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. 2026 Apr 15;136(9):3895–3906. doi: 10.1002/lary.70564

Endothelial Hydrogels Improve Microvascular Regeneration and Perfusion in Tracheal Scaffolds

Molly O Hunter 1, Lumei Liu 2,3, Nathaniel Hochuli 1, Olivia Chan 1, Jazmin Calyeca 2,3, Ada C Sher 1, Tendy Chiang 1,2,3,4,✉
PMCID: PMC13460639  PMID: 41987006

ABSTRACT

Objective

Vascularization remains a major barrier to the clinical translation of airway replacement. We previously demonstrated that engineered tracheal scaffolds support neovascularization, although restoration of microvascular perfusion is delayed in part due to a reduction in capillary branching. Endothelial hydrogels (EH) have demonstrated the potential to enhance vascular regeneration in several domains of tissue engineering, yet their application in airway replacement has not been fully evaluated. In this study, we assessed the impact of EH on microvascular regeneration and perfusion in tracheal scaffolds.

Methods

Tracheal scaffolds were fabricated from C57BL/6J mice. EH were formulated with endothelial cells (EC) from human or mouse lineages and collagen, then applied to tracheal scaffolds. EH‐scaffolds were cultured ex vivo for 7 days and stained with CD31, DAPI, lectin‐FITC, and vascular endothelial growth factor (VEGF). The optimal EH composition was evaluated in vivo using orthotopic tracheal transplantation. At 1‐month, microvascular regeneration was quantified, patterning was assessed via Sholl analysis, and perfusion was quantified using lectin‐FITC.

Results

EH supported EC attachment on tracheal scaffolds. The ideal EH composition for maximal EC coverage was 3.6 × 106 cells/cm2 and 1.25 mg/mL collagen. EC expressed VEGF and formed microvascular networks on tracheal scaffold ex vivo. EH‐scaffolds were successfully implanted without signs of respiratory distress or obstruction. Compared to control, EH‐scaffolds improved graft perfusion and microvascular patterning in tissue engineered trachea.

Conclusion

EH enhances microvascular regeneration and perfusion of engineered tracheal scaffolds in vivo, thus supporting the potential of EH as a strategy to accelerate vascular regeneration in airway reconstruction.

Level of Evidence

N/A.

Keywords: endothelial hydrogels, hydrogels, tissue engineering, tracheal scaffolds

Insufficient vascularization remains a major barrier to successful tracheal transplantation. This study demonstrates that an endothelial hydrogel significantly enhances graft perfusion and neovessel organization compared to non‐endothelial hydrogel controls. This approach may improve scaffold integration and advance clinical translation of engineered airway constructs.

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1. Introduction

In the airway, replacement for long‐segment tracheal defects remains an unmet clinical need. Tissue‐engineered tracheal grafts offer a promising solution for these defects, however successful revascularization of grafts remains a major hurdle, as both synthetic and decellularized tracheal grafts have exhibited poor endothelial cell regeneration and vascularization in vivo [1, 2, 3]. Various attempts to accelerate revascularization, such as utilizing prelamination with vascularized tissue flaps, have shown limited benefit [4].

We have previously established an engineered tracheal scaffold that can sustain microvascular regeneration equivalent to native trachea 3 months after orthotopic implantation, with a similar degree of perfusion to surgical controls [5, 6, 7]. However, we found a delay in perfusion at early timepoints compared to surgical controls, which is a critical interval for successful graft integration [7]. Additionally, microvascular patterning of neovessels within the tracheal scaffold was observed to exhibit less branching compared to control [7]. Delayed perfusion may stem from altered vascular patterns, as vascular organization has been shown to influence perfusion efficiency, nutrient transport, signal communication, angiogenic sprouting, and oxygenation [8, 9, 10, 11].

