhibited significantly higher PT (p<0.5) and APTT (p<0.5) than the control group. These findings demonstrate the anticoagulant properties of the grafts. SEM images of platelets attached to the graft surface showed that almost all platelets were inactivated even after 1 hour of incubation (Figures 5E, 5F, and 5G). Typically, inactivated platelets are round and, upon activation, form pseudopodia, further inducing thrombosis (Figure 5D). The total number of platelets attached to the surface of heparin-loaded grafts was significantly lower than that of pure PG grafts (p<0.001) (Figure 5C).The total platelet counts (per 103 μm2) attached to PG, PGH, and PGHV grafts were 84±3, 19±4, and 25±3, respectively.

[0154]

[0155] 2.5 In vitro biocompatibility

[0156]

[0157] The MTT assay presented in Figure 5I shows no significant differences in cell proliferation after 1 and 3 days for the PG, PGH, and PGHV vascular grafts. However, after 7 days, cell proliferation was significantly reduced in the PG and PGH grafts (p<0.01) compared to the control (tissue culture plate), but significantly increased in the PGHV graft (p<0.001). The improved cell proliferation in the PGHV grafts may be due to the release of VEGF from the grafts into the culture medium. Initially, the proliferation rate was similar to that in the control, but as more VEGF was released from the grafts, proliferation increased significantly. Confocal cell proliferation images, as shown in Figure 5K, also correspond to the MTT results. All cells appeared healthy and formed spindle-shaped, filiform pseudopodia over time. The percentage of F-actin area calculated from the confocal images of PG, PGH, and PGHV grafts (Fig. 5M) was 56.26±5.85%, 57.66±3.45%, and 80.56±2.76%, respectively. Live / dead analysis was performed to count the number of live and dead cells in the grafts (Fig. 5L). The number of live / dead cells present in PG, PGH, and PGHV grafts was 2057±68 / 363±18, 1966±29 / 383±35, and 2110±50 / 320±27, respectively, as shown in Fig. 5N. The cell viability of PG, PGH, and PGHV was 86.04±2.21, 84.17±2.53, and 89.68±2.28, respectively (Fig. 5J). Cell migration tests showed that the addition of VEGF significantly increased the rate of endothelial cell migration after 24 hours (Fig. 6B). Both MTT and live / dead cell viability tests suggested that the graft was biocompatible and could be further studied for in vivo applications.

[0158]

[0159] 2.6 Patency of vascular grafts

[0160]

[0161] After implantation, all rats were thoroughly examined for blood leakage. No leakage was found (Figs. 7A and 5B), and all rats remained in good physical condition throughout the implantation period. Doppler ultrasonography was used to measure blood velocity and patency of the implanted grafts (Fig. 7D). After 2 months, the patency rates of the PG and PGHV grafts were 77.5% and 97%, respectively, and after 4 months, the patency rates of the grafts decreased to 67.5% and 89.5%, respectively (Fig. 7C). These results suggest the possibility of stenosis in the lumen of the PG graft. However, even after 4 months, the PGHV graft maintained nearly 90% of its patency. A recent study showed that vascular grafts coated with a Glycocalyx-like hydrogel maintained approximately 85% of their patency 32 days after implantation (Dimitrievska, S., et al., 2020). In another recent study, Wang et al. The patency of dopamine-mediated vascular grafts was approximately 65% ​​at 3 months after implantation (Wang, Z., et al., 2023). A patency rate of 90% was achieved, which is excellent compared to other studies, especially considering the extended graft duration. Andreadis ST, Swartz DD, and their groups developed a VEGF-heparin-loaded TEVG and achieved remarkable patency similar to that of the native sheep carotid artery (Koobatian, MT, et al., 2016). After 2 months, the peak blood velocity of the PG and PGHV grafts was 38.2 cm , respectively. -1 and 20.8cm -1 After 4 months, the speed was 61.5 cm each. -1 and 30.4cm -1 increased to (Fig. 7E). After 2 months, the average blood velocity of the PG and PGHV grafts was 23.3 cm -1 and 13.4cm -1 After 4 months, the speed was 39.8 cm each. -1 and 17.1cm -1increased (Fig. 7F). Blood velocity increased significantly in the PG graft due to stenosis, but remained normal in the PGHV graft. Fig. 7G schematically illustrates normal laminar blood flow, and Fig. 7H graphically illustrates turbulent blood flow due to thrombosis or intimal hyperplasia within the graft. Consequently, the cause of stenosis is explained to be intimal hyperplasia.

