Tissue engineered vascular grafts transform into autologous neovessels capable of native function and growth

Computational model of G&R

An analysis of neotissue kinetics using a computational G&R model enables long-term predictions of TEVG behaviors. We previously developed and validated a constrained mixture based G&R model that accurately described and predicted native vessel behavior by simulating the evolving mass density of native constituents (nat) at time s according to

where ({m}_{h}^{{{{{{rm{mech}}}}}}}) is the rate of mass density production of the native constituent in the homeostatic state, ({varUpsilon }^{{{{{{rm{mech}}}}}}}(tau )) is a stimulus function at time τ that modulates production according to deviations from the homeostatic values of intramural stress and wall shear stress, and ({q}^{{{{{{rm{mech}}}}}}}(s,tau )in [0,1]) is a survival function that tracks the degradation of a cohort produced at past time τ to current time s^11,41. We subsequently modified this native vessel model to simulate G&R in TEVGs

where ρ^neo(s) is the total evolving mass density of all neotissue constituents, resulting from ρ^infl(s) and ρ^mech(s), that is, constituents produced in response to inflammatory stimuli and mechanical stimuli, respectively¹¹. The model thus built upon our original description of native vessel G&R but with the addition of inflammation-mediated contributions induced by the polymeric scaffold, which we had shown experimentally to be critical for neotissue formation^11,42,43. The current computational G&R model thus highlights the roles of both mechanical and inflammatory stimuli in driving the natural history of the TEVG. The mass density of mechano-mediated constituents was written as

where ({M}^{{{{{{rm{nat}}}}}}}(tau )) describes the native constituent kinetics at each time s from Eq. (3), and an exponential-decay term modifies the time of its influence to account for delayed cellular infiltration. Importantly, the model considers that the scaffold alters G&R via stress-shielding, where decreased intramural stresses from the presence of a stiff polymer can decrease mass density production as in a native vessel subjected to reduced loading⁴⁴. Additionally, the luminal wall shear stress changes as the diameter of the TEVG evolves. With the stenosis observed in early remodeling, wall shear stress increased and intramural stress decreased, both acting as mechanobiological stimuli to decrease mass density production via the mechano-mediated stimulus function ({varUpsilon }^{{{{{{rm{mech}}}}}}}(tau ) , < , 0) in the early remodeling phase. Inflammatory effects from the foreign body response generated a new class of constituents whose mass density were modeled as

where ({m}_{h}^{{{{{{rm{infl}}}}}}}) is a basal rate of inflammatory production, ({varUpsilon }^{{{{{{rm{infl}}}}}}}(tau )) is a gamma function accounting for the transient production of immuno-mediated neotissue until polymer degradation is complete, and ({q}^{{{{{{rm{infl}}}}}}}(s,tau )in [0,1]) is the survival function for the cohort produced at past time τ that persists at current time s. With the assumption that complete polymer degradation leads to eventual resolution of the immune response, we can write ({rho }^{{{{{{rm{infl}}}}}}}(sto infty )=0). Then, since ({rho }^{{{{{{rm{mech}}}}}}}(sto infty )={rho }^{{{{{{rm{nat}}}}}}}(s)) with the exponential decay term approaching zero,

Thus, the behavior of the TEVG can be separated into two periods: the first, hereafter referred to as the period of neotissue formation, during which G&R are mainly influenced by the immune response to the scaffold material and, the second, referred to as the period of neovessel remodeling, which occurs after scaffold degradation when the mechano-mediated stimulus ({varUpsilon }^{{{{{{rm{mech}}}}}}}(tau ), > ,0) due to increasing intramural stress and decreasing wall shear stress with decreasing thickness and increasing diameter. Consequently, a key result of the model is the prediction that neovessel G&R should eventually mimic the purely mechano-mediated G&R found in native vessels. Further description of the G&R model can be found in the “Methods”.

