Perfusable branching microvessel bed forvascularization of engineered tissuesLoraine L. Y. Chiua, Miles Montgomerya,b, Yan Lianga, Haijiao Liub, and Milica Radisica,b,c,1

aDepartment of Chemical Engineering and Applied Chemistry, bInstitute of Biomaterials and Biomedical Engineering, and cHeart and Stroke/Richard LewarCentre of Excellence, University of Toronto, Toronto, ON, Canada M5S 3G9

Edited by Robert Langer, Massachusetts Institute of Technology, Cambridge, MA, and approved October 24, 2012 (received for review June 28, 2012)

Vascularization is critical for the survival of engineered tissues invitro and in vivo. In vivo, angiogenesis involves endothelial cellproliferation and sprouting followed by connection of extendedcellular processes and subsequent lumen propagation throughvacuole fusion. We mimicked this process in engineering an orga-nized capillary network anchored by an artery and a vein. Thenetwork was generated by inducing directed capillary sproutingfrom vascular explants on micropatterned substrates containingthymosin β4-hydrogel. The capillary outgrowths connected be-tween the parent explants by day 21, a process that was accel-erated to 14 d by application of soluble VEGF and hepatocytegrowth factor. Confocal microscopy and transmission electron mi-croscopy indicated the presence of tubules with lumens formedby endothelial cells expressing CD31, VE-cadherin, and von Wille-brand factor. Cardiac tissues engineered around the resulting vas-culature exhibited improved functional properties, cell striations,and cell–cell junctions compared with tissues without prevascula-rization. This approach uniquely allows easy removal of the vas-culature from the microfabricated substrate and easy seeding ofthe tissue specific cell types in the parenchymal space.

microvasculature | contact guidance | controlled release | angiogenicfactors | cardiac tissue engineering

Tissue engineering may offer alternative treatment options fortissue and organ replacement, but challenges related to vas-cularization in vitro and in vivo remain. Organized vasculature isimportant for the integration of blood vessels with other tissue-specific cell types to create complex tissues (1), especially cardiactissues that require small intercapillary distances for metaboli-cally active cardiomyocytes (2). Moreover, organized vasculatureallows easy isolation, manipulation, and implantation of thevascular structures (1). Strategies developed to engineer vascu-larized tissues generally have involved (i) coculture of tissue-specific cells with endothelial cells (ECs) or their precursors (3,4); (ii) incorporation of preexisting endothelial networks orblood vessels (5); or (iii) engineering of proangiogenic scaffoldswith peptides, growth factors, or geometric cues (6–11).The incorporation of ECs can accelerate vascularization of the

implanted biomaterial or engineered tissue and its functionalanastomosis with the host vasculature (5, 12). In vivo, vascular-ization can be enhanced by cultivating cells around an arterio-venous loop (5) or by implanting materials into omentum (10).Alternatively, pore size alone (interconnected pores 30–40 μm indiameter) can promote angiogenesis in porous materials (6).Peptides and growth factors such as thymosin β4 (Tβ4) (13, 14),VEGF (7–9), and angiopoietin-1 (8, 9) lead to increased vesseldensity when incorporated into tissue-engineering biomaterials insoluble, encapsulated, or immobilized form. Recent studies havefocused on improving the organization of ECs using microcontactprinting and microgrooved substrates (15–17).Although these methods suggest that improving vascularization

enhances engineered-tissue function, limitations in the controlover cell position and the ultimate organization of the vasculaturelead to the formation of networks with random spatial distribu-tion, with implications for network function (10, 18). Recent

breakthrough studies enable microfabrication of microvascularstructures with barrier function (19); however, when the geometryof tubules was controlled, there was no facile way to seed cells inthe parenchymal space, because it was occupied by a synthetic ornatural polymer (1, 19). Additionally, these approaches do notenable easy removal of the engineered microvascular bed fromthe microfabricated mold or microfluidic device used to housethe vasculature.Here, we devised a method to engineer a prototype vascular

network consisting of two branching vessels that form a micro-vascular bed suitable for rapid vascularization of engineeredtissues in vitro. Our approach integrates the use of topographicalcues and sustained release of angiogenic factors (Fig. 1A).Rather than forcing isolated ECs into defined positions on apolymeric or hydrogel matrix, we used biomaterials to createa niche that directed sprouting and anastomosis of the vascula-ture from the two branching vessels in vitro. We demonstratedthe utility of this vascular network for improving the function ofengineered myocardium. Functional properties of myocardiumare determined by cardiomyocytes, a cell type sensitive to oxygenlimitations because of their high metabolic activity. The presenceof a microvascular bed in an engineered cardiac tissue in vitromay provide a biomimetic milieu for functional cell assembly,leading to cardiomyocyte orientation and survival.Our strategy involved controlled release of Tβ4 from the

substrate coating to mimic the in vivo environment in whichbiomolecules are present to promote formation of vascularnetwork. Tβ4 is an angiogenic and cardioprotective peptide thatenhances cardiomyocyte survival by inducing coronary vascular-ization and up-regulation of Akt activity (14, 20). Tβ4 was shownpreviously to initiate angiogenesis and sustain vascular stabilitybecause of its ability to recruit endothelial and smooth musclecells (13, 14). Compared with angiogenic growth factors, Tβ4has advantages such as small size, high activity, minimal immu-nogenicity, and high solubility in water.Soluble biomolecules are easily degraded and washed out, re-

quiring multiple doses at increased levels to maintain bioactivity(21). We previously developed a collagen-chitosan hydrogel ca-pable of releasing Tβ4 at nearly zero-order kinetics from day 3 today 28, with the release behavior over 28 d well-fitted to Higuchi’ssquare root equation (22). Although collagen promotes cell at-tachment, survival, and proliferation, the interconnected pores inthe collagen-chitosan hydrogel and the electrostatic interactionsbetween negatively charged Tβ4 and positively charged chitosancontribute to the sustained release of Tβ4 (22). In contrast, Tβ4

Author contributions: L.L.Y.C. and M.R. designed research; L.L.Y.C., M.M., Y.L., and H.L.performed research; L.L.Y.C., Y.L., H.L., and M.R. analyzed data; and L.L.Y.C., M.M.,and M.R. wrote the paper.

The authors declare no conflict of interest.

This article is a PNAS Direct Submission.1To whom correspondence should be addressed. E-mail: m.radisic@utoronto.ca.

See Author Summary on page 20176 (volume 109, number 50).

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1210580109/-/DCSupplemental.

