Vascularized organoid engineered by modular assembly enables blood perfusion

Abstract

Tissue engineering is one approach to address the donor-organ shortage, but to attain clinically significant viable cell densities in thick tissues, laboratory-constructed tissues must have an internal vascular supply. We have adopted a biomimetic approach and assembled microscale modular components, consisting of submillimeter-sized collagen gel rods seeded with endothelial cells (ECs) into a (micro)vascularized tissue; in some prototypes the gel contained HepG2 cells to illustrate the possibilities. The EC-covered modules then were assembled into a larger tube and perfused with medium or whole blood. The interstitial spaces among the modules formed interconnected channels that enabled this perfusion. Viable cell densities were high, within an order of magnitude of cell densities within tissues, and the percolating nature of the flow through the construct was evident in microcomputed tomography and Doppler ultrasound measurements. Most importantly, the ECs retained their nonthrombogenic phenotype and delayed clotting times and inhibited the loss of platelets associated with perfusion of whole blood through the construct. Unlike the conventional scaffold and cell-seeding paradigm of other tissue-engineering approaches, this modular construct has the potential to be scalable, uniform, and perfusable with whole blood, circumventing the limitations of other approaches.

Hierarchical structures self-assembled from discrete modular components are widespread in nature at both the molecular and macroscopic scales. Organs, for example, are built up from modular components such as the nephron or liver lobule. Adapting this principle, mesoscale modular arrangements have been self-assembled artificially for a range of engineering applications (1). Here we have devised a biomimetic modular approach to create a previously undescribed, potentially scalable tissue-engineered construct that ultimately might address donor-organ shortages or could be used as a vascularized in vitro 3D culture model. Laboratory-engineered tissues with dimensions greater than a few hundred micrometers require an internal vascular supply to attain clinically significant viable cell densities. We have assembled microscale modular components with thrombosis-inhibiting characteristics into a vascularized tissue, which was perfused with whole blood without significant platelet loss. The modular construct contained a uniform cell distribution and enabled the incorporation of multiple cell types in a form that is theoretically scalable, unlike other scaffold methods.

Engineering an artificial tissue conventionally involves seeding healthy cells into a preformed, porous, and biodegradable scaffold (2). The utility of such constructs, as a potential organ source to alleviate donor shortages, is limited, however, by the absence of an internal vascular supply, such that cell viability is maintained exclusively by oxygen diffusion from the surrounding host. At the cell densities of normal tissues, this diffusion limitation restricts viable engineered constructs to ≈100–200 μm in thickness (3). Devices with larger dimensions are possible only by reducing cell density or by tolerating a very low oxygen concentration. Moreover, achieving a high-density, uniform cell distribution throughout a scaffold is difficult because cells primarily populate the periphery (<1 mm) of the scaffold (4). Furthermore, typical scaffolds do not facilitate the controlled mixing of multiple cell types, which may promote long-term cell function through paracrine and other interactions, because the specific growth rates of the different cell types can alter their ratio and relative location. Strategies that enable fabrication of large vascularized constructs with high, uniform cell densities, within which multiple cell types can be incorporated, are essential for engineering whole organs comprised of highly metabolic cells. Although each of these scaffold-related problems is being addressed separately elsewhere (4–9), here we report the development of modular assembly as a theoretically scalable strategy to provide a vascularized construct with uniform cell density, here containing two cell types. Furthermore, we demonstrate the capacity of the endothelial cell (EC)-seeded construct to be perfused with whole blood in vitro, which has not previously been reported for a (micro)vascularized engineered construct.

Instead of seeding a preformed scaffold with cells, HepG2 cells (as a model cell) were encapsulated in short cylindrical collagen gel modules (Fig. 1 a and b), and the outer surface of these modules was covered with a confluent layer of human umbilical vein endothelial cells (HUVEC) (Fig. 1 a and c). These modules then were assembled randomly into a larger structure (here a tube) to form the modular construct (Fig. 1d). The interstitial spaces, formed between the assembled modules, constituted HUVEC-lined interconnected channels, on the order of a few hundred micrometers in size, that permeated the modular construct, enabling fluid and particularly blood perfusion. Even though these channels are not capillary-like in scale or structure, and they do not replicate the “tree-like” native vascular structure, the seeded ECs (uniform throughout the construct, unlike native vasculature) were expected to control the dynamic balance of pro- and anti-thrombogenic factors to maintain continuous blood flow without thrombosis. We envision that the modular construct, in an appropriate organ-like shape, will be connected to the vascular supply of the host by using appropriate host vessels or artificial vascular grafts. Modular constructs (albeit on a much smaller scale) resemble packed beds, a common and well understood component in chemical engineering (10).