Natural history of neovessel formation

Scaffolds were fabricated from polyglycolic acid (PGA) fibers that were knitted into a tube and coated with a 50:50 copolymer of polycaprolactone and polylactide (PCLA), which together formed a porous sponge (Fig. 1A, B). The scaffold was designed to degrade by hydrolysis. Our previous studies demonstrated that upon implantation, the scaffold initially functioned as a synthetic vascular conduit; soon thereafter, neotissue formed as the scaffold degraded. Neotissue formation, as characterized by angiography and histology, was highly dynamic over the first 26 weeks after implantation, with marked graft narrowing during the first six weeks followed by spontaneous reversal by 26 weeks¹¹. Beyond the initial 26-week period, morphological changes were more gradual (Fig. 1C), similar to native vessel G&R under nearly constant hemodynamics. Once the scaffold degraded completely, by 52 weeks, the resulting neovessel grossly resembled a native blood vessel (Fig. 1D).

Mechanisms underlying G&R

Serial intravascular ultrasound (IVUS) imaging over the 52-week study period (1, 6, 26, and 52 weeks post-implantation) provided additional information on the morphology of the evolving TEVG (Fig. 2A). Changes in the lumen of a blood vessel can arise from (i) thickening or thinning of the wall (intramural growth defined here as an increase in thickness of the graft wall), (ii) inward or outward remodeling (inward remodeling defined here as an inward change in the outer wall of the graft), and (iii) combinations of the two (Fig. 2B). Serial IVUS data comparing wall thickness to luminal diameter demonstrated that the TEVG lumen narrowed between 1 and 6 weeks primarily due to wall thickening through intramural growth. IVUS imaging between 6 and 26 weeks revealed wall thinning in addition to inward remodeling, namely, a decrease in outer diameter without a decrease in inner diameter. Between 26 and 52 weeks, the wall continued to thin and the lumen expanded (Fig. 2C).

Histological evaluation of the explanted TEVGs demonstrated that between 1 and 6 weeks after implantation, the luminal narrowing causing TEVG stenosis was primarily due to thickening of the scaffold wall. H&E staining demonstrated that this thickening arose from the infiltration and proliferation of cells, resulting in TEVG wall area at 6 weeks being 371 ± 66% of the implanted scaffold wall area by IVUS. There was also evidence of appositional growth of vascular neotissue along the luminal surface of the scaffold, though it accounted for only 16 ± 5% of the total wall area at 6 weeks. The TEVG stenosis spontaneously reversed between 6 and 26 weeks as the wall thinned (IVUS wall thickness 6.5 ± 1.2 mm 6 week vs 2.7 ± 0.5 mm 26 week). This thinning arose from degradation of the scaffold and loss of associated neotissue, yet the lumen did not fully return to its pre-implant area due to persistent inward remodeling. The wall thinned further between 26 and 52 weeks with the lumen enlarging as the scaffold and its associated neotissue waned (Fig. 2D).

Interestingly, the correlation between TEVG lumen area and wall thickness as measured by IVUS changed throughout the time course (Supplemental Fig. 1). At one week, there was no correlation between the two, as the graft parameters were primarily defined by the scaffold. At six weeks, the point of highest measured graft wall thickness, there was a negative relationship between IVUS lumen area and wall thickness, with thicker walls leading to more stenosis. After 26 weeks post-implantation, there again was no relationship; however, after 52 weeks, the relationship was positive, with larger lumens having thicker walls.