Endothelial hydrogels (EH) are created by combining endothelial cells (EC) with an extracellular matrix‐based vehicle [12, 13, 14, 15]. EH have demonstrated benefit in enhancing neovascularization in a variety of animal model organs, including skeletal, cardiac, and peripheral nervous systems [16, 17, 18]. Despite these advancements, the application of EH in airway reconstruction, where prompt restoration of a functional blood supply is critical, remains unexplored. To address this, we aimed to examine the effects of EH on neovascular regeneration, perfusion, and patterning in engineered tracheal scaffolds. To do so, we created an EH incorporating mouse lung endothelial cells (MLEC) and collagen and assessed the impact of the optimal EH composition in a mouse model of airway transplantation.

2. Materials and Methods

2.1. Animal Care and Ethics

The Institutional Animal Care and Use Committee of the Abigail Wexner Research Institute at Nationwide Children's Hospital reviewed and approved the study protocols (AR15‐00090, AR21‐00021). All animals were cared for in accordance with the Guide for the Care and Use of Laboratory Animals (8th edition, 2011; National Institutes of Health, Bethesda, MD), the Public Health Service Policy on Humane Care and Use of Laboratory Animals, and the regulations of the U.S. Department of Agriculture (USDA) outlined in the Animal Welfare Act.

2.2. Tracheal Scaffold Fabrication

Tracheal segments (4–8 rings) were harvested from female C57BL/6J mice, spanning tracheal rings 3–10, and decellularized to create a tracheal scaffold using our previously published sodium dodecyl sulfate–based protocol [19]. Tracheal scaffolds were cryopreserved in Dulbecco's Modified Eagle Medium (DMEM, Gibco) with 10% fetal bovine serum, 1% penicillin/streptomycin, and 5% dimethyl sulfoxide (DMSO, Fisher Scientific) at −80 °C until ready for use.

2.3. HUVEC‐Derived EH (hEH) Preparation and Loading to Tracheal Scaffold

Human umbilical vein endothelial cells (HUVEC) were expanded in complete endothelial growth medium (EGM‐2 Endothelial Cell Growth Medium‐2 BulletKit, Lonza, Walkersville, MD). HUVEC‐derived EH (hEH) were seeded on open‐booked tracheal scaffolds (4 rings) with the submucosa facing up. Collagen (5 mg/mL, Cultrex Rat Collagen I R&D Systems, Minneapolis, MN) was diluted to 1.25 mg/mL and 4 mg/mL with 10 × PBS and ddH2O and pH was adjusted to 7 with NaOH. HUVEC (0, 1.7 × 103, 1.7 × 104, 1.7 × 105, 8.5 × 105) were mixed with 1.25 mg/mL and 4 mg/mL collagen (n = 4/group). Collagen concentrations were selected based on previous publications for HUVEC [20] and other endothelial cell types [21]. One μL of hEH was loaded on tracheal scaffolds yielding a density of approximately 0, 3.6 × 104, 3.6 × 105, 3.6 × 106, 1.8 × 107 cells/cm2. hmEH‐scaffolds were incubated for 15 min (37°C, 5% CO2). Once polymerized, hEH‐scaffolds were cultured at standard conditions (37 °C, 5% CO2) in complete endothelial growth medium for 7 days.

2.4. MLEC‐Derived EH (mEH) Preparation and Loading to Tracheal Scaffold Lumen

RFP‐expressing C57BL/6 MLEC were purchased from Cell Biologics (Chicago, IL). MLEC were cultured and expanded in T25 flasks coated with a gelatin‐based coating solution (Cell Biologics) supplied with complete endothelial cell medium (M1168‐Kit, Cell Biologics). Based on prior optimization experiments using hEH, the highest tested concentration contained minimal collagen, barely sufficient to suspend endothelial cells, whereas the lowest concentration resulted in inadequate cell density. Consequently, two intermediate concentrations, representing feasible ranges, were selected for evaluation with mEHs. mEH were composed of mixing MLEC (0, 1.7 × 105 and 1.7 × 106) with 1.25 mg/mL collagen. Tubular tracheal scaffolds (5 rings) were seeded with 1.5 μL of mEH yielding approximate densities of 0, 3.6 × 105, 3.6 × 106 cells/cm2 (n = 4/group) and mEH‐tracheal scaffolds were incubated at 37 °C for 20 min. mEH‐tracheal scaffolds were maintained at standard culture conditions (37 °C, 5% CO2) in complete endothelial culture medium for 7 days.