Fig. 1.

Modular construct design and fabrication. (a) Collagen with or without HepG2 cells is drawn into the lumen of a sterilized PE tube and incubated at 37°C for 30 min to allow gelation. The PE tubing containing the gel is fed through an automated tubing cutter and sectioned into 2-mm lengths, which are collected in a sterile centrifuge tube. Cell culture medium is added, and the tube is vortexed to release the collagen–cell modules from the lumen of the sectioned PE pieces. PE sections float, whereas collagen modules sink. The collagen cylinders with encapsulated HepG2 cells are subsequently seeded with HUVEC. Once complete coverage of the collagen surface with HUVEC has been achieved (typically 2–3 days), the cell-seeded cylinders are assembled into a larger structure (here a tube) to form the construct. Assembly of the modules creates a network of interconnected channels that permeate the construct. Medium or blood is perfused through this network to supply nutrients to the cells within the construct. (b) Light micrograph of a collagen–HepG2 module before HUVEC seeding. (c) Confocal microscopy image of vascular endothelial (VE)-cadherin-stained module indicating a confluent layer of HUVEC over the module surface at 7 days after seeding. (d) Modular construct in the flow circuit being perfused with PBS. (e) Confocal microscope image of a collagen–HepG2–HUVEC module retrieved from a construct after 7 days of medium perfusion with HepG2 cells labeled with Vybrant CFDA SE.

The quiescent, nonthrombogenic EC lining within the channels of the modular construct is critical to enable whole blood to percolate around the modules with a significantly lower level of thrombosis than that associated with biomaterial surfaces. HUVEC are useful in this context because they express low basal levels of tissue factor, a potent coagulation initiator (11). Collagen was selected for the module base material in the prototype because HUVEC naturally reside on a type IV collagen membrane, albeit not the same type as that (type I) which is readily available. It is possible to incorporate other extracellular matrix components, such as type IV collagen, elastin peptides, and glycosaminoglycans, during module fabrication to further enhance the function of the modules.

This biomimetic modular design strategy has several intrinsic benefits. Constructs are potentially scalable: High-cell-density modules can be assembled into any overall size and shape with a uniform cell distribution simply by using more modules. The small dimensions of the modules and the interconnected channels through which blood flows are expected to allow for relatively high mass transfer coefficients [following the extensive engineering literature on flow through packed beds (10)] and hence good nutrient and oxygen supply to the encapsulated cells, which ideally should be within ≈100–200 μm of a flow channel. Furthermore, modules encapsulating different cell types can be mixed to prepare constructs with multiple cell types in controlled ratios and in different locations within the overall construct.

Results and Discussion

Collagen-HepG2 modules were fabricated (Fig. 1a) by gelling a solution of endotoxin-free collagen, containing suspended HepG2 cells, within the lumen of a small bore polyethylene (PE) tube. The tubing then was cut into 2-mm lengths by using an automated cutter and gently vortexed to remove the cell-containing collagen modules from the tubing lumen.^† Modules with different dimensions can be produced by using different tubing diameters and sectioning lengths. HepG2 cell viability within individual modules was >90%, and even with perfusion for 7 days (Fig. 1e) viable cell numbers were similar to, if not greater than, those of modules cultured under static conditions. After 7 days of culture, cell densities reached high values (0.3–1 × 10⁸ cells per cm³), depending both on cell growth and module contraction,^‡ within an order of magnitude of cell densities within tissues [10⁸ to 10⁹ cells per cm³ (12)].