Role of the scaffold on G&R (in vitro degradation study)

Based on the computational G&R model prediction of the dual roles played by the scaffold, we characterized both its evolving material properties and diminishing mass, which provided stress shielding and inflammatory stimuli, respectively. The accelerated scaffold degradation study utilized the temperature-dependent reaction kinetics of hydrolytic degradation of the PGA/PCLA scaffolds by bathing samples for 0, 1, 3, 5, 7, 9, and 14 days in PBS heated to 70 °C in vitro (Fig. 3A–C). 70 °C was chosen as preliminary studies revealed that one day of accelerated degradation corresponded to approximately one month of real-time degradation at 37 °C (Supplemental Fig. 2). Quantitative analyses revealed complete degradation over 14 days in vitro, with differential rates of degradation for PGA (by day 5 in vitro, estimated month 5 in vivo) and PCLA (by day 14 in vitro, estimated month 14 in vivo) (Fig. 3D). The TEVG was constructed with 25% of the initial mass being the knit central PGA layer, with the remaining 75% of the initial mass representing the PCLA sponge layers. Morphometric characterization via SEM demonstrated that pore size initially increased and then gradually decreased (Fig. 3E) while fiber diameter persisted during the early period (0–5 days) (Fig. 3F) but changed dramatically as bulk erosion continued and the PGA knit was no longer apparent by SEM imaging or chemical analysis (7 days and beyond). Following loss of mechanical integrity of the PCLA sponge (at 7 days in vitro, estimated 7 months in vivo); the scaffold existed solely as small fragments that continued to degrade until no visible fragments remained at 14 days.

Results demonstrated further that the scaffold became progressively more compliant as it degraded (Fig. 3G). There was a sudden increase of compliance (i.e., decreasing slope of the pressure–strain curve) at 3 days in vitro (estimated 3 months in vivo), corresponding to loss of mechanical integrity of the PGA fibers (125 ± 21 kPa day 0, 116 ± 8 kPa day 1, 59 ± 14 kPa day 3; significantly different between days 1 and 3 based on a Tukey post-hoc test with p < 0.05) (Fig. 3H). By day 7, the scaffold lost mechanical integrity and could not withstand pressure (i.e. compliance approached infinity and stiffness decreased to zero) (65 ± 49 kPa day 5, 0 ± 0 kPa day 7). Similar findings emerged for equivalent burst pressure, which dropped precipitously at 3 days with the loss of PGA fiber integrity (>1000 kPa day 0 and day 1, 121 ± 21 kPa day 3, 101 ± 105 kPa day 5) and at 7 days the burst pressure was not measurably due to the scaffold being too degraded for testing with the loss of PCLA sponge integrity (Fig. 3I). Of note, from a physiologic perspective the scaffold remained relatively stiff compared to the native IVC over the physiologically relevant pressure range (1–0 mmHg) until after 5 days of accelerated degradation. Thus, these results suggested the change in inflammatory stimulation (presence of scaffold) and the stress-shielding properties (structural integrity of scaffold) are uncoupled, since the loss of structural integrity occurred well before the scaffold mass disappeared.

Role of the scaffold on G&R (in vivo time course study)

Next we evaluated scaffold-induced inflammation in vivo. TEVGs implanted as sheep IVC interposition grafts were harvested at 1, 6, 26, and 52 weeks after implantation and characterized using immunohistochemical (IHC) stains (Fig. 4A). Markers of the inflammatory response, including CD45 for leukocytes and CD68 for monocytes and macrophages, were present within the neotissue, particularly at earlier time points. Histological evaluation demonstrated degradation of the scaffold in vivo mirrored its degradation in vitro, including differential rates of degradation, with PGA fibers (solid black) degrading prior to the PCLA sponge (black outline) between 6 and 26 weeks (Fig. 4B). However, there was scattered residual scaffold material and patches of inflammatory cells remaining at 52 weeks, emphasizing differences between polymer degradation in vivo and in vitro. Also, the thickness of the polymeric scaffold in vivo increased from the manufactured value of 0.70 mm at implant to 1.3 ± 0.3 mm at 1 week and 3.0 ± 0.6 mm at 6 weeks (1- vs 6-week Tukey adjusted p < 0.001) then decreased to 1.9 ± 0.4 mm by 26 weeks (6 vs 26 week Tukey adjusted p < 0.001). As this thickening did not occur in PBS in vitro, it was likely not due to polymer swelling. Rather, cell infiltration and extracellular matrix (ECM) accumulation (inflammatory neotissue formation) accounted for most scaffold thickening observed following implantation.