[0162]

[0163] 2.7 ECM Reconfiguration

[0164]

[0165] For histological staining, the extracted grafts were dissected from the middle portion. Figures 7I and 8C show H&E stained images. As shown in the images, neointima was formed in both grafts. The neointima thicknesses of the PG and PGHV grafts were 50.90±4.69 μm and 25.97±1.55 μm after 2 months, respectively, and 65.99±8.99 μm and 34.07±4.09 μm after 4 months, respectively (Figure 7N). After both time points, the thickness of the newly formed intima was significantly higher in the PG graft than in the PGHV graft. This increased thickness impeded blood flow and increased blood velocity, which was supported by the Doppler ultrasound data. In contrast, the neointima was considerably thinner and had little effect on blood velocity. Recent studies have reported that the neointimal thickness of hyaluronan-loaded PCL and PLCL-loaded vascular grafts (D=2 mm) was approximately 30 μm and 200 μm, respectively (Qin, K., et al., 2020). Another recent study showed that the neointimal thickness of PLLA grafts (D=1.5 mm) loaded with platelet-rich plasma (PRP) was approximately 40 μm 4 weeks after implantation (Li, G., et al., 2023). Another recent study confirmed that the neointimal thickness of PCL grafts (D=2 mm) loaded with recent ECM was approximately 60 μm 12 weeks after implantation (Liu, S., et al., 2023). Polyglycerol-sebacate-loaded tropoelastin vascular grafts (D = 1.5 mm) implanted in rats also regenerated a 100-μm-thick neointima, but patency remained similar to native vascular grafts due to the rapid degradation of the graft (Wang, Z., et al., 2022). Because regeneration of an abnormally thick neointima indicates intimal hyperplasia, a thinner neointima with adequate endothelialization is a desirable feature in small-diameter vascular grafts. All recent studies have shown that the novel PGHV grafts regenerated a significantly thinner neointima.However, the low porosity and high density microstructure of the two implanted grafts hindered cell infiltration. The porous structure of the vascular grafts facilitates the infiltration of endothelial progenitor cells into the vessel wall and subsequent capillary formation, but the cell infiltration into the graft wall may make it difficult to achieve endothelial lining of the lumen of the graft (Hong-De Wu, M., et al., 1995; Sαnchez, PF, EM Brey, and JC Briceρo, 2018). The porous structure may also facilitate the migration of blood cells, which can cause thrombosis in later stages. Since one of the main objectives of the present invention was to achieve endothelial lining of the lumen of the graft, a high porosity structure was not considered in the present invention to achieve this goal.

[0166] Masson's trichrome staining, shown in Figure 7J, demonstrates that the neointima of the PG graft was thicker, resulting in more collagen (blue) deposition in the PG graft compared to the PGHV graft. Figure 7K shows Von Kossa staining for calcium (dark brown) deposition. After 4 months, the PG graft showed significant calcium deposition. Abnormal proliferation of SMCs is known to be a major cause of vascular calcification. Figures 7L and 5M show the extent of elastin (black) and glycosaminoglycans (orange-red) within the vascular graft wall. Elastin fibrils impart elasticity and resilience, whereas collagen fibrils in the media layer are oriented circumferentially to provide mechanical support. The presence of elastin within the graft wall was observed in a rat aortic model study. Achieving the presence of elastin in diseased human patients or diseased mice can be a very challenging goal. GAGs fill the gaps between fibrils, hydrate the ECM, and can bind to various cell surface receptors with high selectivity. Here, the ECM, containing elastin, collagen, and GAGs, is tightly and orderly organized after transplantation, exhibiting a structure similar to that of the original artery.

[0167]

[0168] 2.8 Endothelialization characteristics of vascular grafts

[0169]

[0170] The endothelium coverage of the explanted grafts was assessed by scanning electron microscopy (SEM) at three separate locations, including the proximal, middle, and distal regions (Fig. 8A). After 2 months, the proximal and distal regions of the PG graft were coated with a confluent layer of cells, whereas the middle region lacked a confluent layer. The cell layer was identified as EC, and its morphology was elongated by blood flow. Furthermore, the native lumen wall of the rat abdominal aorta exhibited a similar texture (Fig. 9B). However, after 4 months, a confluent layer of EC was present throughout the lumen of the PG graft. In contrast, the entire lumen of the PGHV graft exhibited an endothelial layer within 2 months. This layer was also evident after 4 months. Fig. 8B schematically illustrates the endothelialization behavior in the lumen of the PG and PGHV grafts. Fig. 8D demonstrates the presence of the endothelialization marker CD31 throughout the entire cross-section of the vascular graft. Neointima regeneration was nearly uniform but thicker in the PG compared to the PGHV. Figure 8E schematically illustrates (i) the neointimal proliferation phenomenon and (ii) the perfect alignment of the SMC and EC layers of the vascular graft. Immunofluorescence staining for vWF, CD31, and ICAM 1 was used to assess the endothelial coverage. All markers were specific for endothelial cells (Kerdjoudj, H., et al., 2007; Lawson, C. and S. Wolf, 2009; Gao, G., et al., 2017). Figure 10A shows that the CD31 marker was expressed in the lumen (middle part) of the PGHV graft after 2 months, but CD31 was absent in the middle part of the PG graft. The results were consistent with the SEM images in Figure 8A, which showed that the middle part of the PG graft had no endothelial layer after 2 months. After 4 months, the PG graft showed CD31 expression in the middle part of the lumen.