2.5. Hydrogel Staining and in Vitro Preparation

On day 7, EH‐scaffolds were rinsed with PBS and fixed in 4% PFA for 2 h at 4°C. EH‐scaffolds were then washed with PBS 3 times (10 min/wash). Lectin‐FITC staining was used as an angiogenesis assay to visualize functional endothelial networks. Lectin‐FITC from Lycopersicon esculentum for mEH‐scaffolds and Lectin‐FITC from Ulex europaeus agglutinin for hEH‐scaffolds (Vector Laboratories, Plain City, OH) were diluted to 100 μg/mL, and 200 μL lectin‐FITC dilution was added to each EH‐scaffold for 30 min. After washing with 1X PBS 3 times (10 min/wash), the EH‐scaffolds were blocked in 20% donkey serum (Sigma‐Aldrich) and 0.3% Triton X (ThermoFisher Scientific) in PBS at 4°C overnight. Rat anti‐CD31 (primary antibody, 1:100, Abcam, Waltham, MA) and rabbit anti‐VEGFA (primary antibody, 1:200, Abcam) in 1% donkey serum in PBS were added to EH‐scaffold and incubated for 24 h at 4°C. After washing with 1X PBS 3 times (20 min/wash), donkey anti‐rat Alex Fluor 594 and donkey anti‐rabbit (secondary antibodies, 1:200, Invitrogen, Waltham, MA) were added and incubated for 24 h at 4°C. The EH‐scaffolds were stained with 1 μg/mL 4,6‐diamidino‐2‐phenylindole (DAPI, Invitrogen) for 1 h at room temperature, then washed with PBS 3 times (20 min/wash). EH‐scaffolds were first imaged with confocal microscope axially (AX R confocal system, Nikon, Melville, NY). Then EH‐scaffold was cut open on the anterior wall and mounted on coverslips with the submucosa facing down. After adding a drop of antifade mounting medium (VECTASHIELD PLUS, Vector lab), glass slides were placed on the flat EH‐scaffold. Whole mount EH‐scaffolds were imaged with confocal microscope (AX R confocal system, Nikon, Melville, NY). EC coverage on EH‐scaffolds was quantified by lectin‐FITC area over the whole‐mount tissue area. The optimal composition of EH was seeded on trachea scaffolds for 7 days and characterized with Masson's Trichrome staining for histological observation.

2.6. EH‐Tracheal Scaffold Implantation, Whole Mount Harvest and Staining

mEH were loaded onto the lumen of tracheal scaffolds (8 rings) at cell densities 0 and 3.6 × 106/cm2 (n = 5/group). mEH was also seeded in a 16‐well plate (Nunc Lab‐Tek Chamber Slide System) to validate cell function (Figure S1). On day 7, mEH‐scaffolds were trimmed to a 5‐mm segment for orthotopic implantation into a female C57BL/6J mouse using previously published implantation methods [22]. The remaining trimmed tracheal segments were fixed in 4% PFA for mEH characterization (Figure S1). At 1 month, implanted mEH‐scaffolds were harvested following our previously published perfusion protocol [7]. Briefly, FITC‐conjugated tomato lectin (100 μL of 1 mg/mL Vector Laboratories) was injected into the left ventricle with a 30‐gauge ultra‐fine insulin syringe (BD, Franklin Lakes, NJ) for 1 min. Then 20 mL 4% paraformaldehyde in PBS (ThermoFisher Scientific) was perfused via the left ventricle with a 25‐gauge needle. mEH‐scaffolds along with anastomosed host trachea were isolated. Harvested mEH‐scaffolds were stained and mounted according to previously published methods [7]. Primary antibodies were rat anti‐CD31 (1:100, Abam, Waltham, MA) and rabbit anti‐RFP (1:50, Invitrogen, Carlsbad, CA). The secondary antibodies were donkey anti‐rat Alex Fluor 594 (1:200, Invitrogen) and donkey anti‐rabbit Alex Fluor 647 (1:200, Invitrogen). Whole mount tracheal scaffolds were imaged using a confocal microscope.