IHC staining delineated the inflammatory cells throughout TEVG development. CD45+ leukocyte populations and CD68+ monocytes and macrophages rapidly populate the scaffold, residing within the pores. The inflammatory stimulus, measured by CD45+ cell density, continued to rise until reaching the highest measured levels around 6 weeks (437 ± 91 cells/mm²), after which this density decreased (257 ± 110 cells/mm² at 26 weeks, Tukey adjusted p-value vs 6 weeks p < 0.001), becoming closer to, but still higher than, the native background inflammatory cell density by 52 weeks (148 ± 63 cells/mm² TEVG vs 34 ± 28 cells/mm² IVC; 26- vs 52-week TEVGs, Tukey adjusted p < 0.01; 52-week TEVG vs IVCs, Tukey post-hoc p < 0.001). Pro-inflammatory (iNOS+) and anti-inflammatory (CD163+) macrophages and monocytes appeared to rise and fall in tandem throughout TEVG evolution. At 52 weeks, there were slightly higher levels of lingering anti-inflammatory compared to pro-inflammatory macrophages (31 ± 21 iNOS+ cells/mm² vs 65 ± 34 CD163+ cells/mm², t-test p = 0.008), likely related to long-term retention of low levels of foreign body giant cells within the neotissue.

We also evaluated the effect of the in vivo stress-shielding exhibited by the polymeric scaffold. Ex vivo biaxial mechanical testing of neovessels (Fig. 5A) demonstrated that at 6 weeks and 78 weeks the TEVG had a lower in vivo stretch than the native IVC (Fig. 5A) (1.17 ± 0.08 at 6 weeks, 1.11 ± 0.03 at 78 weeks, and 1.43 ± 0.04 for the IVC, Tukey adjusted p-value 0.515 for 6 vs 78 weeks, p = 0.00024 for 6 weeks vs IVC, and p = 0.00024 for 78 weeks vs IVC). The axial and circumferential wall stress were calculated at in vivo stretch and a representative pressure (10 mmHg for axial and 20 mmHg for circumferential). The axial wall stress (1.39 ± 0.60 kPa at 6 weeks, 11.89 ± 2.81 kPa at 78 weeks, and 14.77 ± 3.55 kPa for IVC, Tukey adjusted p-value < 0.0001 for 6- vs 78 weeks, p < 0.0001 for 6 weeks vs IVC, p = 0.224 for 78 weeks vs IVC) and the circumferential wall stress (3.71 ± 1.01 kPa at 6 weeks, 33.79 ± 0.69 kPa at 78 weeks, and 37.77 ± 5.26 kPa for the IVC, Tukey adjusted p-value < 0.0001 for 6- vs 78 weeks, p < 0.0001 for 6 weeks vs IVC, and p = 0.151 for 78 weeks vs IVC) was significantly lower for the 6 week group. Distensibility (0.0077 ± 0.0072 mm Hg⁻¹ at 6 weeks, 0.0090 ± 0.0025 mm Hg⁻¹ at 78 weeks, and 0.0640 ± 0.0609 mm Hg⁻¹ for IVC, with Tukey adjusted p-value = 0.997 for 6 vs 78 weeks, p = 0.024 for 6 weeks vs IVC, p = 0.066 for 78 weeks vs IVC) of the TEVG was significantly lower than the native IVC at both 6 weeks and 78 weeks (Fig. 5B, Table 1). Axial and circumferential stress approached native IVC values at 78 weeks post-implantation, though the in vivo axial stretch and distensibility remained lower than the native IVC, suggestive of an altered matrix composition and deposition history.