One day after fabrication, modules were seeded and incubated with HUVEC under static conditions. Full surface coverage and shrinkage was achieved within 3 days (Fig. 1c). In some cases, HUVEC bridging of modules in close proximity was observed. HUVEC densities reached 4.5 ± 1.5 × 10⁵ cells per cm² (>90% viable) within 7 days, consistent with confluent HUVEC densities observed on tissue culture polystyrene.

Modules were randomly assembled into a modular construct by pipetting a suspension of modules into a larger tube positioned within a continuous-flow circuit (Fig. 1d). The modules produced from 4 m of collagen-filled PE tubing were sufficient to assemble a 0.5- to 1.0-cm long × 0.3-cm diameter construct (construct volume of 0.038–0.075 cm³). Constructs were perfused at physiological pressure differences [i.e., <100 mmHg (1 mmHg = 133 Pa)] with cell culture medium to simulate the flow of blood through a fully functional construct. Pressure difference vs. flow-rate flow profiles, obtained for two separate modular constructs (Fig. 2a), were used to estimate construct porosity and shear levels within the channels of the constructs. Analysis of these profiles by using the Ergun equation (13) indicated construct porosity was 22% and 24% (lower than expected; see below) in the two constructs shown in Fig. 2a. By using these porosity values, the average shear stress on the HUVEC (Fig. 2a Inset) was calculated to be in the range of 3–30 dynes/cm² (1 dyne = 10 μN) depending on the flow rate. The channels within a similar modular construct (prepared with a stiffer material) are shown in a microcomputed tomography image in Fig. 2b, and velocity estimates from Doppler ultrasound are shown in Fig. 6, which is published as supporting information on the PNAS web site. Both figures illustrate the interconnectedness of the channels and the laminar, well defined percolating flow profile in a modular construct. Exposure to flow (24 h at a flow rate of 0.08–0.11 cm·s⁻¹; shear ≈2–3 dynes/cm²) in the construct increased F-actin levels and elongated and flattened the HUVEC on the module surface (see Fig. 7, which is published as supporting information on the PNAS web site).

Fig. 2.

Characterization of constant porosity and shear stress on the surface of the modules during construct perfusion. (a) Flow and shear profiles through two collagen modular constructs. Flow rate of PBS through two separate constructs (construct length, 0.5 cm; construct diameter, 0.3 cm) as a function of applied pressure difference (hydrostatic head); open and filled points represent different constructs. Each point is the mean of two flow-rate measurements made at each pressure difference. (Inset) The slope of the fitted line was used to calculate construct porosity by using the Ergun equation (13) from which the shear stress on the surface of the modules was calculated for each construct. (b) Microcomputed tomography image of microfil cast of a poloxamine (22) modular construct (without HUVEC). Poloxamine is a stiffer material, enabling microfil casting. Light-colored regions correspond to the microfil (i.e., the channels), and dark regions correspond to modules, illustrating the interconnectedness of the flow channels that are normally lined with ECs. Porosity based on the number of light pixels was 22.6%. A relatively high pressure was required to fully infiltrate the viscous microfil, and hence this technique was not suitable for assessing porosity in a lower-stiffness HUVEC-coated collagen construct.

Because flow rate determines both the pressure difference generated across the construct during perfusion and the shear stress on the surface of the modules, the maximum flow rate that can ultimately be achieved is limited by the maximum shear stress to which the ECs can be exposed [typically 5–50 dynes/cm² (14), depending on the vessel from which the cells originate] before they are dislodged from the channel wall or exhibit an altered phenotype such as increased tissue factor expression (15). It is for this reason that cylindrical, instead of spherical, modules were selected during the initial design stages. Cylindrical modules were expected to result in greater construct porosity (10, 16), which, for a given flow rate, produces a lower shear stress on the module/endothelial surface. An aspect ratio (L/D) of 1.5 (as used here) has an associated (theoretical) porosity of ≈0.40 (cross-reference to ref. 10) compared with ≈0.36 (17) obtained with randomly packed incompressible spheres. This porosity difference translates into a 31% increase in achievable throughput rate at a given shear stress for cylinders over spheres. Not unexpectedly, the porosity in the prototype construct was lower than the theoretical, due to the compressibility of the modules and the presence of the HUVEC.