2.7. Cell Seeding Efficiency and Microvascular Quantification

Cell seeding efficiency was quantified using the percent of FITC+ area over total tracheal scaffold area using ImageJ software (National Institutes of Health, Bethesda, MD). Microvascular areas (CD31 and lectin‐FITC) were quantified using percentage of CD31+ area over high power field image area and FITC+ area over total graft area. Samples were excluded from lectin‐FITC quantification if the following applied: (1) Perfusion failure: animals not successfully perfused with lectin‐FITC in the proximal and distal host regions; (2) Cell seeding failure: distal region of implanted grafts lacked evidence of successful seeding (Figure S1A). Samples lacking evidence of successful seeding (Figure S1A) were also excluded from CD31 quantification. Microvascular patterning was quantified using Sholl analysis following previously published methods [7]. All analyses were conducted in a blind manner.

2.8. Statistical Analysis

Statistical analysis was performed using GraphPad Prism 10 software (GraphPad Software LLC, CA). Ordinary one‐way ANOVA was used to compare multiple groups (Tukey's multiple comparisons test). Microvascular patterns analyzed using Sholl were compared using paired t‐tests. Statistical difference was defined as p < 0.05. Experimental data were expressed as mean ± standard deviation (SD).

3. Results

3.1. hEH Delivered and Retained HUVEC on Tracheal Scaffold

After 1 week ex vivo, we found that hEH had successfully retained HUVEC on tracheal scaffolds as evidenced by CD31+ and Lectin‐FITC+ endothelial cells remaining on the surface (Figure 1). The density of retained HUVEC on the scaffolds increased with cell seeding density, with dose saturation observed at 3.6 × 106 cells/cm2 (Figure 1A–D).

FIGURE 1.

FIGURE 1

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Confocal imaging of HUVEC on tracheal scaffolds. Confocal images of hEH‐scaffolds using 1.25 mg/mL collagen and varying cell concentrations with high powered images are shown. Representative images of collagen only control (A), 3.6 × 104 cells/cm2 hEH (B), 3.6 × 105 cells/cm2 hEH (C), 3.6 × 106 cells/cm2 hEH (D), 1.8 × 107 cells/cm2 hEH (E) are shown on tracheal scaffolds. (Green: Lectin‐FITC; Blue: DAPI‐stained nuclei; Red: CD31).

Additionally, we evaluated the impact of collagen concentration (1.25 mg/mL and 4 mg/mL) on hEH performance. Both collagen concentrations supported successful seeding of hEH on tracheal scaffolds and showed no difference in seeding efficiency (Figure S1). Due to the high viscosity of the 4 mg/mL concentration, 1.25 mg/mL collagen was chosen for use in the subsequent experiments.

3.2. hEH Delivered and Retained Functional Endothelial Cells on Tracheal Scaffold Basement Membrane

To determine the optimal cell concentration for hEH, we quantified cell coverage on the tracheal scaffolds (Figure 2A). hEH with a concentration of 3.6 × 106 cells/cm2 exhibited the highest cell coverage (46.6% ± 13.9%.), supporting the observations in Figure 1. The lowest cell concentration (3.6 × 104 cells/cm2) exhibited minimal coverage (1.04% ± 1.04%), and was significantly lower than higher concentrations (3.6 × 106 and 1.8 × 107 cells/cm2; p = 0.0002, p = 0.0052). At 3.6 × 105 cells/cm2, coverage was 15.11% ± 7.22%, lower than 3.6 × 106 cells/cm2 (p = 0.0029). The highest cell concentration (1.8 × 107 cells/cm2) reached 29.8% ± 8.64%, which was not different from 3.6 × 106 cells/cm2 (p = 0.13).

FIGURE 2.