The biomechanical properties of the TEVG arise from a combination of the material behavior of the scaffold and that of the neotissue constituents. As the polymer is initially very stiff relative to the neotissue, it bears most of the pressure-induced load. Hence, the material behavior of the TEVG derives almost exclusively from the scaffold shortly after implantation. Furthermore, the increased thickness of the TEVG wall relative to the native vessel thickness in combination with the presence of the stiff polymeric constituents reduce the intramural stresses experienced by the cells (Fig. 5B). As such, the cells are stress-shielded during the early remodeling process, but the degradation of the polymer and transfer of the load to the deposited ECM allows long-term mechanical loads to contribute to ECM remodeling towards a native-like structure. With marked stress-shielding through 6 weeks, the ECM is initially disorganized, but becomes the predominant load bearing constituent after polymer degradation. Thus, there is significant remodeling and maturation over time to form a highly ordered structure (Fig. 5C, D), likely due to mechano-mediated G&R predicted by modeling.

The total area of neotissue, measured from the trichrome stain, increased up to the 6 week time point (12.7 mm² at 1 week vs 138.0 ± 37.7 mm² at 6 weeks), when the inflammation was at its highest recorded levels, then decreased as the inflammation waned and the stenosis self-resolved (65.4 ± 31.9 mm² at 26 weeks vs 30.3 ± 11.2 mm² at 52 weeks, Tukey adjusted p = 0.003). Cell content was high (81%) 1 week after implantation whereas collagen content was low (19%), as measured by trichrome staining. This ratio reversed over time, approaching at 52 weeks the low cellularity, high collagen content (78.8 ± 6.6% TEVG vs 91.3 ± 4.8% IVC trichrome collagen area, Tukey adjusted p < 0.001) of the native IVC. Picro-Sirius Red staining showed nearly the same collagen content in the TEVG at 52 weeks as the native IVC (85.0 ± 10.6% TEVG vs 86.5 ± 8.0% IVC, Tukey adjusted p = 0.649). The ratio of thick to thinner collagen fibers also changed as the TEVG evolved, with thicker (mainly type I) collagen fibers becoming an increasingly larger percentage of total collagen at later times. This ratio remained lower than that seen in the native IVC at 52 weeks (thick:thin of 23.5 ± 8.5 for the TEVG vs 43.3 ± 52.2 for the IVC, Tukey adjusted p = 0.224), although there was variation in the ratios for the IVC samples.

Hemodynamic changes

To assess effects of the evolving TEVG on hemodynamics and vice versa, we performed subject-specific computational simulations and calculated 3D maps of major hemodynamic indices. 3D anatomical models of TEVGs at 1, 6, and 52 weeks post-implantation (Fig. 6A) detailed structural changes. Subject-specific computational fluid–structure interaction (FSI) simulations in the respective geometries allowed calculation of velocity, pressure, and wall shear stress (WSS) and enabled in-depth spatio-temporal comparisons of morphological and hemodynamic parameters. Simulations highlighted characteristic changes seen in the natural history of TEVG development in vivo, from the cinching of the graft and IVC at the anastomoses to development of stenosis, evidenced by marked narrowing of the lumen cross-sectional area (Fig. 6B) and thickening of the wall (Fig. 6C), which reversed by 52 weeks. Effects of these morphological changes on hemodynamics are seen in (Fig. 6D, E), where the conical-like 6 week geometry (from proximal anastomosis to stenosis) caused a spike in WSS at its focal point (Fig. 6D). Time-averaged wall shear stress (TAWSS) along the TEVG generally increased from 1 to 6 weeks, particularly in the stenotic region, and again from 6 to 52 weeks, as the TEVG elongated and narrowed. Effects of pressure on the vessel wall were quantified using the Cauchy stress (Fig. 6E). Given the Laplace estimation of circumferential Cauchy stress for a thin-walled cylinder (Pr/h), that is, luminal pressure multiplied by radius and divided by wall thickness, the large increase in Cauchy stress from 1 to 6 weeks arose in part due to the dramatic increase in pressure at 6 weeks. Cauchy stress subsequently decreased from its 6-week level back to the 1-week level by 52 weeks of implantation. The 1- and 52-week values were indicative of how stiff the TEVG was compared to the IVC. Further quantifications of MRI measurements given in Supplemental Fig. 3.

