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. 2019 Jan 10;33(4):5089–5100. doi: 10.1096/fj.201801856R

Implantation of VEGF-functionalized cell-free vascular grafts: regenerative and immunological response

Randall J Smith Jr *, Tai Yi †, Bita Nasiri ‡, Christopher K Breuer †, Stelios T Andreadis *,‡,§,1
PMCID: PMC6436645  PMID: 30629890

Abstract

Recently, our group demonstrated that immobilized VEGF can capture flowing endothelial cells (ECs) from the blood in vitro and promote endothelialization and patency of acellular tissue–engineered vessels (A-TEVs) into the arterial system of an ovine animal model. Here, we demonstrate implantability of submillimeter diameter heparin and VEGF-decorated A-TEVs in a mouse model and discuss the cellular and immunologic response. At 1 mo postimplantation, the graft lumen was fully endothelialized, as shown by expression of EC markers such as CD144, eNOS, CD31, and VEGFR2. Interestingly, the same cells coexpressed leukocyte/macrophage (Mϕ) markers CD14, CD16, VEGFR1, CD38, and EGR2. Notably, there was a stark difference in the cellular makeup between grafts containing VEGF and those containing heparin alone. In VEGF-containing grafts, infiltrating monocytes (MCs) converted into anti-inflammatory M2-Mϕs, and the grafts developed well-demarcated luminal and medial layers resembling those of native arteries. In contrast, in grafts containing only heparin, MCs converted primarily into M1-Mϕs, and the endothelial and smooth muscle layers were not well defined. Our results indicate that VEGF may play an important role in regulating A-TEV patency and regeneration, possibly by regulating the inflammatory response to the implants.—Smith, R. J., Jr., Yi, T., Nasiri, B., Breuer, C. K., Andreadis, S. T. Implantation of VEGF-functionalized cell-free vascular grafts: regenerative and immunological response.

Keywords: monocytes, endothelialization, immunoengineering, inflammation

Over the past decade, several groups have demonstrated the feasibility of completely acellular, off-the-shelf grafts in various animal models as well as human clinical trials. Decellularized and devitalized tissue-engineered constructs have been utilized with increasing frequency and have demonstrated improved patency and regeneration potential (1–13). Devitalized constructs are based on bioengineered tissue constructs that are stripped of the cellular components while leaving extracellular matrix (ECM) components intact and then implanted into the animal model of choice (14). It is important to note that these vessels, although successful, still require intensive culture times—a key issue when considering the ability of a graft to be truly off-the-shelf ready. In addition to devitalized and decellularized grafts, nonbiologic grafts composed of various polymeric biomaterials have also been used to engineer cell-free vascular grafts (15–23). Besides biocompatibility, lack of immunogenicity, and mechanical properties matching those of native vessels, these materials must promote endothelialization of the lumen to achieve patency and promote development of the vascular wall through extensive, long-term remodeling. These qualifications limit the potential choices of polymer-based biomaterials.

The biggest challenge facing implantation of vascular grafts is endothelialization of the lumen in order to prevent thrombosis and occlusion (24). To this end, growth factors or other proteins have been immobilized to the luminal surface. One such growth factor, SDF1-α, has been used to home circulating stem cells to the graft lumen via the SDF1-α receptor CXCR4. However, these studies showed incomplete endothelialization, especially in the center of the grafts, indicating that endothelialization may be limited to anastomotic end ingrowth of endothelial cells (ECs) (25, 26).

In addition to growth factors, synthetic peptides such as REDV and full-length ECM proteins such as fibronectin have also been used to promote endothelialization (27–30). Use of fibronectin resulted in incomplete endothelialization in the middle of the graft, possibly due to a lack of specificity to integrin-binding domain RGD, which may allow a wide variety of cells to bind and outnumber the scarcely present ECs in the blood (26). Indeed, because platelets also carry the integrin α5β1 that binds the RGD domain, use of full-length fibronectin presents the risk of homing thrombogenic cells to the graft lumen (29, 31). Furthermore, the use of the glycosaminoglycan heparin has been utilized in small animal models to prevent thrombosis (8, 32, 33) but has failed to prevent thrombosis in large animal models, such as sheep (34).