The seeded ECs maintained their nonthrombogenic phenotype as demonstrated by various assays, including ones involving whole-blood perfusion. The tissue factor activity (factor Xa generation chromogenic assay) of HUVEC-seeded modules cultured under static conditions was low (see Fig. 8, which is published as supporting information on the PNAS web site). HUVEC-covered modules produced significantly longer times to clotting (0.75 units/ml heparin; rocking platform arrangement; see Methods) than collagen-only modules (Fig. 3a; P = 1.4 × 10⁻⁵). In 9 of 14 trials using HUVEC modules, clotting had not occurred at test termination compared with 1 of 15 trials for collagen-only modules. The presence of the HUVEC significantly reduced the thrombogenicity of the module surface.

Fig. 3.

Characterization of module thrombogenicity using whole-blood studies. (a) Clot formation times. The presence of HUVEC on the modules significantly increased the time to clot formation (P = 1.4 × 10⁻⁵) of slightly heparinized whole blood (0.75 units/ml) in a clotting test. In some cases, clot formation never actually occurred, and the test was terminated between 4,500 and 5,400 s; in these instances, the recorded time was the test termination time. Mean clot time is represented by the thick central line within the box. Open circles and stars represent outliers and extreme outliers, respectively. (b) Fresh whole blood (0.75 units/ml heparin) perfused through a HUVEC-covered modular construct (filled circles) maintains platelet levels no different from those measured in the absence of modules (open circles, flow circuit blank; includes polypropylene mesh required to keep modules in place). Blood perfusion through a control modular construct in which HUVEC have been removed by dispase–collagenase action (open squares), however, results in significant reductions in platelet number, indicating platelet activation and the thrombogenic response that occurs in the absence of HUVEC. Error bars indicate SEM (n = 3, 4, and 7 for background, dispase-treated modular constructs, and HUVEC-covered modular constructs, respectively).

Lastly, and most significantly, slightly heparinized (0.75 units/ml) whole blood was perfused through the constructs at a rate of 0.334 ml/min (equivalent to ≈7 dynes/cm²), and the effluent was analyzed for platelet concentration (Fig. 3b). When constructs assembled from HUVEC-covered modules were perfused, there was no reduction in platelet concentration relative to the background changes associated with the flow circuit itself (i.e., measured in the absence of modules). Blood perfusion through collagenase-dispase–treated HUVEC modules (to remove the HUVEC layer after module shrinkage) significantly reduced platelet concentration in the collected perfusate. The reduction in effluent platelet concentration is an indicator of thrombogenicity in the absence of HUVEC; the absence of this reduction (relative to the background) is an indicator of the functional efficacy of the HUVEC-seeded modules in inhibiting platelet activation. Obvious thrombus formation (at 30 min) was seen in the majority of flow circuits without ECs, but not when ECs were present. The presence of the HUVEC significantly reduced the thrombogenicity of the construct. Although there are vascularization strategies involving VEGF delivery (5), EC implantation (18), and micromachining (19) followed by implantation within the omentum (20), such a demonstration of in vitro blood perfusion through a (micro)vascularized tissue engineering construct together with evidence of low thrombogenicity has not been described previously.

The potential for scalability arises because, unique to the modular approach, the underlying design principles can be delineated. The three main constraints that influence the design of the modular construct are nutrient supply, incorporating clinically significant numbers of cells within a construct of implantable volume, and the shear force on the HUVEC layer. Nutrient supply, determined by mass transfer within the construct, was estimated not to be a significant design constraint. Channel dimensions are expected to be of the same size as the modules (i.e., on the order of a few hundred micrometers) allowing good oxygen mass transfer, the likely limiting nutrient (12), within the construct channels. Moreover, HepG2 cells remained viable within an assembled construct over 7 days, suggesting that mass transfer to the encapsulated cells was sufficient, at least for the cell-seeding density and module size used. Because it has been predicted that a patient could survive on 10% of normal liver function (12), an engineered liver with the cell densities achieved in our construct (3–10% of tissue densities) could conceivably have sufficient cell mass to support patient survival. Studies of liver cell function and comparison with other liver constructs were beyond the scope of this investigation.