Computational G&R analysis of inflammation-driven, mechano-mediated neotissue

Previous clinical studies and computational simulations demonstrated that the TEVGs were prone to early stenosis¹¹, but simulations revealed a possible spontaneous reversal as the balance of an immuno-dominant response (({rho }^{{{{{{rm{infl}}}}}}}gg {rho }^{{{{{{rm{mech}}}}}}}), ({Upsilon }^{{{{{{rm{mech}}}}}}}, < ,0)) shifted towards a mechano-mediated response (({rho }^{{{{{{rm{mech}}}}}}}gg {rho }^{{{{{{rm{infl}}}}}}}), ({Upsilon }^{{{{{{rm{mech}}}}}}}, > ,0)) between 25 and 50 weeks with scaffold degradation and geometric changes (Fig. 7A). We probed the importance of these mechanisms in silico by isolating the effect of each stimulus. When the immune response to the scaffold was eliminated numerically (set ({rho }^{{{{{{rm{infl}}}}}}}(s)=0)), the simulated TEVG experienced an early-onset rapid dilation due to a lack of early neotissue formation as the scaffold degraded. Over time, however, mechano-mediated neotissue progressively restored the TEVG towards its original diameter, which was mediated to reach to homeostatic WSS. Conversely, including only immuno-mediated neotissue production (set ({rho }^{{{{{{rm{mech}}}}}}}(s)=0)) led to an early stenosis followed by reversal, but the lack of mechano-mediated neotissue production led to substantial dilatation of the graft at late times as the immuno-mediated neotissue degraded without new production to replace it. Finally, without mechano-mediation to reduce the production of native-like neotissue (set ({Upsilon }^{{{{{{rm{mech}}}}}}}(tau )=0)) during the peak inflammatory response, the stenosed geometry was “locked-in”. These simulations highlight the relative importance of the extent and timing of both immuno- and mechano-mediated neotissue formation in TEVG behavior at different times during TEVG evolution.

From the G&R modeling results, we then quantified relative contributions of inflammation-driven and mechanical-mediated constituents from the predicted TEVG evolution on neotissue G&R over the simulated 52-week period, which confirmed that neotissue formation was driven primarily by inflammation during the first 26 weeks after implantation with a rapid transition to mechano-mediated neotissue formation thereafter (Fig. 7B).

Comparison of our computational G&R predictions of inflammatory versus mechanically stimulated neotissue production with explant histology (CD45 for inflammatory cells and calponin for mechanically stimulated neotissue) revealed good agreement. The relative amount of mechanical and inflammatory neotissues measured from histology was qualitatively similar at 26 weeks while the model predicted this similarity at approximately 36 weeks after implantation (Fig. 7C). In vivo measurements also demonstrated a longer-lasting inflammatory response than seen in the modeling predictions (Fig. 7C). Mechano-mediated growth predictions based on the FSI results and 3D anatomical models (Fig. 7D, E) further supported our timeline for inflammation-driven and mechano-mediated responses, with increasing inflammation-driven responses up to 6 weeks but modest at 52 weeks, when the mechano-mediated response accounts for most, but not all, of the in vivo thickness measurements. To test whether the long-term stress-mediated thickness and radius reflected homeostatic values, we evaluated the computational simulations using theoretical results inferred from native vessels²⁸. That is, for fold-increases in flow and pressure (ε and γ, respectively) relative to original (homeostatic) values, mean WSS and intramural stress can be written as