In our laboratory, we developed a truly off-the-shelf vascular graft that is based on small intestinal submucosa (SIS) with immobilized heparin and VEGF on the graft lumen to capture VEGF receptor–expressing circulating angiogenic cells from the blood. When implanted into the carotid arteries of an ovine animal model, such small-diameter (4.5 mm) grafts exhibited high patency rates, fully endothelialized within 1 mo, and developed a functional and contractile medial layer by 3 mo postimplantation (34–36). In this study, we utilized a small-animal model to examine the role of the inflammatory response in promoting endothelialization and tissue regeneration of VEGF-decorated acellular tissue–engineered vessels (A-TEVs). Although not as clinically relevant as sheep, mice may be useful in addressing mechanistic questions mostly because of the availability of multiple transgenic models that are not available in sheep.

In this study, we demonstrate the implantability of submillimeter diameter vascular grafts comprising SIS functionalized with heparin and VEGF in a mouse animal model. In contrast to grafts containing only heparin, the presence of immobilized VEGF promoted formation of the luminal endothelial and medial layers, with spatial organization that recapitulated the structure of the native artery. VEGF also promoted an anti-inflammatory micro-environment, which might have contributed significantly to the enhanced regeneration of cell-free vascular grafts.

MATERIALS AND METHODS

Graft production

Grafts were engineered using SIS (Cook Biotech, West Lafayette, IN, USA) wrapped around a perforated plastic mandrel and further subjected to cross-linking and drying steps as shown in Fig. 1. Initially, plastic tubing with an outer diameter of 800 µm was perforated with 12-gauge needles in an equally distributed pattern. SIS was then rolled around the mandrel manually, tightening each turn as it was rolled for at least 3 times. SIS has a thickness of ∼50 μm. The SIS was then allowed to dry on the mandrel for 2 h at room temperature before it was cross-linked in a solution containing 20 mM 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide (MilliporeSigma, Burlington, MA, USA), 10 mM N-hydroxysuccinimide (MilliporeSigma), in 50 mM 2-ethanesulfonic acid buffer (pH 4.5) for 2 h at room temperature with gentle rocking. The partially cross-linked SIS and mandrel were removed from the solution and assembled into a vacuum device consisting of a 1-mm diameter tubing, a vacuum line, and a clamp. The mandrel was first connected to the vacuum line, and the larger tubing was then slipped over the SIS and mandrel and sealed around the vacuum line piece. Finally, a clamp was placed at the opposite end of both the SIS and mandrel and the larger sheath tubing. Application of vacuum for 24 h allowed the SIS to dry through the perforations on the mandrel, and the dry graft was then easily slipped off of the mandrel for further processing. The final dried SIS tube had a diameter ranging from 850 to 900 μm when assessed under the surgical microscope right before implantation.

Figure 1.

Figure 1

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Schematic of submillimeter-sized VEGF-decorated A-TEV. EDC, 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide; NHS, N-hydroxysuccinimide.

Functionalization of SIS graft

SIS conduits were further processed using a protocol that we previously published (34, 36). Briefly, the grafts were placed in a solution of 5-mg/ml heparin (low MW; MilliporeSigma), 20-mM 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide, and 10 mM N-hydroxysuccinimide in 50 mM 2-ethanesulfonic acid buffer (pH 4.5). The reaction proceeded for 24 h at room temperature with gentle rocking. After repeated rounds of washing with sterile water, the graft was either stored in PBS [SIS + heparin group (SH) graft] or further functionalized with 0.5-mg/ml VEGF (produced in house) in PBS for 2 h at 37°C. VEGF binds to the heparin via the heparin-binding domain [SIS + heparin + VEGF group (SHV) graft]. Prior to implantation, the vessels were washed in PBS to remove unbound VEGF.

Arterial interpositional tissue-engineered vascular graft implantation

All animal experiments were approved by Nationwide Children’s Hospital institutional guidelines for the use and care of animals. In this experiment, there were 2 groups: heparin-only control (SH) and heparin/VEGF-coated grafts (SHV) (n = 5 each). A-TEVs were implanted into 8–10-wk-old female C57BL/6 mice. The mice were anesthetized using ketamine xylazine cocktail, with ketoprofen as preanesthesia analgesic. Hair in the surgical area was removed by shaving, and the area was disinfected by betadine and alcohol pads. A midline laparotomy incision from below the xyphoid to the suprapubic region was made, and a self-retaining retractor was inserted. The intestines were wrapped in saline-moistened gauze and retracted. The infrarenal aorta and inferior vena cava were bluntly defined.