[0171] Figure 10B shows ICAM1 expression, and Figure 10C shows vWF expression in the lumen of the graft. Both markers followed a similar expression trend to the vWF marker. Finally, it can be predicted that PGHV grafts loaded with VEGF and heparin can induce more rapid endothelialization than pure PG grafts. We hypothesize that both anastomotic in-growth and fallout endothelialization mechanisms are involved during vascular reconstruction with PGHV grafts (Figure 12B). In the trans-anastomotic mechanism, the host intima responds to the trauma caused by the graft surgery by growing from the anastomotic site toward the center of the graft. This process is a natural process, and in addition to host ECs, it also involves a complex web of chemotactic and mechanotransducing signals, including monocytes, platelets, and SMCs. According to the fallout endothelialization mechanism, circulating endothelial progenitor cells (EPCs) in the bloodstream can also contribute to endothelialization. VEGF significantly supports the endothelialization process by recruiting EPCs and differentiating them into mature ECs. Andreadis ST, Swartz DD, and their groups reported that VEGF can recruit circulating monocytes and differentiate them into endothelial cells (Smith Jr, RJ, et al., 2020). A recent study also reported the effects of VEGF released from bacterial nanocellulose vascular grafts on EC proliferation and migration (Hu, G., et al., 2022). In PGHV grafts, both mechanisms operated simultaneously to accelerate endothelialization, whereas in PG grafts, endothelialization occurred solely through the anastomosis process, resulting in a slower rate of endothelialization.

[0172]

[0173] 2.9 Regeneration of vascular smooth muscle

[0174]

[0175] Immunofluorescence staining for α-SMA and MYH11 was used to assess vascular smooth muscle regeneration. Figure 11A shows α-SMA expression in the lumen of vascular grafts. After 2 months, the smooth muscle cell layer was significantly thicker in PG grafts than in PGHV grafts. After 4 months, the thickness of PG grafts had nearly doubled, indicating neointimal hyperplasia, which resulted in increased blood velocity and decreased patency of PG grafts (Figure 7C). The number of cells found in PG was also significantly higher than in PGHV grafts, likely due to the thicker regenerated intima of PG (Figure 10D). Figure 11C shows that significantly more α-SMA+ cells were present in PG grafts than in PGHV grafts (p<0.001). Figure 11D shows significantly higher expression of α-SMA regions in PG grafts compared to PGHV grafts after 2 months (p<0.01) and 4 months (p<0.001). Figure 11B shows MYH11 expression in the lumen of vascular grafts. MYH11 marker expression followed a similar expression trend to α-SMA. Figure 11E shows that a significantly higher number of MYH11+ cells were present in PG grafts compared to PGHV grafts at both time points (p<0.001). Figure 11F shows that the MYH11 expression area was significantly higher in PG grafts compared to PGHV grafts after 2 months (p<0.001) and 4 months (p<0.001). Delayed endothelialization may contribute to the intimal hyperplasia of PG grafts. It is well known that endothelial cells regulate the proliferation of smooth muscle cells. Inadequate endothelialization leads to abnormal smooth muscle cell proliferation, ultimately leading to intimal hyperplasia. It was anticipated that vascular remodeling in PG grafts would occur solely through the anastomotic mechanism. This also induces intimal hyperplasia, including subsequent SMC proliferation through a complex inflammatory response and migration.In the case of PGHV transplantation, it can be roughly assumed that rapid endothelialization can occur through both the endovascular and transanastomotic mechanisms, which control smooth muscle cell proliferation and prevent intimal hyperplasia. This suggests that these endothelialization mechanisms may play a key role in PGHV transplantation.