Controlling shear stress is the most critical constraint to achieving a quiescent, confluent, nonthrombogenic layer of HUVEC. Quiescent cells are unlikely to overgrow and block the channels of the construct. Here, HUVEC growth on module surfaces reached a plateau, which suggests that overgrowth is not a concern. Maintenance of a confluent layer of HUVEC, under flow, is also critical to allow long-term blood flow through the interconnected channels of the construct. A major limitation of endothelialized vascular grafts is incomplete cell coverage under flow conditions due to insufficient adhesion (21), resulting in limited protection from thrombosis. Full coverage of the modules was achieved within 3 days of seeding, and strong adhesion to collagen films was observed, as expected, in a centrifugation assay.^§ Most importantly, the HUVEC must exhibit a nonthrombogenic phenotype. Whole-blood perfusion of a prototype construct, assembled from HUVEC-covered modules, at shear rates equivalent to ≈7 dynes/cm², resulted in no significant increases in platelet loss above background levels, whereas constructs assembled from modules from which ECs were removed showed significant platelet loss within a short period. Together with the tissue factor and clotting time results, this result suggests that a functional nonthrombogenic layer of EC was generated on the module surface that was sufficient to maintain continuous blood flow through the engineered modular tissue.

We have demonstrated the use of microscale modular components in a biomimetic fashion to assemble uniform, potentially scalable (micro)vascularized tissue-engineered constructs containing multiple cell types, which were perfused with whole blood. The current prototype enabled maintenance of cell viability at high cell densities and whole-blood perfusion with minimal blood activation. The next step is exploiting the modular concept in a form that is suitable for in vivo use (e.g., adding components to enable anastomoses to the host vasculature; using biocompatible components) and understanding how the modular construct and the EC-lined channels remodel once implanted. Subsequently, it will be necessary to demonstrate the higher-level functions (e.g., polarity, spatial heterogeneity) and longer-term nonthrombogenicity of the endothelialized channels (at different shear stresses) and ultimately the utility of this approach to create functioning tissue or organ (e.g., liver, pancreas) equivalents, including extending the concept beyond HUVEC/HepG2 cells. Modular tissue assembly is a biomimetic alternative to traditional scaffold-based strategies, which offers many advantages for engineering whole-organ and large-tissue grafts and potentially transforms the conventional cell seeding/porous scaffold paradigm of tissue engineering.

Methods

Cell Culture.

The human hepatoma cell line, HepG2 (American Type Culture Collection, Manasses, VA), was cultured in 25-cm² tissue culture flasks in RPMI 1640 culture medium with l-glutamine (Invitrogen Canada, Burlington, ON, Canada) supplemented with 15% bovine calf serum (HyClone) and 2% penicillin/streptomycin (Invitrogen Canada) at 37°C in a 5% CO₂/95% air humidified atmosphere. HUVEC (Cambrex, Walkersville, MD) were cultured in 75-cm² tissue culture flasks in EGM-2 medium, as suggested by the suppliers, supplemented with EGM-2 bullet kit (Cambrex) at 37°C in a 5% CO₂/95% air humidified atmosphere. In modules where both cell types were present, both cell types were cultured in HUVEC culture medium.

Module Fabrication.

Vitrogen collagen solution (Type I, bovine dermal, 3.1 mg collagen per ml; Cohesion Technologies, Palo Alto, CA) was mixed with 10× minimum essential medium (125 μl of 10× medium per ml of collagen; Invitrogen Canada) and neutralized by using 0.8 M NaHCO₃ (Sigma-Aldrich Canada). Pelleted HepG2 cells were mixed with the neutralized collagen (2 × 10⁶ cells per ml), and the solution was drawn into the lumen of an ethylene oxide gas-sterilized PE tube (0.76 mm inner diameter × 1.22 mm outer diameter) connected to a syringe at one end. After 30-min incubation to allow collagen gelation, the gel-filled tubing was cut into 2-mm lengths by using a custom-built automated cutter (Fig. 1a; FCS Technology, London, ON, Canada). Sections were vortexed gently in cell culture medium to remove the gel-cell module cores from the tubing lumen. The collagen–cell modules were allowed to settle, separated from the PE tubing, and cultured in Petri dishes under static conditions. Collagen-only modules were fabricated identically (same collagen concentration) without the addition of the HepG2 cell pellet.