Microsurgery was performed using an operating microscope with zoom magnification. The aorta was separated from the inferior vena cava, and vascular control was achieved with microvascular clamps before the infrarenal aorta was transected. An aortic interposition graft 2 mm in length was implanted with proximal and distal end-to-end anastomoses using sterile 10-0 monofilament sutures on tapered needles. Any hemorrhaging was controlled by applying topical, absorbable, sterile hemostatic agents (Surgicel). Then, intestines were returned to the abdominal cavity, and the abdominal musculature and skin were closed in 2 layers using 6-0 prolene sutures.

Ultrasound

To monitor graft patency during the implantation period, mice were imaged at 1 d, 2 wk, and 4 wk with a high-frequency Doppler ultrasound system (Vevo 2100; VisualSonics, Toronto, ON, Canada). After anesthetizing the mice (1.5% isoflurane; Baxter, Deerfield, IL, USA), the abdominal hair was clipped, and ultrasound gel (Aquasonic Clear, Parker Laboratories, Fairfield, NJ, USA) was applied on the abdomen. Long-axis images were acquired in both B mode and color Doppler, and the graft patency was determined by the presence of blood flow through the graft lumen.

Tissue collection and preparation for histology

The grafts were explanted at 1 mo postimplantation. After euthanizing the mice, the grafts were perfusion fixed with 10% formalin, placed in 10% formalin overnight, and embedded in paraffin. The paraffin-embedded sections were stained with hematoxylin and eosin for histologic examination. Tissues were sectioned from anastomotic ends (observed via suture location) and tissue sections (5 μm thick) were collected starting at 15 μm after proximal suture site (proximal end) or 15 μm prior to distal suture site (distal end). The middle of grafts was at ∼1 mm after the proximal suture site.

Immunohistochemistry

Paraffin sections were first blocked with 5% (v/v) goat serum in PBS prior to incubation with the following primary antibodies: anti-VEGFR1 (1:100; Thermo Fisher Scientific, Grand Island, NY, USA), anti-VEGFR2 (1:100; Thermo Fisher Scientific), anti–smooth muscle actin (1:200; Thermo Fisher Scientific), anti-CD144 (1:50; Cell Signaling Technology, Danver, MA, USA), anti-CD16 (1:200; Abgent, San Diego, CA, USA), anti-CD14 (1:100; Abgent), anti-CD38 (1:200; Abcam, Cambridge, MA, USA), anti-EGR2 (1:100; Abcam), anti-CD31 (1:200; Thermo Fisher Scientific), and anti-eNOS (1:500; BD Biosciences, Franklin Lakes, NJ, USA) in 5% (v/v) goat serum in PBS overnight at 4°C. Following 3 washes, tissue sections were incubated with Alexa Fluor secondary antibodies (1:200; Thermo Fisher Scientific) for 1 h at room temperature. Nuclei were counterstained with Hoechst 33342 (1:200; Thermo Fisher Scientific) for 5 min at room temperature, and images were obtained with a Zeiss Axio Imager microscope (Carl Zeiss GmbH, Jena, Germany).

Analysis and statistics

Images of immunostained tissue sections were quantified using ImageJ software (National Institutes of Health, Bethesda, MD, USA). All cells per image were quantified using DAPI, and 5 images were used per tissue section (proximal, middle, distal) for each animal (n = 3 animals) for a total of 45 images per antibody panel. Luminal cells were defined as the topmost single cell layer, with wall and subluminal cells defined as all cells beneath the luminal cells. Statistical significance (P < 0.05) was calculated using paired Student’s t test.

RESULTS

Cell-free tissue-engineered vessels with very small inner diameters (diameter: 850–900 µm; length: 2 mm) were prepared from the clinical grade natural biomaterial SIS using perforated tubing with the same outer diameter (Fig. 1). The SIS was rolled around the mandrel 3 times to achieve a wall thickness of 150 μm (50 μm/layer), then cross-linked, dried, and vacuum pressed to seal the tubular construct into a single conduit. The conduit was then cross-linked with heparin (SH) and further functionalized with VEGF (SHV). As we previously reported (34), under these conditions, the surface concentration of VEGF on the SIS surface was 11.5 μg/cm2, and only 14% was released over 120 h. In addition, human VEGF could not be detected in the blood of sheep receiving the A-TEV, suggesting negligible release of immobilized VEGF from the graft into the circulation (unpublished results).