where Q is the volumetric flowrate, P the pressure, r the luminal radius, and h the wall thickness, with subscript 0 indicating homeostatic values. A return to homeostatic values of intramural stress and wall shear stress requires remodeled radius and thickness values to be (r={varepsilon }^{1/3}{r}_{0},h={varepsilon }^{1/3}gamma {h}_{0}). We took 52-week native IVC values to represent the homeostatic state. The difference between predicted (stress-mediated) and measured wall thickness thus represented the thickness contribution from the immune response. Similarly, the difference in predicted luminal radius from the mechano-adaptive case and the radial evolution measured experimentally demonstrate the effect of inflammation on vessel narrowing. The role of the immune response, i.e. the large difference in the mechano-adaptive prediction (dashed line, Fig. 7D, E) and the actual geometric evolutions (symbols, Fig. 7D, E) was apparent at 6 weeks but decreased dramatically by 52 weeks, further supporting that the immune response dominated early G&R before giving way to more mechano-mediated responses stimulated by the changing hemodynamics.

The relative contributions of, and relationships between, inflammation-driven and mechano-mediated G&R were demonstrated further by correlating our morphometric and IHC data over time. These data revealed a significant positive correlation between iNOS and intramural growth (linear regression p < 0.0001, R²=0.628), supporting the model-based prediction of inflammation-mediated luminal narrowing and experimental observations of inflammation-driven wall thickening. There was also a significant positive correlation between calponin and inward remodeling, although with a weaker R² correlation coefficient (linear regression p = 0.0023, R² = 0.278), supporting the model prediction of mediation of geometric changes by mechano-sensitive smooth muscle cells and our experimental data demonstrating the role of inward remodeling in stenosis around 6 weeks after implantation (Supplemental Fig. 4). Of note, a decrease in thickness of a pressurized tube while retaining identical material properties would have the effect of increasing the outer diameter through reduced structural stiffness. As the TEVG decreased in outer diameter and wall thickness, this suggests that a change in material stiffness is likely, as confirmed via biomechanical testing. Furthermore, eNOS staining for endothelial cells demonstrated little to no staining at 1 week, scattered staining along the lumens of the TEVGs at 6 weeks, and complete luminal staining at 26 weeks and beyond, similar to the staining along the lumen of the native IVC in appearance (Supplemental Fig. 5). Evolution of an intact endothelium corresponded with the cells’ potential ability to respond to mechanical stimuli exerted by WSS.

Neovessels evolve to resemble native vessels in structure and function

Our computational G&R model suggested that after scaffold degradation, G&R of the TEVG would progressively yield a neovessel that mimics the native vessels. At 52 weeks post-implantation, surface SEM demonstrated a contiguous luminal covering of endothelial cells, and en face immunofluorescent staining of the luminal surface with CD31 and eNOS suggested functional endothelial cells, although not necessarily having fully native levels of functionality (Fig. 8A). IHC comparisons of the neovessel and the native vessel demonstrated that the 52-week old neovessel had a thin laminated wall composed of layers similar to those seen in the native IVC (Fig. 8B). The intima was composed of a monolayer of CD31+ endothelial cells surrounded by concentric layers of calponin+ smooth muscle cells. Time course IHC studies revealed further that calponin+ smooth muscle cells increased rapidly from 6 to 26 weeks (calponin+ area fraction was 0.021 ± 0.014 at 6 weeks vs 0.060 ± 0.015 at 26 weeks, t-test p < 0.001), after which the levels remained similar to those seen in the native IVC (Fig. 8C: calponin+ area fraction of 0.055 ± 0.020 for the 52-week TEVG vs 0.050 ± 0.019 for the IVC, t-test p = 0.485). When examining the total amount of calponin+ area, it increased from 1 to 6 weeks, then stayed steady until 26 weeks before declining at 52 weeks as the neovessel matured.