FIGURE 2

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hEH characterization on tracheal scaffold. A. EC coverage of 3.6 × 104 cells/cm2, 3.6 × 105 3.6 × 106 cells/cm2 and 1.8 × 107 cells/cm2 on tracheal scaffold surface area. B. VEGF characterization. a. A representative image of VEGF (appearing in white) on a collagen only control; b. A representative image of VEGF (appearing in white) on hEH‐scaffold with ideal EC concentration (3.6 × 106 cells/cm2). C. Representative Masson's trichrome images of no collagen control (a) and hEH on basement membrane of tracheal scaffold (b, c). Yellow arrowheads denote basement membrane and green arrowheads denote HUVEC. * denotes higher HUVEC coverage of 3.6 × 106 and 1.8 × 107 cells/cm2 groups than 3.6 × 104 cells/cm2 group (*** p = 0.0002, ** p = 0.0052); ** denotes 3.6 × 106 cells/cm2 group had higher HUVEC coverage than 3.6 × 104 cells/cm2 (p = 0.0029).

After identifying 3.6 × 106 cells/cm2 as the ideal cell concentration for hEH, we confirmed that HUVEC remained functional in hEH as evidenced by their expression of vascular endothelial growth factor (VEGF) on the scaffold (Figure 2B). Additionally, we demonstrated that the hydrogel itself did not cause obstruction of the tracheal scaffold lumen, as there was no collagen in the lumen (Figure 2C). Furthermore, we identified that endothelial cells were directly attached to the basement membrane (Figure 2C).

3.3. mEH Successfully Delivered Functional MLEC to Tracheal Scaffold

After 7 days in vitro, EC formed a FITC‐lectin+ network suggesting mEH preserved the function of MLEC (Figure S2). Similar to in vitro, FITC‐lectin demonstrated a robust network formation by endothelial cells in mEH with selected cell concentrations (3.6 × 105 cells/cm2 and 3.6 × 106 cells/cm2) as denoted by the yellow arrowheads in Figure 3B,C. EC extended and interconnected to form capillary‐like tubular structures with multiple branch points and junctions resembling microvascular networks. The observed morphology was suggestive of in vitro angiogenesis on tracheal scaffold, another proxy for the preservation of EC function (Figure 3). Furthermore, mEH supported EC on top of the basement membrane, as shown in Figure 4.

FIGURE 3.

FIGURE 3

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Characterization of mEH‐tracheal scaffolds. Whole mount images of collagen only (A), mEH 3.6 × 105 cells/cm2 (B), and mEH 3.6 × 106 cells/cm2 on tracheal scaffold (C). Yellow arrowheads denote angiogenesis of mEH on tracheal scaffolds.

FIGURE 4.

FIGURE 4

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mEH on tracheal scaffolds. A. MLEC coverage on tracheal scaffold surface. B. Representative Masson's trichrome images of collagen only and mEH on basement membrane of tracheal scaffold. Yellow arrowheads denote basement membrane and green arrowheads denote MLEC. **** denotes higher MLEC coverage of 3.6 × 105 cells/cm2 and 3.6 × 106 cells/cm2 groups than collagen only group (p < 0.0001), and higher MLEC coverage of 3.6 × 106 cells/cm2 than 3.6 × 105 cells/cm2 (p < 0.0001).

The EC coverage of 3.6 × 106 cells/cm2 on tracheal scaffold was higher than 3.6 × 105 cells/cm2 (p < 0.0001), and both were higher than no cell control (p < 0.0001). We selected the EC concentration with the highest EC coverage for the ex vivo characterization and in vivo implantation. EC (green arrowheads) were distributed along the basement membrane, with only a few EC observed in the submucosa near the graft ends.

3.4. mEH Accelerated Neovascularization and Improved Microvascular Branching

RPF‐labeled endothelial cells were proven functional prior to implantation (Figure S3, Figure 3). Ten of the eleven mice in the mEH group and all animals of the collagen only group (n = 9) survived to the planned end point, showing no signs of respiratory distress or weight loss (Figure S4A). All grafts were patent upon tissue harvest (Figure S4B). However, at 1‐month post‐implantation, no RPF+ EC remained on tracheal scaffolds (Figure S5).

mEH enhanced neovascularization on the tracheal scaffolds (Figure 5B), similar to the collagen only group (Figure 5A), although the neovasculature appeared morphologically different from the host control (Figure 5C). Quantification showed that microvascular area in the mEH group was comparable to host control (p = 0.0872), whereas the collagen only group had less neovascular area than host control (p = 0.0424) (Figure 5D). Both collagen only and mEH groups demonstrated increased microvascular area compared to our previous scaffold‐only data (p = 0.0228, 0.025).