Grafts were then implanted interpositionally in the descending aorta of adult mice (Fig. 2). At the time of implantation, the diameter of the graft was larger than that of the native artery (700–800 μm). Upon closure, grafts were observed via ultrasound and Doppler radar at 24 h, 2 wk, and 4 wk postimplantation. At 4 wk, mice were euthanized, and the grafts were explanted for further analysis.

Figure 2.

Figure 2

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Gross anatomic images of A-TEVs prior to implantation, upon implantation, and at 4 wk after implantation. Black arrows represent anastomotic sites. Scale bars, 2 mm.

Ultrasound revealed that 2 SHV (n = 5) and 2 SH (n = 5) grafts occluded within the first 24 h. The remaining 3 grafts were patent throughout the 1-mo study, as evidenced by Doppler ultrasound (Fig. 3A). Interestingly, as time progressed, the initial diameter mismatch decreased toward native size, as evidenced by ultrasound measurements (Fig. 3B). Blood velocity within the grafts was consistent throughout the study at ∼160 ± 14 mm/s, as evidenced by color Doppler, demonstrating no evidence of stenosis (Fig. 3C).

Figure 3.

Figure 3

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Patency and blood flow measurements in implanted A-TEV. A) Ultrasound and color Doppler images of SHV and SH grafts at 0 d, 2 wk, and 4 wk postimplantation and representative graphs of blood velocity through the grafts. B) Diameter analysis over time as determined by ultrasound. C) Mean velocity over time as determined by ultrasound.

Explanted grafts were assessed via hematoxylin and eosin staining (Fig. 4). As shown in Fig. 4B, both the luminal cellular layer as well as the underlying remaining SIS (separated by the dotted line) differed significantly between SH and SHV grafts. Specifically, SHV grafts had a significantly thicker cellular layer (250 ± 24 µm) compared with SH grafts (170 ± 19 µm) (n = 10, P < 0.05), which correlated with increased cell density in SHV (3996 ± 550 cells/mm2) compared with SH grafts (2950 ± 450 cells/mm2) (n = 27 fields of view, n = 3 grafts, P < 0.05). In addition, ∼33% of the SIS remained in the SHV grafts compared with 73% remaining in the SH grafts at 1 mo postimplantation.

Figure 4.

Figure 4

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Histologic analysis of A-TEV explants. A) Hematoxylin and eosin analysis of explanted grafts. B) Measurements of thickness of cell layer and the remaining SIS scaffold. SIS and cell layer are separated by a dotted line.

SHV grafts exhibit native-like topological organization of the reconstituted lumen and vascular wall

Immunostaining for endothelial markers VEGFR1 (red) and VEGFR2 (green) indicates expression of VEGFR1 and VEGFR2 in the luminal cells and only VEGFR1 in cells immediately beneath the lumen of SHV grafts (Fig. 5A), which is similar to that observed in native arteries (Fig. 5B). In contrast, the demarcation was not clear in SH grafts, as VEGFR2 and VEGFR1 were expressed in the lumen and subluminal areas. In addition, SH grafts contained multiple layers of VEGFR1/VEGFR2-expressing cells.

Figure 5.

Figure 5

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SHV grafts exhibit native-like topological organization of the reconstituted lumen and vascular wall. A, B) Immunostaining for VEGFR1 (red) and VEGFR2 (green) in SHV and SH grafts (A) or native arteries (B). C, D). Immunostaining for α-SMA (red) and CD144 (green) in SHV and SH grafts (C) or native arteries (D). A, C) The arrow indicates 3 images from the proximal to the distal end of SHV and SH grafts. E) Image quantification and analysis for n = 45 images per antibody panel. Percent positive cells of DAPI positive cells per specified vessel area. L, graft lumen. Scale bars: 10 μm (white), 30 μm (red). *P < 0.05.

Similarly, there was a clear demarcation in the expression of EC marker CD144 (green) on the lumen and α-SMA (red) in the wall of SHV grafts, which is similar to that observed in native arteries (Fig. 5C, D). In contrast, both markers appeared in the luminal as well as subluminal areas of SH grafts (Fig. 5C). Notably, SHV grafts contained a continuous layer of CD144+ cells on their lumen, but in SH grafts, CD144 expression was not consistent across the entirety of the luminal surface.