E3414–E3423 | PNAS | Published online November 26, 2012 www.pnas.org/cgi/doi/10.1073/pnas.1210580109

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mailto:m.radisic@utoronto.cahttp://www.pnas.org/content/109/50/E3414/1http://www.pnas.org/lookup/suppl/doi:10.1073/pnas.1210580109/-/DCSupplementalhttp://www.pnas.org/lookup/suppl/doi:10.1073/pnas.1210580109/-/DCSupplementalwww.pnas.org/cgi/doi/10.1073/pnas.1210580109

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Fig. 1. Substrate topography and controlled release of angiogenic peptide Tβ4 guide capillary outgrowths from arteries and veins. (A) Experimental setup. (i) PDMSsubstrates were fabricated using standard soft lithography methods and (ii) were coated with collagen-chitosan hydrogel with or without Tβ4. (iii) Arteries and veinswere isolated and placed on the two ends of the substrate and (iv) were cultivated for 2 wk with HGF or VEGF supplementation or for 3 wk without any growthfactors. (v) Cardiomyocytes were seeded onto the engineered vascular bed and cultured for additional 7 d to grow a beating, vascularized cardiac tissue. (B) Rep-resentative images of cell outgrowths frommouse artery and vein explants on nonpatterned substrates at day 14 of cultivation (n = 3 per group). The asterisks indicatethe location of explants; arrows indicate locations of outgrowths. (Scale bars, 200 μm.) (C) Density of migrated cells from artery explants at various time points. (D)Density of migrated cells from vein explants at various time points. (E and F) Autocrine VEGF secretion was dependent on the Tβ4 dose and correlated with celloutgrowth density. (E) Total amount of VEGF measured with artery explants between days 12 and 14 (n = 3 per group; P = 0.0034, one-way ANOVA; P < 0.05 forcontrol vs. both Encap100 and Encap1500). (F) Total amount of VEGF measured with vein explants between days 12 and 14 (n = 3 per group, P = 0.0036 for control vs.Encap1500; P = 0.0055 for Encap100 vs. Encap1500). (G–J) Tβ4 increased the branch length of capillary outgrowths frommouse artery and vein explants at day 14, withthe most pronounced effect on substrates with grooves 50-μmwide. (G) Representative images of outgrowths (n = 10–12 per group). (Scale bars, 100 μm.) (H) Branchdensity on grooves with different widths (a branch is defined by the distance between two nodes or between a node and the end of a tube; P = 0.0002, two-wayANOVA; P < 0.01, Bonferroni posttest for control vs. Tβ4 gel groups on 100-μm grooves; P = 0.7078 within Tβ4 gel groups). (I) Average branch length on grooves withdifferent widths (P < 0.0001, two-way ANOVA; P < 0.05 for control vs. Tβ4 gel on 25-μm grooves; P < 0.001 for control vs. Tβ4 gel on 50-μm and 100-μm grooves; P <0.001 for 50-μm grooves vs. both 25-μm and 100-μm grooves in Tβ4 gel groups). (J) Average branch width on grooves with different widths (P = 0.3782). Micro-patterned PDMS substrates had grooves 25, 50, and 100 μm in width and 65 μm in height. Experimental groups include hydrogel without Tβ4 (control) and hydrogelwith 1,500 ng encapsulated Tβ4 (Tβ4 gel). The asterisks in graphs indicate statistically significant differences between groups (P < 0.05, two-way ANOVA withBonferroni posttests for cell density, branch density, branch length, and branch width; one-way ANOVA with post hoc Tukey tests for VEGF expression).

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was released quickly (within 3 d) from collagen-only hydrogel thatwas negatively charged and did not exhibit the interconnectedporous structure of collagen-chitosan hydrogel (22). Here, weused Tβ4 to generate vascularized cardiac tissues because ofits angiogenic and cardioprotective potential, and our hydrogeldelivery system enabled localized and sustained activity. We ap-plied the Tβ4-encapsulated hydrogel onto micropatterned poly-dimethylsiloxane (PDMS) substrates with 25-, 50-, or 100-μm–wide grooves (Fig. 1A and Fig. S1). Subsequently, mouse, rat, orhuman arterial and venous explants were placed at the two endsof the substrate to promote outgrowth of capillaries.

ResultsTo determine if the inclusion of Tβ4 improved cell recruitmentand accelerated network formation, we exposed mouse arterialand venous explants to different amounts of Tβ4 encapsulatedin collagen-chitosan hydrogel (0 ng for control, 100 ng forEncap100, and 1,500 ng for Encap1500). Increased cell outgrowthdensity was observed with a higher dose of Tβ4 (1,500 ng vs. 0 ngand 100 ng; Fig. 1 B–D). In the control group, higher outgrowthdensity was observed in venous explants than in arterial explants.However, there were no significant differences between the out-growth densities in arterial vs. venous explants when Tβ4 wasapplied (compare Fig. 1 C and D).Although Tβ4 alone can stimulate EC migration and differ-

entiation (14, 23), Tβ4 also stimulates the synthesis and auto-crine secretion of VEGF, as previously reported (24). Here, weobserved enhanced VEGF secretion from explants cultivated onTβ4 hydrogels compared with Tβ4-free gels (Fig. 1 E and F). Ourbasal culture medium contains serum, which may be the sourceof VEGF and was reported previously to stimulate quiescentECs to proliferate (25). We measured the basal VEGF con-centration in the culture medium as 0.06 ± 0.006 ng/mL byELISA. However, this level was increased by autocrine secretionin the presence of cultivated explants (Fig. 1 E and F). Ap-proximately 20% of the secreted VEGF was released to themedium, and 80% remained in the hydrogel (Fig. S2). In gen-eral, the basal level of VEGF secretion was higher with venousexplants than with arterial explants cultivated without Tβ4(compare Fig. 1 F and E), correlating with the greater celldensity on control hydrogel in the presence of venous explants ascompared with arterial explants (white bars in Fig. 1 C and D).The amount of VEGF measured in the culture medium for ar-tery explants at day 14 was increased with the application ofeither 100 ng or 1,500 ng Tβ4. For vein explants, the amount ofVEGF was increased with 1,500 ng Tβ4 gel as compared witheither control or 100 ng Tβ4-gel. Thus, there was a correlationbetween VEGF secretion and cell outgrowth densities. A posi-tive linear correlation was found when plotting the averageVEGF concentration produced between days 12 and 14 for eachgroup (Fig. 1 E and F) against the corresponding average celldensity at day 14 (Fig. 1 C and D) (r = 0.85, P = 0.03; Fig. S3).The Encap1500 group (referred to as “Tβ4 gel”) was used forfurther studies because of the observed dose response of bothVEGF secretion and cell outgrowth densities to Tβ4.We next aimed to determine if we can guide and control the

sprouting from mouse arterial and venous explants through theintegrated use of topographical cues (25-, 50-, and 100-μm–widegrooves) and the controlled release of Tβ4. Previously, exoge-nous, soluble Tβ4 was found to increase sprouting from coronaryartery rings in an angiogenesis assay (23), but the directionality ofthe outgrowths was not controlled. We evaluated three parame-ters to describe the sprouting process: branch density, branchlength, and branch width. Increased branch density correlateswith a higher level of sprouting often found during angiogenesis.Increased branch length is necessary for rapidly forming con-nections between the parent explants. Ideally, the branch width

should be similar to that of the capillaries in the nativevasculature.Both the branch density and length within the outgrowths

from mouse vascular explants (Fig. 1 H and I) were increasedwith the addition of encapsulated Tβ4 in hydrogel coating.Branch density for the Tβ4 gel groups was not affected by groovewidth (Fig. 1H). However, branches were significantly longer on50-μm grooves than on other groove widths (Fig. 1I). The lowerbranch lengths on substrates with smaller groove size likely wererelated to the hindrance of tube growth by spatial limitations forthe sprouts growing inside the grooves. At larger groove widths,capillary outgrowths tended to branch more because of the widechannels available for expansion of the vascular structure, thuslowering branch length (Fig. 1 G and I). There was no significantdifference in tube widths in the control and Tβ4 gel groups (Fig.1J). Tubes grown on substrates with 25-, 50-, and 100-μm groovesshowed comparable widths of ∼10–12 μm, typical of capillaries inthe native mouse myocardium (∼10–20 μm) (26). Thus, we used50-μm–wide grooves in further studies. Arterial and venousexplants did not differ significantly in branch density, length, orwidth, and the results are shown as lumped rather than sepa-rately for arteries or veins.Although both cell outgrowth density and autocrine VEGF