[0176]

[0177] 2.10 Inflammatory response to vascular grafts

[0178]

[0179] Pro-regenerative M2 macrophages and pro-inflammatory M1 macrophages were counted using immunofluorescence staining for the expression of CD206 (Fig. 11D) and CD68 (Fig. 11C). The percentages of CD68+ cells within the anastomotic sites of PG and PGHV grafts after 2 months were 16.67±2.33 and 16±3.6, respectively (Fig. 11I). However, CD68 expression significantly decreased after 4 months in both grafts. The percentages of CD68+ cells within the anastomotic sites of PG and PGHV grafts after 4 months were 5±1.16 and 4.67±1.2, respectively (Fig. 11I). CD206 (an M2 macrophage marker) expression was lower in both grafts after 2 months and increased after 4 months (Fig. 11J). All images and graphs demonstrate similar expression patterns of M1 and M2 macrophage markers. Both grafts can modulate inflammatory responses by shifting them to an M2 regulatory pro-regenerative state. The ability of M2 macrophages to develop into SMCs, which play a role in vascular graft formation, is supported by some evidence, despite current discrepancies (Hristov, M., W. Erl, and PC Weber, 2003; Smith Jr, RJ, et al., 2020). Macrophages may play a key role in SMC development. Meanwhile, grafts measuring between 7 and 10 mm in length are not suitable for implantation in a rat abdominal aorta model.

[0180]

[0181] While specific aspects of the present invention have been described in detail above, it will be apparent to those skilled in the art that these specific descriptions merely represent preferred embodiments and are not intended to limit the scope of the present invention. Therefore, the substantial scope of the present invention is defined by the appended claims and their equivalents.

[0182]

[0183] The national research and development projects that supported this invention are as follows.

[0184] [Project ID] 1345235458

[0185] [Assignment Number] 2015R1A6A1A0303252223

[0186] [Ministry Name] Ministry of Education

[0187] [Name of Project Management (Specialist) Institution] National Research Foundation of Korea

[0188] [Research Project Name] University Key Research Institute Support Project (Science and Engineering)

[0189] [Research Project Title] Development of Metal-Ceramic-Polymer Hybrid Biomaterials for Clinical Treatment

[0190] and tissue regeneration research

[0191] [Name of the project performing organization] Soonchunhyang University Industry-Academic Cooperation Foundation

[0192] [Research Period] March 1, 2023 - February 29, 2024

[0193]

[0194] [National Research and Development Project Supporting This Invention]

[0195] [Project ID] 1711190896

[0196] [Assignment Number] 2021R1G1A109489413

[0197] [Ministry Name] Ministry of Science and ICT

[0198] [Name of Project Management (Specialist) Institution] National Research Foundation of Korea

[0199] [Research Project Name] Individual Basic Research (Ministry of Science and ICT)

[0200] [Research Project Name] Growth Factor-Loaded Decellularized ECM / Electrospun PCL-Based Bilayer Structure

[0201] Manufacturing and preclinical studies of artificial blood vessels

[0202] [Name of the project performing organization] Soonchunhyang University Industry-Academic Cooperation Foundation

[0203] [Research Period] March 1, 2023 - February 29, 2024

Claims

1. A step of manufacturing a core structure of nanofibers by dissolving polycaprolactone (PCL) in dichloromethane; A step of preparing a shell structure of nanofibers by dissolving polycaprolactone (PCL) and gelatin in 2, 2, 2-trifluoroethanol (TFE); A step of preparing a solution of each of the core structure and shell structure of the nanofiber using a magnetic stirrer; A step of adding VEGF to a polycaprolactone (PCL) and gelatin solution; A step of stirring each solution of the core structure and shell structure of the nanofiber on a magnetic stirrer; and A method for producing core-shell structured nanofibers for preventing endometrial hyperplasia, comprising the step of adding heparin to a polycaprolactone (PCL) and gelatin solution.

2. In paragraph 1, In the above stirring step, A method for producing core-shell structured nanofibers, wherein the mixing ratio of core structure and shell structure is 70:

30.

3. In paragraph 1, A method for producing core-shell structured nanofibers, wherein the shell structure comprising polycaprolactone (PCL) and gelatin induces a constant release of VEGF and heparin.

4. In paragraph 1, A method for producing core-shell structured nanofibers, wherein the polycaprolactone (PCL) core maintains mechanical strength.

5. In paragraph 1, A method for producing core-shell structured nanofibers, wherein the core-shell structured nanofibers reduce thrombosis.

6. In paragraph 1, A method for producing core-shell structured nanofibers, wherein the core-shell structured nanofibers induce endothelialization.

7. Nanofibers manufactured according to any one of the manufacturing methods of clauses 1 to 6.

8. A graft composition comprising the nanofiber of clause 7.

Images