SHV grafts regenerate a well-defined luminal endothelium coexpressing anti-inflammatory monocyte and macrophage proteins

Next, we sought to examine whether blood monocytes (MCs) infiltrated the grafts and whether the presence of VEGF on the graft lumen affected the phenotype of the graft-infiltrating macrophage (Mϕ). Interestingly, immunostaining for CD14 (green) and CD16 (red) revealed a significant difference (P < 0.05) between SHV and SH grafts (Fig. 6A). Specifically, the lumen of SHV grafts contained doubly stained CD14+/CD16+ cells (100%, n = 153 luminal cells), with few such cells also present in the wall (4 ± 1%, n = 1463 wall cells). In contrast, SH grafts displayed a stark difference (CD14+/CD16+: 10 ± 2%, n = 201 luminal cells), with the overwhelming majority of cells on the lumen and the wall expressing only CD14 (CD14+/CD16−: 73 ± 10%, n = 1486 total luminal and wall cells). As expected, little or no expression of CD14 or CD16 was observed in native arteries (Fig. 6B).

Figure 6.

Figure 6

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Coexpression of MC and EC markers by the cells populating the SHV grafts. A, B) Immunostaining for the MC markers CD14 (green) and CD16 (red) in SHV and SH grafts (A) or native arteries (B). C, D) Immunostaining for M1 (Mϕ) marker CD38 (red) and the M2/EC marker EGR2 (green) in SHV and SH grafts (C) or native arteries (D). A, C) The arrow indicates 3 images from the proximal to the distal end of SHV and SH grafts. E) Image quantification and analysis for n = 45 images per antibody panel. Percent positive cells of DAPI positive cells per specified vessel area. L, graft lumen. Scale bars: 10 μm (white), 30 μm (red). *P < 0.05.

This result prompted us to examine the nature of Mϕs in the grafts by costaining for the Mϕ M1 marker CD38 (red) and the M2/EC marker EGR2 (green) (Fig. 6C, D). Immunostaining indicated a significant difference (P < 0.05) between SH and SHV grafts. The luminal surface of SHV grafts consisted entirely of EGR2+ cells (100%, n = 164 luminal cells), with CD38+ cells present only beneath the surface, which is similar to the expression observed in native vessels. Some EGR2+ cells were also present in the subluminal areas. In contrast, EGR2 expression was significantly reduced in SH grafts and limited to only subluminal areas (11 ± 8%, n = 1486 wall cells, P < 0.05), whereas all luminal cells (100%, n = 201) and several subluminal cell layers expressed CD38 (81± 3%, n = 1486 wall cells), indicating that MCs turned into proinflammatory Mϕs in SH grafts. Interestingly, the lumen of SHV and SH grafts expressed the MC/endothelial marker CD31 (green) but were statistically different (P < 0.05, n = 1668 cells) in the expression of CD14 or CD16, as discussed above. All CD14+ and CD16+ cells costained for CD31 in both SHV and SH grafts (Supplemental Fig. S1).

SHV grafts develop functional luminal endothelium

Next, we examined whether the luminal cells displayed EC function by staining for the active, phosphorylated form of eNOS (green). In SHV grafts, active eNOS was expressed only by luminal cells, which also expressed CD31 (100%, n = 153 luminal cells) (Fig. 7A, B) and CD16 (100%, n = 151 luminal cells) (Fig. 7C). However, in SH grafts, which lacked CD16+ cells (5 ± 2%, n = 1467 total cells), eNOS was observed in subluminal cell layers, suggesting atypical tissue organization (Fig. 7A, C). As expected, in native arteries only, luminal EC coexpressed active eNOS and CD31 (Fig. 7B) but not CD16 (Fig. 7D).

Figure 7.

Figure 7

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Development of function endothelium in A-TEV grafts. A, B) Immunostaining for eNOS (green) and CD31 (red) in SHV and SH grafts (A) or native arteries (B). C, D). Immunostaining for eNOS (green) and MC marker CD16 (red) in SHV and SH grafts (C) or native arteries (D). A, C) The arrow indicates 3 images from the proximal to the distal end of SHV and SH grafts. E) Image quantification and analysis for n = 45 images per antibody panel. Percent positive cells of DAPI positive cells per specified vessel area. L, graft lumen. Scale bars: 10 μm (white), 30 μm (red). *P < 0.05.