concentration were higher with vein explants (Fig. 1 D and F),there were no significant differences in branch density, length,and width in artery and vein explants (Fig. 1H–J). VEGF receptor2 (VEGFR2) is considered to be the major mediator of physio-logical effects such as proliferation, migration, survival, andtube formation caused by VEGF165. Binding of the VEGF toVEGFR2 causes receptor dimerization and autophosphorylation,followed by the activation of a number of downstream pathwaysto mediate different processes. For example, activation of theERK pathway regulates proliferation (27); activation of the p38MAPK, PI3K, and Rac pathways regulates migration (28–30);and activation of Src regulates both migration and vascular per-meability (29). Tube formation by VEGFR2–VEGF165 signalingwas reported to be mediated by MAPK (31, 32), p38 (31), andPI3K (30, 32, 33). Therefore, because of the complexity and inter-connectivity of the intercellular signaling pathways, increased cellproliferation and migration may not translate proportionally toincreased tube formation.Our results clearly indicate that the enabling factor in tube

formation and guidance was the presence of topographical cues,whereas the presence of angiogenic factor Tβ4 simply acceler-ated the formation of the tubes and enhanced their length (Fig.1). Introduction of topographical cues was essential for the for-mation of branches with open lumens, because Tβ4 gel alone ledto cell outgrowth without the formation of luminal structures(Fig. 1B). Topographical cues themselves can enable tube for-mation by contact guidance, changing the local mechanicalproperties of the environment, or locally increasing the con-centration of autocrine growth factors. We aimed to understandfurther the factors by which topographical cues affect tube for-mation by considering appropriate controls: (i) flat PDMS sub-strates with control or Tβ4 hydrogel, and (ii) uncoated PDMSsubstrates with 50-μm grooves and Tβ4 placed in the culturemedium (Fig. 2A). Flat controls demonstrated cell migrationfrom the artery or vein explants, with more cell outgrowths onTβ4 gel-coated substrates than on control gel-coated substrates.However, the cells did not form any vascular tube structures onflat substrates, thus indicating the importance of topography fortube formation and organization. Although controls with un-coated grooved substrates led to tube formation by migrated cells,these vascular structures were not well organized. This resultshows the importance of the localization of the Tβ4 molecules tothe groove/ridge space, because Tβ4 was present in the culturemedium but was not localized within the hydrogel coating in thiscase (Fig. 2A).

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We also measured local Young’s modulus on hydrogel-coatedgrooved and flat substrates by atomic force microscopy nano-indentation (Fig. 2B) and found no significant differences in thestiffness of the groove area and the space between grooves. Tβ4encapsulation in the hydrogel is not expected to change themechanical properties appreciably, based on our previous study(22). Only plain collagen-chitosan hydrogel was tested here,because we previously found no significant differences in our

rheological measurements with or without Tβ4 encapsulation(22). All hydrogel-coated substrates had similar local mechanicalproperties, irrespective of the groove dimensions. The stiffnessof uncoated flat PDMS substrate was measured as 5.85 ± 0.16MPa. This value is comparable to the local Young’s modulus (6.6MPa) previously obtained for the same PDMS material usingatomic force microscopy but is much higher than the bulkYoung’s modulus of PDMS (2.5 MPa) as measured by tensile

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Fig. 2. Topographical cues promote tube formation by providing contact guidance and locally increasing the concentration of autocrine growth factors but notby affecting the local mechanical properties. (A) Contact guidance. YFP mouse artery and vein explants were cultivated on flat PDMS substrates or uncoatedsubstrates. Flat substrates were coated with either control gel (Top) or Tβ4 gel (Middle). Explants were cultivated for 21 d. The vein on uncoated substrate(Bottom Right, Inset) shows unorganized tube formation. Asterisks indicate the locations of artery or vein explants, and yellow arrows indicate the presence oftube formation by outgrown cells. (Scale bars, 50 μm.) (B) Local mechanical stiffness. Local Young’s modulus for hydrogel-coated PDMS substrates withoutgrooves and with 25-, 50-, and 100-μm–wide grooves, as measured by atomic force microscopy in a 50 × 50 μm area spanning a part of the gel-coated groove anda ridge for each substrate. Uncoated flat PDMS substrate also was measured for comparison. The local stiffness of uncoated flat PDMS substrate was significantlyhigher than that of all coated substrates (P < 0.0001, one-way ANOVA), but no differences were found among coated substrates. (C) Local concentration ofautocrine growth factors. A simplified mathematical model of the VEGF concentration gradient in grooved and smooth samples, showing the effect of grooveson the local increase in the concentration of autocrine growth factors. (i and ii) Concentration profiles generated for different substrates. (i) Cross-sectionsshowing a single groove (small rectangular region at the bottom) and the height of the culture medium on top of the substrate for each substrate. (ii) Cropped100 × 200 μm close-up view showing the cross-section of a single groove for each substrate. The double-ended arrow indicates the width of the channel (25, 50,or 100 μm). The vertical dotted line indicates symmetry. The centrally positioned cell is shown as a black semicircle. The bottom of the image represents the topsurface of the substrate for the flat substrate case. (iii) Horizontal concentration profile of VEGF along the bottom of the channel for grooved substrates and atthe surface of the flat substrate, shown as relative VEGF concentrations with all values normalized to the value for flat substrate at the point (0, 0). In this steady-state model, VEGF was assumed to be secreted at a zero rate from a cell centrally positioned at the bottom of the groove or on a flat substrate.

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testing (34). Previous studies comparing local mechanical prop-erties measured by atomic force microscopy and bulk mechanicalproperties as determined by a tensile tester found that localmeasurements usually give higher values (34). Because the localstiffness of the hydrogel was found to be unaffected by thegrooves, tube formation by cells was not the result of the differ-ences in local mechanical properties. However, cells degraded thehydrogel to migrate and form tubes, as shown by the steadydegradation of the hydrogel over 21 d with the cultivation of ar-tery and vein explants on top of the gel (Fig. S4).In addition to providing physical guidance for the sprouting

capillaries, the grooves also may influence the local concentra-tion of autocrine growth factors released from the endothelialcells actively participating in angiogenesis (Fig. 2C). A simplifiedmathematical model was derived to show that the presence ofgrooves and the groove width affected the local concentration ofVEGF165 assumed to be secreted as a point source from onesingle cell. The model values are not absolute, because severalsimplifying assumptions were used. However, the relative localVEGF concentrations, as normalized to the value for flat sub-strate at point (0, 0), show that the grooves can increase the localconcentration of the autocrine growth factors. Thus, one way thatthe grooves contribute to the differences observed in capillarygrowth is through the effect of physical barriers, as provided bythe PDMS groove walls, on the profiles of the autocrine growthfactor concentration. This process likely affects the optimalgroove width for the tube formation: Although the narrowergrooves concentrate the autocrine growth factors more, groovesthat are too narrow impose physical constraints on the migratingcells; thus the optimal groove width would be in the intermediaterange (e.g., 50 μm), as observed here (Fig. 1I).When an artery and a vein were placed at the opposite ends of