EC Seeding.

HUVEC (Passage 1–6; 1.5–2.0 × 10⁶ cells per ml of settled modules) were added to modules with or without encapsulated HepG2 cells in a 15-ml centrifuge tube and incubated for 60 min with gentle shaking every 10 min. Modules then were transferred into a nontissue culture polystyrene Petri dish. Medium was replaced every 1–3 days.

Module Dimensions.

After incubation overnight, a sample (n = 96) of modules containing HepG2 cells was selected, and light microscopy images were taken of each module in a 96-well plate (one module per well) with an Olympus microscope. Modules then were seeded with ≈1.5 × 10⁶ HUVEC per ml of settled modules and incubated for 4 days, after which they were reimaged. Measurements of module diameter and length, before and after EC seeding, were made by using ImagePro software (Media Cybernetics, San Diego, CA).

Cell Viability and Enumeration Within Modules.

Cell metabolism of encapsulated cells was measured by using the Alamar blue (AB) assay at days 1, 3, and 7. Briefly, a micropipette was used to add 10 modules (3 replicates) containing HepG2 cells in a 200-μl volume into a 24 well plate. 10% AB (BioSource International, Camarillo, CA) was added, and the sample was incubated for 7 h. Supernatant samples were transferred into a 96-well plate and read by using a Sunrise ELISA plate reader (Tecan, Maennedorf, Switzerland) at 570 and 600 nm. Module samples then were digested with collagenase (final concentration 0.236 mg/ml in culture medium; Sigma-Aldrich Canada), incubated overnight, and stained with Trypan blue. The numbers of live and dead cells were counted manually with a hemocytometer.

To assess cell viability within the assembled construct, modules containing HepG2 cells or collagen-only modules seeded with HUVEC were cultured under static conditions for 6 days and then within a flow circuit (see below) for 24 h. Modules were retrieved from the circuit and tested immediately for viability by digestion and staining with Trypan blue as above. The viability of HepG2 cells cultured within an assembled construct for 1 week was assessed by using Vybrant carbofluorescein diacetate succinimidyl ester (CFDA) SE prelabeled cells (10 μM; Molecular Probes, Burlington, ON, Canada). One day after fabrication, the HepG2 modules were seeded with HUVEC and then, after 2 days incubation, were assembled into a construct within a flow circuit to allow module shrinkage. After 1 week of medium perfusion, modules were retrieved from the flow circuit, fixed in 3.7% paraformaldehyde–PBS (Electron Microscopy Science, Hatfield, PA) for 30 min, washed in PBS, and observed by using fluorescence microscopy (Zeiss, Axiovert 135).

Construct Assembly and Flow Circuit Perfusion.

Fifty-milliliter centrifuge tubes with two holes punctured in the cap through which to thread Masterflex L/S-13 and L/S-16 tubing (Labcor, Anjou, QC, Canada) and ≈0.015 g of glass wool (≈0.075 cm³) (to hold the modules in place) were assembled into a continuous-loop flow circuit with a number of other connectors and stopcocks (various suppliers). A Masterflex peristaltic pump was used to circulate medium through the flow loop from a 19-ml reservoir. Modules (0.5–1.0 ml) were loaded into the circuit (total circuit volume 20 ml), within a laminar flow hood, by using a 10-ml pipette through a luer lock connector. Modules were maintained in the flow circuit at 37°C in a 5% CO₂/95% air humidified atmosphere for 24 h or 1 week. Medium was added to the reservoir every 1–2 days.

Flow Profile Measurements and Porosity Determination.

Pressure difference across the construct was recorded with low-pressure gauges (H. O. Trerice, Oak Park, MI) inserted on either side of the construct. Duplicate measurements of flow rate through the construct were measured for a range of pressure differences by the timed collection of 0.5 ml of medium from the circuit through a T-connector output. The gradient of flow rate vs. pressure difference (Darcy's permeability) was calculated, and the gradient of similar curves measured in the absence of a construct (i.e., with glass wool only present) was subtracted to isolate the pressure difference contribution from the construct. The Ergun equation (13) then was solved for porosity by iteration using the Solver program in Microsoft Excel. The values used for constants present in this equation were as follows: length of construct, 0.5 cm; fluid viscosity, 0.01 g/cm³; module diameter, 0.0411 cm; and shape factor, 0.874.