DISCUSSION

In this study, we utilized the clinical grade biomaterial known as decellularized porcine SIS, which has been used extensively in the past (24, 37–42). Using SIS with immobilized heparin and VEGF, we developed cell-free off-the-shelf vascular grafts with submillimeter inner diameters (850–900 µm) that were implanted into the arterial system of a mouse. Similar to our previous study with an ovine animal model, grafts demonstrated host cell integration and a confluent endothelium within the lumen (34, 36). Cell-free vascular grafts of such small sizes are a continuing challenge for the field of tissue engineering. Prior to this study, a polymer-based graft composed of an inner core of poly(glycerol sebacate) and a nondegrading outer sheath of thin poly(ε-caprolactone) implanted in mice was patent over a 12-mo period. However, the polymeric outer sheath did not degrade during that time, indicating a lack of integration and long-term remodeling (19).

The anticoagulant heparin has also been used to functionalize the lumen of vascular grafts, with mixed results. One study implanted a heparin-functionalized elastomer in a rat model that proved to be effective in promoting endothelialization (21). However, heparin-functionalized grafts occluded within the first few hours after implantation into the carotid artery of sheep (34, 36). On the other hand, in the current study we found no difference in graft patency between heparin-only (SH) and heparin/VEGF (SHV) grafts using the same biomaterial and immobilization strategy, despite the much smaller graft diameter. Although the reasons are not well understood, significant differences in wall shear stress between the mouse descending aorta (40–50 dyn/cm2) and sheep carotid arteries (10–15 dyn/cm2) may provide a plausible explanation for the patency differences between the 2 studies. Furthermore, EC ingrowth from the anastomotic sites differs between the 2 animal models, with ingrowth being well documented in mice but lacking in sheep and humans (43).

Grafts with immobilized VEGF showed significantly improved tissue organization at 1 mo postimplantation, as evidenced by the presence of a functional cell monolayer on the lumen expressing key endothelial markers (VEGFR2 and CD144) and a medial layer consisting of α-SMA+ cells. In contrast, SH grafts contained multiple layers of disorganized endothelial-like cells with no clear demarcations between the lumen and medial layers (VEGFR1, VEGFR2, and CD144 were seen throughout multiple cell layers) at 1 mo postimplantation. In addition, medial cells were stained for the quintessential smooth muscle cell marker α-SMA and appeared to align perpendicularly to the luminal layer, indicating circumferential alignment. In addition, the thicker cellular component of SHV grafts indicates a more robust cellular infiltration, likely due to the presence of VEGF, a well-known mitogen for ECs and strong chemoattractant for MCs and Mϕs (44, 45). Recruited MCs and Mϕs might have induced robust proliferation of medial-layer cells (especially smooth muscle cells and fibroblasts) in a well-orchestrated series of events typical of the wound healing process (15, 46–48). Increased cell infiltration was accompanied by reduced thickness of the SIS matrix remaining in the grafts at 1 mo postimplantation, suggesting that SHV grafts remodeled more efficiently than SH grafts. It is well documented that the initial phase of rapid cellular proliferation within the vessel wall is followed by decreased proliferation and wall thickness toward more of a homeostatic state (47). Longer implantation times are required to study the long-term remodeling of VEGF-based A-TEVs in this animal model.

Notably, the cellular makeup was significantly different between the SHV and SH grafts. SHV grafts contained significantly more cells expressing EGR2, a marker shared between the anti-inflammatory M2-Mϕs and ECs, and few, if any, cells expressing the proinflammatory M1-Mϕ marker CD38. Cells in the lumen of SHV grafts also expressed CD14 and the M2-Mϕ marker CD16. In contrast, SH grafts contained a large number of cells expressing the M1–proinflammatory marker CD38, significantly fewer cells expressing EGR2, and very few, if any, cells expressing CD16. Collectively, these results indicate that the SIS-based grafts were infiltrated by MCs that turned into M2-Mϕs in the presence of VEGF but M1-Mϕs in grafts containing heparin alone. These results also suggest that immobilized VEGF might have directed the incoming MCs toward an anti-inflammatory phenotype.