a PDMS stamp with Tβ4 hydrogel, the outgrowths from eachvessel followed the topographical cues, approaching each otherover time (Fig. 3A, days 7 and 14). By day 21, the capillary out-growths from the artery explants connected with those from thevein explants, forming an arteriovenous loop with capillariesaligned in the direction of the microgrooves (Fig. 3A). The av-erage branch length (Fig. 3C) was similar to the capillary length of∼600 μm in the native rat myocardium (35). The achieved branchdensity was much lower than the capillary density of >4,000capillaries/mm2 in the native mouse heart (36). Multiple engi-neered vascular structures could be stacked and cultivated foran additional period to increase capillary density. Additionally,hypoxia could be used to increase sprouting from the formedvascular bed. Another option for increasing capillary density issimply to increase groove density on the PDMS substrates bydecreasing the spacing between grooves.We next aimed to determine if we could accelerate the

sprouting rate further to achieve connection between the arteryand vein in a shorter time frame. For this purpose, we introducedthe angiogenic growth factors VEGF and hepatocyte growthfactor (HGF) to the culture medium at 100 ng/mL (37) or 20 ng/mL (38), respectively. The doses were chosen based on concen-trations previously used for angiogenesis assays (37, 38). VEGFpromoted the infiltration of ECs into collagen gels to form cap-illary-like structures (39) and induced sprouting of rat aortic rings(40). In addition, the increased sprouting in our system was cor-related with autocrine VEGF secretion (Fig. 1 B–F). HGF stim-ulates migration and proliferation of vascular ECs and acceleratestheir organization into capillary-like tubes in vitro (38). It alsoinduces the formation of blood vessels in vivo (38). Previousstudies demonstrated that HGF-induced vascularization in vivoinvolved the induction of VEGF expression in ECs (41).HGF or VEGF supplementation during explant cultivation

increased the branch length significantly at day 14 (Fig. 3C). Asa result, the vessel outgrowth rates were accelerated with HGFor VEGF, and connections between an artery and a vein could

be attained after 14 d of culture (Fig. 3 B–G). There was nodifference between VEGF and HGF in accelerating anastomosis,branch density, or branch length in this system (Fig. 3 B and C ).Our strategy was not species dependent. Similar results were

shown using rat femoral artery and vein explants and mousecardiac tissue explants (SI Results and Figs. S5 and S6), sug-gesting that it is not necessary to isolate specific arteries andveins. However, there are advantages in using vascular explantsrather than cardiac or other specific-tissue explants. First, arteriesand veins are more readily available for isolation. Second, theanchorage of the resulting capillary structure by parent vascularexplants could allow implantation and perfusion of the structure.The maximal branch density obtained using a single mouse or ratvascular explant was similar, ∼10/mm2 (Fig. 1 and Fig. S5). Car-diac explants, in contrast, gave a density of ∼30/mm2 (Fig. S6),likely because of the presence of a large number of blood vesselsin cardiac explants that could all sprout concurrently.Confocal microscopy indicated that the mouse capillary out-

growths had characteristics of developed vascular structures withopen lumens (Fig. 4 A–F and Movie S1). Transmission electronmicroscopy demonstrated the presence of lumens formed bycells in the peripheral position (Fig. 4H), as is typical of capil-laries, which are made of a single layer of ECs. Cells formingthe capillary outgrowths were positive for common endothelialmarkers, including CD31, VE-cadherin, and von Willebrand fac-tor (Fig. 4G, I, and J). Importantly, the formation of lumens by vonWillebrand factor-positive ECs was clearly evident (Fig. 4G) byconfocal microscopy z-stacks. Smooth muscle actin- and NG2-positive cells associated with these capillary outgrowths appearedin the peripheral position, typical of pericytes (Fig. S7). Smoothmuscle actin-positive cells alsowere found between the vessels andthe basement of the grooves (Fig. S7A).Because the microvascular bed originates from artery and vein

explants, we investigated the phenotype of the newly generatedsprouts. Ephrin-B2 and EphB4 selectively mark arterial andvenous ECs (42). As expected, outgrowths on the artery and veinsides of the engineered capillary structure were stained positivelyfor Ephrin-B2 and EphB4 respectively at day 21 (Fig. S8). In-terestingly, there also were Ephrin-B2+ cells within capillaries onthe vein side and EphB4+ cells on the artery side, suggesting anintegration of arterial and venous outgrowths. It has beendemonstrated that Ephrin-B2 and EphB4 can interact physicallyin growing blood vessels (43), as is consistent with the lack ofpreferential localization of these markers to either the venous orarterial side observed here. Ephrin-B2 is known to be up-regu-lated during pathological and physiological angiogenesis (42, 44).It also regulates the internalization and signaling activity ofVEGFR2 (45), the receptor of VEGF165 that was up-regulatedusing Tβ4 gels here.The capillary structure was removed easily without damage to

the morphology by peeling the hydrogel coating off the PDMSsubstrate (Fig. 4 K and L). More importantly, the engineeredvascular structures were perfusable with fluorescently labeleddextran (Movie S2).Using the same strategy, we engineered an oriented capillary

bed using human umbilical arteries and veins (Fig. 5 A and B).The capillary outgrowths were composed of CD31+ ECs (Fig. 5C).Connections between the parent arterial and venous explantswere made by day 21 (Fig. 5D) and the outgrowths containedopen lumens (Fig. 5E).To determine whether cardiac tissues with improved function

could be engineered using the established capillary structure,cardiomyocytes were cultivated for 7 d on the PDMS substratecontaining mouse vascular explants and the associated capillaryoutgrowths created after 14 d of culture with HGF supple-mentation (Fig. 6). Culture medium without HGF was used tocultivate the cardiomyocytes. This type of coculture preciselydefines the position of ECs and cardiomyocytes, with beating

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cardiomyocytes (Movie S3) found in parenchymal spaces aroundthe capillaries, as in the native heart (Fig. 6A). Cardiac tissuesgrown on vascular structures had a significantly lower excitationthreshold than those grown on control and Tβ4 gel-coated sub-strates without vascular structures, indicating improved func-tional properties (Fig. 6B). The maximum capture rates for thecardiac tissues were similar with or without vascular structures(Fig. 6C) and were comparable to values from our previousstudies (46). Troponin T staining demonstrated that car-diomyocytes grown on vascular structures contained better-organized sarcomeres than those grown on hydrogel coatingonly (Fig. 6 D and E). Increased Connexin-43 staining in car-diomyocytes cultivated on vascular structures (Fig. 6F) indicatedbetter cell–cell junctions.