FIGURE 5.

FIGURE 5

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Neovascularization whole‐mount characterization. Representative images of whole mount and high‐power field image of collagen only on tracheal scaffold (A), mEH on tracheal scaffold (B), and host control (C). (D) Quantification of CD31+ area of high‐power field image. # denotes higher neovascular area of collagen only and mEH groups than scaffold only control (p = 0.0028, 0.0250); * denotes lower CD31+ area in collagen only group than host control (p = 0.0424).

3.5. mEH Improved Microvascular Pattern

We measured microvascular pattern using Sholl analysis (Figure 6A). Compared to collagen only, EH improved microvascular pattern at the regions of trachealis (p < 0.001), intercartilaginous area (p < 0.0001), cartilage (p < 0.0001), and proximal (p < 0.0001) and distal anastomoses (p < 0.0001; Figure 6B).

FIGURE 6.

FIGURE 6

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Microvascular pattern analysis. A. Representative images of microvascular pattern of collagen only group (a) and mEH group (b). B. Microvascular pattern (intersection number) of collagen only and mEH groups at trachealis (a), intercartilaginous area (b), cartilage (c), proximal anastomosis (d), and distal anastomosis (e). * denotes mEH improved microvascular pattern (p < 0.001).

3.6. mEH Promoted Microvascular Perfusion in Tracheal Scaffolds

Whole‐mount lectin‐FITC imaging revealed robust perfusion in graft regions and distal host control (Figure 7A–C), indicating successful tracheal perfusion. In contrast to the similar findings regarding microvascular density, mEH improved perfusion in tracheal scaffolds compared to collagen only (p = 0.0006). Both collagen only and mEH improved perfusion compared to previous scaffold only data (p = 0.0002 and p < 0.0001). Collagen‐only scaffolds showed reduced perfusion when compared to the adjacent host airway (p = 0.0035), while mEH treated scaffolds demonstrated the same perfusion quantitatively when compared with the host (p = 0.680).

FIGURE 7.

FIGURE 7

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Whole‐mount perfusion analysis. Representative images of whole‐mount perfusion of tracheal scaffold with collagen only (A) and mEH (B) host control (C). (D) Quantification of perfusion in graft using FITC+ area. # denotes improved perfusion of collagen only and mEH, compared to scaffold only control *p = 0.0002, < 0.0001; red * denotes lower perfusion of collagen only group than host control (p = 0.0035); black * denotes increased perfusion of mEH than collagen only group (p = 0.0006).

4. Discussion

Rapid restoration of tracheal blood supply is critical for the survival and integration of tracheal grafts; however, it remains a major hurdle in clinical translation [1, 2, 3, 4, 23]. Several characteristics of the trachea pose inherent challenges to transplantation. First, the native blood supply limits options for microvascular free tissue transfer, often resulting in graft devascularization and increasing the risk of infection, necrosis, and stenosis [5, 24, 25]. Second, grafts must support a functional respiratory epithelium, as delayed epithelialization disrupts mucociliary clearance and increases inflammation and infection risk [5, 24]. Third, maintaining the mechanical properties of the cartilaginous framework is required for proper respiratory function [26]. Failure to retain native mechanics has resulted in graft collapse (malacia) and narrowing (stenosis) [27, 28, 29]. Finally, the airway is a non‐sterile environment exposed to irritants, debris, and pathogens, which can impair graft regeneration through infection and inflammation [26]. Therefore, graft integration must be robust enough to withstand these conditions.

To address these challenges, our lab developed a tracheal scaffold that is non‐immunogenic, supports host‐derived neotissue formation, regenerates all native tracheal cell types, and remains patent in vivo [6, 19, 22, 30]. We further demonstrated that this scaffold can be combined with biomaterials to maintain mechanical properties comparable to synthetic constructs [30, 31, 32]. Our scaffold supports microvascular regeneration and perfusion equivalent to native trachea and surgical controls [5, 6]. However, restoration of perfusion can take months, which is untenable for clinical applications [7]. Reduced perfusion is associated with disorganized neovascular networks, potentially leading to inadequate nutrient and oxygen delivery and graft failure [7]. Therefore, we sought to accelerate airway microvascular regeneration using EH, which leverages the biocompatibility of hydrogels and the ability of EC to deliver physiological levels of pro‐angiogenic factors [33, 34, 35].