To further characterize neovessel functionality, we subjected three long-term implants (>1.5 years) to vasoreactivity testing (Fig. 8D–G). Results demonstrated that the neovessels had similar contractile responses (as measured by force in a ring myograph) to that of the native IVCs in response to both potassium chloride (KCl) and endothelin-1 (ET-1) stimulation (Fig. 8D, E). The neovessels also vasodilated (as measured by force) similar to the native IVCs in response to acetylcholine (ACh) (Fig. 8F), an endothelial dependent generator of eNOS/NO, and sodium nitroprusside (SNP), an endothelial-independent NO donor (Fig. 8G). Together with the histological findings, resolution of inflammation, comparison of wall shear stress measurements, and the observation that the implanted scaffold is composed of polymer and therefore pharmacologically inert at implant, these results suggested the development of native-like structure and function.

Neovessels exhibit biological growth

In addition to investigating neotissue deposition and remodeling into functional neovessels, we sought to evaluate the biological growth potential of the TEVG. Biological growth refers to the progressive change in size, shape, and function that occurs during the development and maturation of an organism. Because we implanted the TEVG in juvenile (4-month old) lambs, we were able to evaluate the biological growth potential of the neovessels as the lambs matured to adult sheep (Fig. 9A). The lambs more than doubled in body mass during the first year following implantation (26.8 ± 3.8 kg at 1 week, 64.2 ± 5.5 kg at 52 weeks, t-test p < 0.001) and continued to grow steadily out to two years before leveling off (79.6 ± 9.2 kg at 104 weeks, 76.8 ± 11.2 kg at 156 weeks, t-test p = 0.446) (Fig. 9B). Comparing our implanted animals and age-matched non-implanted controls revealed similar growth, suggesting that the TEVG implant in the IVC did not cause any growth restriction of the animal.

Volumetric reconstructions based on serial 3D angiography of the TEVG demonstrated that the TEVG lumen initially decreased in volume, reaching its nadir at 6 weeks after implantation (3.6 ± 0.9 mL at 1 week vs 1.8 ± 0.9 mL at 6 weeks, t-test p < 0.001), then increased in volume over the ensuing 150-week period (3.5 ± 1.3 mL at 26 weeks vs 5.0 ± 2.5 mL at 156 weeks, t-test p = 0.018) (Fig. 9C, D). During the same period, the TEVG became progressively more compliant. Comparison of the luminal area deformation of the TEVG and IVC over the cardiac cycle by MRI revealed that the TEVG was relatively stiff upon implantation compared to the IVC at 1 week (0.24 ± 0.08 IVC vs 0.10 ± 0.02 TEVG, fractional area deformation, Mann-Whitney test p < 0.0001), but by 52 weeks post-implantation the neovessel appeared to pulse similarly to the surrounding native IVC (0.23 ± 0.12 IVC vs 0.26 ± 0.06 TEVG, fractional area deformation, Mann-Whitney test p = 0.400) (Fig. 9E, F). This increase in compliance was important because the IVC is a highly compliant vessel that changes its volume dramatically based on the hemodynamic forces, which allows it to function as a capacitance vessel. Yet, assessing growth based on volume or diameter alone could be confounded by differences in the hemodynamic states of an animal at different ages. In contrast, the length of a vessel was not affected by the hemodynamic state and therefore represented a better measure of biological growth capacity. Thus, we measured the change in TEVG length over time and compared it to the change in vertebral body height measured on the same angiogram. Serial measurements revealed that the TEVG initially decreased in length during the first 6 weeks (21.4 ± 2.2 mm at 1 week, 18.1 ± 3.1 mm at 6 weeks, t-test p = 0.001) then subsequently increased in length (21.0 ± 4.1 mm at 26 weeks vs 28.3 ± 2.3 mm at 156 weeks, t-test p < 0.001) at a rate similar to the rate of change in the vertebral body, which coincidentally had a length similar to that of the implanted TEVG (21.7 ± 1.6 mm at 26 weeks vs 27.12 ± 2.4 mm at 156 weeks) over the ensuing time course (Fig. 9G).