DiscussionHere we focused on proving that the engineered vascular net-work is suitable for cultivation of parenchymal cells, lookingat the effects of the network alone. The phenotype of car-diomyocytes in this system could be enhanced further in futurestudies by applying electrical (46, 47) or mechanical (48) stim-ulation. EC cords previously led to synchronized contraction ofcardiomyocytes and increased Connexin-43 expression becauseof the improved survival and spreading of cardiomyocytes (49).In future studies, the vascular structures could be connected toa perfusion system to engineer more complex and metabolicallyactive cardiac tissues. This strategy can provide a native-likesupply of oxygen and nutrients while preventing the cells fromexperiencing hydrodynamic shear. Importantly, anchoring of the

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Fig. 3. Engineering of a connected microvascular bed using topographical cues and hydrogels with encapsulated Tβ4. (A) Fluorescence microscopy imagesshowing the time course of capillary outgrowths extending between a YFP+ mouse artery (on the right) and a vein (on the left) during in vitro cultivation onsubstrates with 50-μm grooves and hydrogel coating containing 1,500 ng encapsulated Tβ4. The connection was achieved at day 21. Arrows indicate locationsof artery and vein explants. (Scale bars, 200 μm.) The artery and vein explants were identified by the presence of bright fluorescence in the YFP mouseexplants. The artery was placed on the right and the vein was placed on the left of the PDMS substrate. The formed vascular structure was identified by the presenceof fluorescent tube-like structures extending between the artery and the vein. The progress of the formation of the vascular structures was monitored by fluo-rescence microscopy at the same location of the sample at different time points. (B–G) Soluble angiogenic growth factors enhance the outgrowths. (B) Branch densityat different time points when the explants were cultivated in culture mediumwith no growth factors or supplemented with VEGF or HGF. (C) Average branch lengthat different time points with no growth factors or supplemented with VEGF or HGF (P < 0.0001, two-way ANOVA; P < 0.05 for Tβ4 gel vs. Tβ4 gel + VEGF on day 14;P < 0.001 for Tβ4 gel vs. Tβ4 gel + HGF on day 14). An asterisk indicates a statistically significant difference between groups. (D–G) Fluorescence microscopy images ofYFP+ mouse artery and vein outgrowths with soluble growth factor supplementation. The connection was achieved at day 14 using VEGF supplementation (E) andHGF supplementation (G). (Scale bars, 100 μm.) These figures are composites of multiple images to illustrate the entire capillary bed structure. n = 6 per group.

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parent artery and vein explants provides a means for surgicallyconnecting the host vasculature to the engineered vascularstructure, thus potentially allowing the engineered tissue to beintegrated fully with the native tissue.A number of notable approaches have been used previously to

create branching microchannels via microfabrication in polymerssuch as polystyrene (50) and silk fibroin (51). Endothelializedmicrovessels also have been created in hydrogels such as collagen(19, 52, 53) and alginate (54). Embedding EC-laden alginatemicrofibers into smooth muscle cell-laden agar-based matrix hasbeen used for defined vascular cell coculture (55). Theseapproaches rely on providing defined areas for the attachment ofECs. In polymeric devices, the parenchymal space often is occu-pied by polymer, making cell seeding in the area around the mi-crovasculature difficult. Although hydrogels are more amenableto coculture, most are often fragile and difficult to handle, espe-

cially when fabricated as microfibers or long, hollow cylinders. Inthese approaches, the engineered vascular bed cannot be removedfrom the hydrogel or the microfluidic device and manipulated. Incontrast, we did not force cells to attach inmicrofabrication-definedattachment regions. Instead, our approach was inspired by nature:Topographical cues, hydrogel, and angiogenic factors were used tocreate a supportive niche for the controlled and directed sproutingof arterial and venous explants.Vascularization in vivo involves EC proliferation and angio-

genic sprouting in which tip cells acquire motile behavior underthe influence of VEGF growth factors (56). Anastomosis is ac-complished via the connection of extended cellular processesfollowed by lumen propagation through intercellular and in-tracellular vacuole fusion (57, 58). Here, we used the angiogenicfactor Tβ4 to enhance outgrowth cell density and increaseautocrine VEGF secretion, and topographical cues were criticalfor the formation and propagation of extended cellular processes.In this approach, ECs are not forced to attach and proliferate ona prefabricated structure. Instead they migrate and proliferateunder the direction of externally added and autocrine cytokines

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Fig. 4. Capillary outgrowths from mouse arteries and veins contained openlumens formed by endothelial cells and remained intact upon removal fromthe substrate. (A–F) Confocal microscopy showing lumens of YFP+ capillaryoutgrowths (n = 3). (A) Full view z-stacked x–y image. (Scale bar, 50 μm.) (B)Cross-sectional lumens in the y–z plane at the location indicated by thedotted line labeled “yz” in the z-stacked image. (Scale bar, 20 μm.) (C and D)Cross-sectional lumens in the x–z plane at the locations indicated by thedotted lines labeled “xz-1” (C) and “xz-2” (D) in the z-stacked image. (Scalebars, 50 μm.) (E and F) Longitudinal lumens at the locations indicated byorange boxes labeled “xy-a” (E) and “xy-b” (F) in the z-stacked image. (Scalebars, 10 μm.) (G) Confocal microscopy images of von Willebrand factorstaining (red) (n = 3). (i) Full view z-stacked x–y image. (Scale bar, 50 μm.) (ii)Cross-sectional lumens in the y–z plane at the location indicated by thedotted line labeled “yz” in the z-stacked image. (Scale bar, 20 μm.) (iii) Cross-sectional lumens in the x–z plane at the location indicated by the dotted linelabeled “xz” in the z-stacked image. (Scale bar, 50 μm.) (iv) Longitudinallumen at the location indicated by the orange box labeled “xy” in the z-stacked image. (Scale bar, 20 μm.) (H) Lumens formed by endothelial cells asshown by transmission electron microscopy (n = 3). (Scale bar, 2 μm.) (I and J)Representative immunostaining for (I) CD31 and (J) VE-cadherin (n = 3). Redindicates positive staining; blue indicates counterstaining of the nuclei withDAPI; arrows show localization of protein; dotted lines outline a capillaryoutgrowth. (Scale bars, 20 μm.) (K and L) The capillary network maintainedits structural integrity after removal from the PDMS substrate. Fluorescencemicroscopy images showing capillary outgrowths before (K) and after (L)removal (n = 3). (Scale bars,100 μm.)

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Fig. 5. Oriented and connected capillary outgrowths were formed from humanumbilical artery and vein explants. Day 21 microvascular bed on micropatternedPDMS substrates with a coating of collagen-chitosan hydrogel containing 1,500ng encapsulated Tβ4 (n = 5 per group). (A) Brightfield images of outgrowthson PDMS substrates with groove widths of 25, 50, or 100 μm. (Scale bars, 50 μm.)(B) Viable staining of outgrowths on PDMS substrates with groove widths of 25,50, or 100 μm. Green represents carboxyfluorescein diacetate (CFDA) staining oflive cells. (Scale bars, 50 μm.) (C) Fluorescence microscopy image showing positiveCD31 staining of endothelial cells that make up the capillary outgrowths onPDMS substrate with a groove width of 50 μm. Red indicates positive CD31staining; blue indicates counterstaining of nuclei with DAPI. Arrows indicateCD31-positive endothelial cells that make up the outgrowths. (Scale bar, 100 μm.)(D) Brightfield image showing connection of capillary outgrowths between theartery (on the right) and vein (on the left) explants on PDMS substrate with a50-μm groove width. Connected outgrowths are shown between pairs of dottedlines. Arrows indicate the parent explants. (Scale bar, 100 μm.) The figure isa composite of multiple images to illustrate the entire capillary bed structure. (E)Confocal microscopy image of CFDA-stained sample showing lumens of thecapillary outgrowths: (i) z-stacked x-y image; (ii) cross-sectional lumens in the y–zplane at the location indicated by the dotted line in the z-stacked image. (Scalebars, 20 μm.)