In this study we found that EH successfully delivered EC of human and mouse origin onto tracheal scaffolds. Although both collagen and EH enhanced microvascular regeneration and perfusion compared to our previous scaffold‐only data [7], EH further improved microvascular regeneration, vascular patterning, and perfusion relative to the collagen only group. The improvements by EH despite the absence of direct incorporation of EC into neovasculature may be attributed to the indirect impacts of the pro‐angiogenic factors released by the seeded EC, such as VEGF and PDGF. Endothelial cells secrete VEGF to promote angiogenesis [36, 37, 38, 39] and PDGF to stabilize newly formed vessels [40].

Direct delivery of pro‐angiogenic factors to tracheal scaffolds poses challenges, including dose variability, reliance on single or dual factor(s) delivery, and drug safety concerns. For example, VEGF has limited clinical application because of VEGF‐induced vascular permeability [41, 42, 43]. Moreover, high VEGF levels are often associated with cancer, and thus no FDA‐approved therapies currently use VEGF to promote angiogenesis. Instead, most FDA‐approved drugs target VEGF inhibition for oncology. In contrast, PDGF is FDA approved for topical treatment of chronic skin wounds such as diabetic foot ulcers [44], though its potential to enhance airway neovascularization has not been explored.

EH addresses these challenges by leveraging the intrinsic capacity of EC to dynamically regulate their secretion of pro‐ and antiangiogenic factors in response to environmental cues. This adaptive signaling is crucial for controlling angiogenesis through a finely tuned interplay of molecular pathways. By supplying tracheal scaffolds with early, physiologically regulated levels of pro‐angiogenic factors regulated from EC, EH may have promoted earlier angiogenic responses than in our previous graft models, leading to improved vascular organization and enhanced microvascular perfusion. Therefore, by supplying the tracheal scaffold with early pro‐angiogenic signals via the EH, this physiological process may have been initiated earlier than our previous no‐EH models, leading to improved vascular organization and thus improved microvascular perfusion.

One barrier of translating EH improvements on vascularization to larger animal models, and eventual clinical translation to humans, is immunogenicity and availability of compatible EC types. The EC would need to be genetically compatible with the host to avoid an immunogenetic response, or an immunosuppressed host would need to be utilized. One possible method of EC collection is the endothelial biopsy, in which a biopsy is taken of tissue from the host, and EC are isolated and expanded [45, 46, 47]. However, this method is invasive and would require that EC are taken from the individual for which they are intended for, with limited replicative potential, making this a suboptimal method [48]. Induced pluripotent stem cells (IPSCs) offer a solution to these challenges, allowing for differentiation into EC with a more robust replicative potential [49]. Obtaining these cells provides a more straightforward approach, as they have been produced from human dermal fibroblasts [50]. Although significant challenges remain in both tracheal‐tissue engineering and the application of induced pluripotent stem cells, their convergence holds substantial promise for advancing EH into clinical translation for human patients.

The study introduces several limitations. Although improved microvascular perfusion was observed following implantation of mEH constructs, the precise mechanism underlying this effect remains unclear. RFP‐labeled endothelial cells were not detected at later time points, suggesting that direct long‐term structural incorporation into host vasculature may be limited. Therefore, paracrine signaling from the transplanted endothelial cells is likely to contribute to the observed vascular improvement. However, specific angiogenic factors (e.g., VEGF or other mediators) were not directly investigated in this study. Further mechanistic studies will be required to identify the dominant signaling pathways involved. Another limitation of this study is that scaffold‐only controls were historical rather than performed concurrently. To minimize variability, the same procedures, methods, and operator were used. Future studies will include simultaneous scaffold‐only controls to more directly assess the specific contribution of EH. Finally, the effect of EH on epithelial cells and chondrocytes was not assessed in this cohort and remains unclear. Future studies are needed to evaluate these aspects of graft regeneration.