MCs, a key component of the inflammatory response, express VEGFR1 and respond to VEGF in a chemotactic fashion (44, 45, 49–61). In addition, endothelial progenitor cells make up <0.1% (62), whereas MCs make up between 10 and 20% (63), of mononuclear cells of the blood; therefore, they may be more likely to be captured by VEGF on the graft lumen via the VEGF receptor (VEGFR1) on their surface. Therefore, initially our grafts might be populated mostly by MCs, which may polarize into M2-Mϕs in the presence of VEGF but M1-Mϕs in the presence of heparin alone (64). M2-Mϕs may be aiding EC ingrowth through paracrine action, as they are known to secrete a multitude of growth factors, including fibroblast growth factor, platelet-derived growth factor, TGF-β, stromal cell-derived factor 1–α, and VEGF (65–68). They also establish new ECM, predominantly fibronectin, that serves as substrate for EC attachment and migration (27, 69). MCs and Mϕs are indispensable for vascular graft patency and remodeling, providing key signals for circulating endothelial progenitor cells and ingrowth of anastomotic ECs, as shown in previous studies (15, 46–48, 70–72). In a key study, Hibino et al. (70) found that Mϕ infiltration was crucial for neotissue formation. When Mϕs were depleted prior to implantation, the vessels lacked both the endothelium and medial layers. Furthermore, they noted that when the grafts were preseeded with bone marrow mononuclear cells, which are known to produce VEGF, among other growth factors and cytokines, the Mϕ phenotype was predominantly M2, as opposed to cell-free naked grafts in which the majority of infiltrating Mϕs turned into M1 phenotype. Taken together, we propose that immobilized VEGF may enable the capture of MCs from the blood stream and direct their conversion to an M2-Mϕ phenotype, which may ultimately affect cell fate and tissue regeneration.

Alternatively, VEGF may be coaxing MCs and Mϕs into differentiating to an EC fate. There is a growing body of literature suggesting that MCs may be capable of differentiating into EC-like cells that express both MC/Mϕ markers and EC markers in vitro (56, 57, 73–82). In agreement, our results revealed that the lumen of SHV grafts contained cells that expressed both M2-Mϕ and EC markers, but in subluminal areas they expressed only Mϕ markers. More experiments are required to address this intriguing hypothesis, and these are being pursued in our laboratories using microfluidic technology and transgenic mouse models. However, regardless of the underlying mechanism, our current study suggests that VEGF may play a crucial role in the regeneration and remodeling of cell-free vascular grafts by modulating the inflammatory response to generate a proregenerative environment.

CONCLUSIONS

In this study, we demonstrated that cell-free, tissue-engineered vascular grafts made of heparin/VEGF-functionalized SIS maintained patency and promoted regeneration of a functional endothelium at 1 mo postimplantation in a mouse model. Heparin and VEGF grafts exhibited improved remodeling and endothelialization compared with heparin-only grafts. We also observed that the graft lumen was populated by cells of dual phenotype expressing both MC/Mϕ and endothelial proteins, indicating a more direct role of immune cells in the regeneration of cell-free vascular grafts. Our results suggest that immobilized VEGF may promote regeneration of cell-free vascular grafts by modulating the inflammatory response of the host.

Supplementary Material

This article includes supplemental data. Please visit http://www.fasebj.org to obtain this information.

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ACKNOWLEDGMENTS

This work was supported by grants from the U.S. National Institutes of Health, National Heart, Lung, and Blood Institute to S.T.A. (R01 HL086582) and to R.J.S. (F31 HL134323 Fellowship). The authors declare no conflicts of interest.

Glossary

A-TEV

acellular tissue–engineered vessel

EC

endothelial cell

ECM

extracellular matrix

Mϕ

macrophage

MC

monocyte

SH

small intestinal submucosa + heparin group

SHV

small intestinal submucosa + heparin + VEGF group

SIS

small intestinal submucosa

Footnotes

This article includes supplemental data. Please visit http://www.fasebj.org to obtain this information.

AUTHOR CONTRIBUTIONS

R. J. Smith, Jr., performed all data analysis and figure creation; R. J. Smith, Jr., T. Yi, and B. Nasiri performed research work from graft design and production to implantation and explantation; R. J. Smith, Jr., C. K. Breuer, and S. T. Andreadis designed the research and overall study; R. J. Smith, Jr., and S. T. Andreadis wrote the manuscript; and T. Yi and C. K. Breuer edited the manuscript.

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