A construct of the same diameter and length was prepared by using modules of a stiffer material (20% poloxamine modules, the preparation of which is described in ref. 22), which enabled perfusion with the viscous microfil solution (“low viscosity”; Flow Tech, Carver, MA; component:diluent ratio of 4:15, 10% curing agent) used for microcomputed tomography [Mice Imaging Centre (MiCe), Hospital for Sick Children, Toronto, ON, Canada]. By using MicroView software (GE Healthcare, Chalfont St. Giles, U.K.), the number of pixels above the threshold corresponding to the microfil was used to calculate the volume fraction of microfil and hence the construct porosity.

Clotting Time.

Fresh whole blood (10 ml) was collected from consenting donors (with ethics approval by the University of Toronto), who had not taken medication within 72 h of phlebotomy, into a syringe containing heparin (final concentration 0.75 units/ml), after discarding the first milliliter. A 350-μl sample of slightly heparinized blood was mixed with 200 μl of collagen or HUVEC-coated modules in a microcentrifuge tube. A 400-μl sample of this mixture then was pipetted into a 25-cm length of polypropylene tubing (1.57 mm ID) connected at either end with Silastic tubing (1.57 mm ID) to 200-μl pipette tips connected to a rocking platform (23). Rocking was initiated, and the time until blood motion ceased or significant clot deposition occurred within the tubing was recorded as the clotting time.

Construct Perfusion.

Constructs were assembled from HUVEC-covered modules that were untreated or treated for 15 min in 100 mg/ml collagenase dispase solution (Roche, Mississauga, ON, Canada) to remove all HUVEC from the surface (for control modules), yet retain the size and stiffness of the contracted collagen. The short treatment time ensured that module dissolution did not occur, and microscope observation confirmed removal of the HUVEC layer. Constructs were assembled within a 0.2-ml length of a 1-ml graduated pipette and held in place at both ends with 1-cm² sections of polypropylene mesh (PPM-3; Biomedical Materials, Slatersville, RI). Silastic tubing (10 cm, 3.18 mm ID) was used to connect the pipette section to the syringe pump (824E Infusion pump model A-99; Razel Scientific Instruments, Fairfax, VT). The construct was prefilled with PBS to prevent air bubble formation.

Blood was collected in a 10-ml syringe from consenting volunteers that had taken no medication into 0.75 units/ml heparin. It was necessary to use a small amount of heparin (0.75 units/ml is much less than the 5 units/ml needed to stop all coagulation) to prevent premature clotting during the blood draw or while the blood was sitting in the syringe pump. The Silastic tubing was filled with blood from the syringe before being connected to the PBS prefilled pipette/construct section. The syringe then was placed on the syringe pump located on a rocking platform (to minimize blood settling) within a 37°C oven, and blood was perfused through the construct at a rate of 0.334 ml/min. At regular intervals during perfusion, 400-μl samples of the perfusate were collected in 0.6-ml graduated microcentrifuge tubes containing 8 μl of 200 mM EDTA. An initial sample was collected from the syringe before connecting it to the construct. The experiment was terminated when all of the blood from the syringe had been used or if circuit blockage occurred. Constructs were removed and dissected for evidence of thrombus formation within the construct or the polypropylene mesh.

Statistics.

Student's t test was used to determine significant difference when only two treatment groups were being compared. ANOVA was used to test for significant differences among multiple test groups. Q-Q plots were used to assess the normality of the data. Levene's test for homogeneity was used to test for equal variance among samples (24). When equal variance could be assumed, the Tukey HSD post hoc test was used to identify significant differences among multiple test groups. When equal variance could not be assumed, the Games–Howell (25) post hoc test was used to identify significant differences among multiple test groups. All tests were two-tailed, and P = 0.05 was considered significant. Further details on the methods are available in Supporting Methods, which is published as supporting information on the PNAS web site.

Abbreviations

EC

endothelial cell

HUVEC

human umbilical vein ECs

AB

Alamar blue

PE

polyethylene