5. Conclusion

In this study, we found that EH successfully delivered active and functional endothelial cells of human and mouse onto tracheal scaffolds. EH improved microvascular regeneration, vascular patterning, and perfusion relative to the collagen only group. Future studies should be focused on translation to larger animal models.

Funding

This research was supported by the Medical Student Pelotonia Scholars Program (Molly Hunter is the recipient) as well as by NIH NHLBI R01HL157039 (Tendy Chiang is the recipient).

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Figure S1: Workflow of mEH seeding, culturing, and implantation.

Figure S2: Collagen concentration selection. Representative images of hEH‐scaffolds: A and B show cell densities of 3.6 × 104 cells/cm2 and 3.6 × 105 cells/cm2, respectively, in 4 mg/mL collagen. Corresponding images for the same cell densities in 1.25 mg/mL collagen are shown in Figure 1. C. Quantification of HUVEC coverage on tracheal scaffold surface. *denotes higher HUVEC coverage on hEH‐scaffold with 3.6 × 105 cells/cm2 compared to the no‐cell control and the 3.6 × 104 cells/cm2 condition, at both 1.25 mg/mL and 4 mg/mL collagen concentrations.

Figure S3: mEH supported EC angiogenesis function in standard cell culture condition. Yellow arrowhead denotes angiogenesis.

Figure S4: mEH did not affect survival and graft patency. A. Survival curve of animals; B. A representative patent graft at explantation.

Figure S5: RFP+ cell characterization. Whole‐mount images of collagen only on tracheal scaffold (A) and mEH on tracheal scaffold (B) showed negative RPF staining in both the graft and host, with similar background noise observed in each region.

LARY-136-3895-s001.docx (6.2MB, docx)

Acknowledgments

We would like to thank Dr. Brenda J. Lilly from the division of cardiovascular medicine at Nationwide Children's Hospital for her kind contributions of the human umbilical vein endothelial cells (HUVEC) used in this series of experiments. We would like to express our gratitude to the Cores of Animal Resource, Morphology and Microscopy at Nationwide Children's Hospital.

Hunter M. O., Liu L., Hochuli N., et al., “Endothelial Hydrogels Improve Microvascular Regeneration and Perfusion in Tracheal Scaffolds,” The Laryngoscope 136, no. 9 (2026): 3895–3906, 10.1002/lary.70564.

This article was presented at: American Society of Pediatric Otolaryngology, May 1st, 2025, Montreal, QC, Canada.

Data Availability Statement

The data that support the findings of this study are available on request from the corresponding author. The data are not publicly available due to privacy or ethical restrictions.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Figure S1: Workflow of mEH seeding, culturing, and implantation.

Figure S2: Collagen concentration selection. Representative images of hEH‐scaffolds: A and B show cell densities of 3.6 × 104 cells/cm2 and 3.6 × 105 cells/cm2, respectively, in 4 mg/mL collagen. Corresponding images for the same cell densities in 1.25 mg/mL collagen are shown in Figure 1. C. Quantification of HUVEC coverage on tracheal scaffold surface. *denotes higher HUVEC coverage on hEH‐scaffold with 3.6 × 105 cells/cm2 compared to the no‐cell control and the 3.6 × 104 cells/cm2 condition, at both 1.25 mg/mL and 4 mg/mL collagen concentrations.

Figure S3: mEH supported EC angiogenesis function in standard cell culture condition. Yellow arrowhead denotes angiogenesis.

Figure S4: mEH did not affect survival and graft patency. A. Survival curve of animals; B. A representative patent graft at explantation.

Figure S5: RFP+ cell characterization. Whole‐mount images of collagen only on tracheal scaffold (A) and mEH on tracheal scaffold (B) showed negative RPF staining in both the graft and host, with similar background noise observed in each region.

LARY-136-3895-s001.docx (6.2MB, docx)

Data Availability Statement

The data that support the findings of this study are available on request from the corresponding author. The data are not publicly available due to privacy or ethical restrictions.

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