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and assemble into luminal structures directed by topographicalcues. Thus, the approach described here enables the engineeringof a prototype vascular network consisting of two branching ves-sels suitable for the cultivation of different parenchymal cell types.In summary, we have shown that Tβ4 in collagen-chitosan

hydrogels in conjunction with the application of topographicalcues guided endothelial outgrowths from an artery and a vein,enabling their organization into microvasculature. The resultingvascular bed was used to cultivate engineered vascularized car-diac tissues with improved function. The engineered microvas-cular bed was perfusable and removable from the PDMSsubstrate. Its functionality and ease of manipulation and thepresence of the flanking artery and vein overcome key limitationsof currently available approaches for in vitro vascularization.

Although we have demonstrated seeding and cultivation of car-diomyocytes as a parenchymal cell type here, the vascular bedis generic and could be used for the vascularization of othertissue types.

Materials and MethodsPreparation of Hydrogel. The hydrogel solutions were prepared bymixing 10×PBS, Tβ4 (stock solution of 1 mg/mL in distilled water) (HOR-275; ProSpec),chitosan in distilled water (75–90% deacetylation; PROTASAN, catalog no.G113; NovaMatrix), collagen I (4.08 mg/mL rat tail collagen I in acetic acid)(catalog no. 354236; BD Biosciences), and 1 N NaOH. The volume of 10× PBSin the hydrogel solution was 1/10th of the final volume, and the volume ofNaOH was 0.025 times the volume of collagen I. The volumes of Tβ4, chi-tosan, and collagen I were calculated based on the final desired concen-trations of 10 μg/mL (Encap100 group) or 150 μg/mL (Encap1500 group),

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Fig. 6. Engineering of vascularized cardiac tissue. Neonatal rat cardiomyocytes were seeded around day 14 microvascular bed and were cultivated for addi-tional 7 d (n = 6 per group). HGF was added to the culture medium during cultivation of capillary outgrowths. (A) A movie frame of the beating cardiac tissuefrom the group cultivated on Tβ4 gel with capillaries, showing the position of the cardiomyocytes relative to the capillary outgrowths that run parallel betweenthe parent explants. The asterisk indicates the location of the vascular explant. YFP+ capillary outgrowths are shown in green and are indicated by arrows.Cardiomyocytes are shown as brightfield with an added dark background to enhance contrast. (Scale bar, 200 μm.) (B and C) The functionality of engineeredcardiac tissues was evaluated by measuring (B) the excitation threshold (P = 0.0036, one-way ANOVA; P < 0.01 post hoc Tukey test for control vs. Tβ4 gel withcapillaries; P < 0.05 for Tβ4 gel vs. Tβ4 gel with capillaries) and (C) maximum capture rate (P = 0.2226, one-way ANOVA). An asterisk indicates a statisticallysignificant difference between groups. (D) Troponin T immunostaining shows striations in cardiomyocytes grown on a microvascular bed. The Tβ4 hydrogel aloneexhibits poorly developed cells. Troponin T staining is shown in red. Hoechst dye staining of the cell nuclei is shown in blue. (Scale bars, 20 μm.) (E) High-magnification confocal microscopy images showing Troponin T immunostaining (red). (Scale bars, 20 μm) (F) Images of Connexin-43 staining show more cell–celljunctions in the cardiac tissues grown on the microvascular bed. Arrows indicate positive punctate Connexin-43 staining. Connexin-43 staining is shown in red.Hoechst dye staining of the cell nuclei is shown in blue. (Scale bars, 50 μm.) Experimental groups include seeding of cardiomyocytes on PDMS substrates witha coating of Tβ4-free hydrogel (control gel), with a coating of Tβ4-encapsulated hydrogel (Tβ4 gel), and with day 14 capillary outgrowths on a coating of Tβ4-encapsulated hydrogel (Tβ4 gel with capillaries).

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1.25 mg/mL and 2.5 mg/mL, respectively. Distilled water was added insteadof Tβ4 in the control group.

Fabrication of Micropatterned PDMS Substrates. PDMS substrates were fab-ricated as previously described (59), using standard soft lithography. Theresulting substrates consisted of lanes 25, 50, or 100 μm in width and 65 μmin height. The substrates were cut into 5 × 5 mm squares and were fixed tothe bottom of individual wells. Hydrogel solution, 10 μL, was pipetted evenlyonto each PDMS substrate to coat the surface, followed by incubation at37 °C for 1 h. To show the influence of topography on the tube formationand organization, flat PDMS substrates coated with control or Tβ4 hydrogelwere added as controls.

Isolation and Cultivation of Explants. Cardiac tissues, thoracic arteries, andinferior vena cava were isolated from 5-wk-old YFP transgenic mice [129-Tg(CAGEYFP) 7AC5Nagy/J; Jackson Laboratory], and femoral arteries and veinswere isolated from Sprague–Dawley rats according to a protocol approvedby the University of Toronto Committee on Animal Care. Human umbilicalarteries and veins were kind gifts from John Davies at Tissue RegenerationTherapeutics (Toronto, ON, Canada). For studies with vascular explants, onepiece of artery and one piece of vein were placed on the two sides of thePDMS substrate, ∼0.5–1 mm apart. For studies with cardiac explants, the toppart of the heart containing the dense vasculature was used. As a control todetermine the importance of the ECM interactions, the explants were cul-tivated on PDMS substrates with 50-μm grooves without hydrogel coat-ing. In this case, the YFP mouse vascular explants were pinned down at thetwo ends of the PDMS substrates, and soluble Tβ4 was added to the culturemedium. The amount of Tβ4 added to the medium was based on our pre-vious study (22), in which the average rate of Tβ4 release from Tβ4-encap-sulated hydrogel was found to be ∼37.5 ng/mL/d over 28 d. The mouseexplants were cultivated in culture medium consisting of DMEM with 1%(vol/vol) penicillin/streptomycin, 1% (vol/vol) Hepes, and 15% (vol/vol) FBS.In some cases, culture medium was supplemented with 20 ng/mL HGF or100 ng/mL VEGF. The rat explants were cultivated in culture medium con-sisting of DMEM with 1% (vol/vol) penicillin/streptomycin, 1% (vol/vol)Hepes, and 10% (vol/vol) FBS. The human explants were cultivated in Clo-netics Endothelial Cell Growth Medium-2 (EGM2) with BulletKit (catalog CC-3162; Lonza), suitable for the culture of human endothelial cells.

Image Analysis. At different time points (days 7, 9, 11, 14, 21), the sampleswere imaged under optical or fluorescence microscope (Olympus IX2-UCB),and the images were analyzed using ImageJ (SI Materials and Methods).

Immunohistochemical Analysis. The staining was carried out as describedpreviously (22). Briefly, the samples were fixed in 10% formalin and stainedwith specific antibodies against CD31 (1:50; Abcam), VE-cadherin (1:50;Abcam), von Willebrand factor (1:100; Abcam), SMA (1:100; Abcam), NG2Chondroitin Sulfate Proteoglycan (1:200; Millipore), EphB4 (1:50; Hycult Bio-tech), Ephrin-B2 (1:100; GenScript), Troponin T (1:100; Fisher), and connexin-43 (1:100; Abcam) followed by rhodamine- or Texas red-labeled secondaryantibodies (1:100; rhodamine goat anti-mouse, Jackson ImmunoResearch;rhodamine goat anti-rat, Jackson ImmunoResearch; or Texas red goat anti-rabbit, Abcam). Nuclei were stained with Hoechst dye or DAPI (1:100).

Dextran Perfusion. Rhodamine-labeled dextran (Invitrogen) in distilled water,200 μg/mL, was injected into the artery explant using a 30-G 1/2-in needle.Fluorescence microscopy was used to capture the perfusion of dextranthrough the capillary outgrowths.

Transmission Electron Microscopy. After 21-d cultivation, samples were fixedin glutaraldehyde for 1 h and washed in PBS. Osmium tetroxide was added tothe samples for 1 h. The samples then were serially dehydrated in 70, 90, and

100% ethanol. After dehydration, the samples were imbedded in resin,sectioned, mounted on glass slides, and stained.

VEGF ELISA. Culture medium was collected at 12 and 14 d of explant culti-vation on hydrogel. VEGF ELISA was performed on the culture mediumsamples using the Murine VEGF ELISA Development Kit (Peprotech) to de-termine the VEGF secretion between days 12 and 14. The time points werechosen based on the plateau in cell outgrowth and the initiation of themajority of tube formation.

Seeding Cardiomyocytes onto Capillary Structures. After mouse arterial andvenous explants were cultured for 14 d with HGF supplementation to formconnected capillary outgrowths, 100,000 neonatal rat cardiomyocytes wereseeded on the capillary structure for each sample (SI Materials and Methods).The cardiomyocytes were cultivated for 7 d in 1 mL culture medium con-sisting of DMEM with 1% penicillin/streptomycin, 1% Hepes, and 15% FBS.At the end of the 7-d cultivation, the engineered cardiac tissues were re-moved from the PDMS substrates, and their functional properties weremeasured (SI Materials and Methods) as previously described (46).

Atomic Force Microscopy. PDMS substrates without grooves or with 25-, 50-,and 100-μm–wide grooves were fabricated, coated with plain collagen-chi-tosan hydrogel solution, and incubated at 37 °C for 1 h for gelation. Culturemedium was added to the dish to immerse the PDMS substrates fully inliquid. Local mechanical stiffness of the hydrogel coating was measured fora selected 50 × 50 μm area, spanning a part or all of one groove/ridge re-gion, on each sample using an atomic force microscope (Bioscope Catalyst;Bruker) under force volume mode. The pyramidal tip used (MSCT-D; Bruker)had a semivertical side angle of 17.5°, a typical height of 5–8 μm, anda nominal cantilever spring constant of 0.03 N/m. The measured local me-chanical stiffness values were averaged for each substrate type, because nopattern was found in stiffness with regards to the location of measurementon the substrates.

Mathematical Modeling of Topographical Effects on Local ConcentrationProfiles of Autocrine VEGF. A simplified mathematical model was formu-lated to examine the potential differences in local autocrine growth factorconcentrations caused by the presence of PDMS grooves. A representative 2Dsteady-state model of VEGF165 diffusing out of a single cell was solved usingfinite element model software, COMSOL Multiphysics 3.5. The solver usedwas DIRECT(UMFPACK). All initial concentrations were equal to 0. Becausethe purpose of this model was simply to investigate the effect of geometryon local VEGF concentration, several complexities (e.g., kinetic receptor andECM binding, basement membrane formation, proteolysis, internalization,multiple VEGF splice isoforms) were ignored. The details for the mathe-matical modeling are given in SI Materials and Methods.

Statistical Analysis. Statistical analysis was performed with Graphpad Prism5.0 software. One-way ANOVA with post hoc Tukey tests and two-wayANOVA with Bonferroni posttests were used to compare the differencesamong groups, with P < 0.05 considered statistically significant. The data arepresented as mean ± SE.

ACKNOWLEDGMENTS. We thank Ms. Anne Hsieh for help in microfabrica-tion, Dr. John Davies for human umbilical cords, and Drs. Yu Sun and CraigSimmons for use of the atomic force microscope. This work was funded byNatural Sciences and Engineering Research Council (NSERC) Strategic GrantSTPGP 381002-09, NSERC-Canadian Institutes of Health Research Collab-orative Health Research Grant CHRPJ 385981-10, NSERC Discovery GrantRGPIN 326982-10, and Discovery Accelerator Supplement RGPAS 396125-10.L.L.Y.C. is the recipient of an NSERC Canada Graduate Scholarship andQueen Elizabeth II Graduate Scholarship in Science and Technology. M.R.is the recipient of a McLean Award.

1. Raghavan S, Nelson CM, Baranski JD, Lim E, Chen CS (2010) Geometrically controlled

endothelial tubulogenesis in micropatterned gels. Tissue Eng Part A 16(7):2255–2263.2. Rakusan K, Korecky B (1982) The effect of growth and aging on functional capillary

supply of the rat heart. Growth 46(3):275–281.3. Asakawa N, et al. (2010) Pre-vascularization of in vitro three-dimensional tissues

created by cell sheet engineering. Biomaterials 31(14):3903–3909.4. Takahashi T, et al. (1999) Ischemia- and cytokine-induced mobilization of bone marrow-

derived endothelial progenitor cells for neovascularization. Nat Med 5(4):434–438.5. Morritt AN, et al. (2007) Cardiac tissue engineering in an in vivo vascularized chamber.

Circulation 115(3):353–360.6. Madden LR, et al. (2010) Proangiogenic scaffolds as functional templates for cardiac

tissue engineering. Proc Natl Acad Sci USA 107(34):15211–15216.

7. Miyagi Y, et al. (2011) Biodegradable collagen patch with covalently immobilized

VEGF for myocardial repair. Biomaterials 32(5):1280–1290.8. Chiu LL, Radisic M (2010) Scaffolds with covalently immobilized VEGF and

Angiopoietin-1 for vascularization of engineered tissues. Biomaterials 31(2):

226–241.9. Chiu LL, Weisel RD, Li RK, Radisic M (2011) Defining conditions for covalent

immobilization of angiogenic growth factors onto scaffolds for tissue engineering.

J Tissue Eng Regen Med 5(1):69–84.10. Dvir T, et al. (2009) Prevascularization of cardiac patch on the omentum improves its

therapeutic outcome. Proc Natl Acad Sci USA 106(35):14990–14995.11. Richardson TP, Peters MC, Ennett AB, Mooney DJ (2001) Polymeric system for dual

growth factor delivery. Nat Biotechnol 19(11):1029–1034.

E3422 | www.pnas.org/cgi/doi/10.1073/pnas.1210580109 Chiu et al.

Dow

nloa

ded

by g

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12. Chen X, et al. (2009) Prevascularization of a fibrin-based tissue construct acceleratesthe formation of functional anastomosis with host vasculature. Tissue Eng Part A15(6):1363–1371.

13. Smart N, et al. (2010) Thymosin beta4 facilitates epicardial neovascularization of theinjured adult heart. Ann N Y Acad Sci 1194:97–104.

14. Smart N, et al. (2007) Thymosin beta4 induces adult epicardial progenitormobilization and neovascularization. Nature 445(7124):177–182.

15. Co CC, Wang YC, Ho CC (2005) Biocompatible micropatterning of two different celltypes. J Am Chem Soc 127(6):1598–1599.

16. Moon JJ, Hahn MS, Kim I, Nsiah BA, West JL (2009) Micropatterning of poly(ethyleneglycol) diacrylate hydrogels with biomolecules to regulate and guide endothelialmorphogenesis. Tissue Eng Part A 15(3):579–585.

17. Dike LE, et al. (1999) Geometric control of switching between growth, apoptosis, anddifferentiation during angiogenesis using micropatterned substrates. In Vitro Cell DevBiol Anim 35(8):441–448.

18. Levenberg S, et al. (2005) Engineering vascularized skeletal muscle tissue. NatBiotechnol 23(7):879–884.

19. Zheng Y, et al. (2012) In vitro microvessels for the study of angiogenesis andthrombosis. Proc Natl Acad Sci USA 109(24):9342–9347.

20. Bock-Marquette I, Saxena A, White MD, Dimaio JM, Srivastava D (2004) Thymosinbeta4 activates integrin-linked kinase and promotes cardiac cell migration, survivaland cardiac repair. Nature 432(7016):466–472.

21. Lin CC, Metters AT (2006) Hydrogels in controlled release formulations: Networkdesign and mathematical modeling. Adv Drug Deliv Rev 58(12-13):1379–1408.

22. Chiu LL, Radisic M (2011) Controlled release of thymosin β4 using collagen-chitosancomposite hydrogels promotes epicardial cell migration and angiogenesis. J ControlRelease 155(3):376–385.

23. Grant DS, et al. (1999) Thymosin beta4 enhances endothelial cell differentiation andangiogenesis. Angiogenesis 3(2):125–135.

24. Sosne G, Qiu P, Goldstein AL, Wheater M (2010) Biological activities of thymosin beta4defined by active sites in short peptide sequences. FASEB J 24(7):2144–2151.

25. Morrison RF, Seidel ER (1995) Vascular endothelial cell proliferation: Regulation ofcellular polyamines. Cardiovasc Res 29(6):841–847.

26. Ma N, et al. (2005) Human cord blood cells induce angiogenesis following myocardialinfarction in NOD/scid-mice. Cardiovasc Res 66(1):45–54.

27. Cross MJ, Dixelius J, Matsumoto T, Claesson-Welsh L (2003) VEGF-receptor signaltransduction. Trends Biochem Sci 28(9):488–494.

28. Rini BI, Small EJ (2005) Biology and clinical development of vascular endothelialgrowth factor-targeted therapy in renal cell carcinoma. J Clin Oncol 23(5):1028–1043.

29. Olsson AK, Dimberg A, Kreuger J, Claesson-Welsh L (2006) VEGF receptor signalling -in control of vascular function. Nat Rev Mol Cell Biol 7(5):359–371.

30. Gille H, et al. (2001) Analysis of biological effects and signaling properties of Flt-1(VEGFR-1) and KDR (VEGFR-2). A reassessment using novel receptor-specific vascularendothelial growth factor mutants. J Biol Chem 276(5):3222–3230.

31. Yang S, et al. (2001) Vascular endothelial cell growth factor-driven endothelial tubeformation is mediated by vascular endothelial cell growth factor receptor-2, a kinaseinsert domain-containing receptor. Arterioscler Thromb Vasc Biol 21(12):1934–1940.

32. Ilan N, Mahooti S, Madri JA (1998) Distinct signal transduction pathways are utilizedduring the tube formation and survival phases of in vitro angiogenesis. J Cell Sci 111(Pt 24):3621–3631.

33. Tanimoto T, Jin ZG, Berk BC (2002) Transactivation of vascular endothelial growthfactor (VEGF) receptor Flk-1/KDR is involved in sphingosine 1-phosphate-stimulatedphosphorylation of Akt and endothelial nitric-oxide synthase (eNOS). J Biol Chem 277(45):42997–43001.

34. Song J, Tranchida D, Vancso GJ (2008) Contact mechanics of UV/ozone-treated PDMSby AFM and JKR testing: Mechanical performance from nano- to micrometer lengthscales. Macromolecules 41(18):6757–6762.

35. Batra S, Rakusan K (1992) Capillary length, tortuosity, and spacing in rat myocardiumduring cardiac cycle. Am J Physiol 263(5 Pt 2):H1369–H1376.

36. Kubis N, et al. (2002) Decreased arteriolar density in endothelial nitric oxide synthaseknockout mice is due to hypertension, not to the constitutive defect in endothelialnitric oxide synthase enzyme. J Hypertens 20(2):273–280.

37. Carretero-Ortega J, et al. (2010) Phosphatidylinositol 3,4,5-triphosphate-dependent

Rac exchanger 1 (P-Rex-1), a guanine nucleotide exchange factor for Rac, mediates

angiogenic responses to stromal cell-derived factor-1/chemokine stromal cell derived

factor-1 (SDF-1/CXCL-12) linked to Rac activation, endothelial cell migration, and in

vitro angiogenesis. Mol Pharmacol 77(3):435–442.38. Grant DS, et al. (1993) Scatter factor induces blood vessel formation in vivo. Proc Natl

Acad Sci USA 90(5):1937–1941.39. Pepper MS, Ferrara N, Orci L, Montesano R (1992) Potent synergism between vascular

endothelial growth factor and basic fibroblast growth factor in the induction of

angiogenesis in vitro. Biochem Biophys Res Commun 189(2):824–831.40. Nicosia RF, Nicosia SV, Smith M (1994) Vascular endothelial growth factor, platelet-

derived growth factor, and insulin-like growth factor-1 promote rat aortic angiogenesis

in vitro. Am J Pathol 145(5):1023–1029.41. Toyoda M, et al. (2001) Overexpression of hepatocyte growth factor/scatter factor

promotes vascularization and granulation tissue formation in vivo. FEBS Lett 509(1):

95–100.42. Gale NW, et al. (2001) Ephrin-B2 selectively marks arterial vessels and neovascularization

sites in the adult, with expression in both endothelial and smooth-muscle cells. Dev Biol

230(2):151–160.43. Taylor AC, Murfee WL, Peirce SM (2007) EphB4 expression along adult rat microvascular

networks: EphB4 is more than a venous specific marker. Microcirculation 14(3):253–267.44. Shin D, et al. (2001) Expression of ephrinB2 identifies a stable genetic difference

between arterial and venous vascular smooth muscle as well as endothelial cells, and

marks subsets of microvessels at sites of adult neovascularization. Dev Biol 230(2):

139–150.45. Sawamiphak S, et al. (2010) Ephrin-B2 regulates VEGFR2 function in developmental

and tumour angiogenesis. Nature 465(7297):487–491.46. Chiu LL, Iyer RK, King JP, Radisic M (2011) Biphasic electrical field stimulation aids in

tissue engineering of multicell-type cardiac organoids. Tissue Eng Part A 17(11-12):

1465–1477.47. Radisic M, et al. (2004) Functional assembly of engineered myocardium by electrical

stimulation of cardiac myocytes cultured on scaffolds. Proc Natl Acad Sci USA 101(52):

18129–18134.48. Zimmermann WH, et al. (2002) Tissue engineering of a differentiated cardiac muscle

construct. Circ Res 90(2):223–230.49. Narmoneva DA, Vukmirovic R, Davis ME, Kamm RD, Lee RT (2004) Endothelial cells

promote cardiac myocyte survival and spatial reorganization: Implications for cardiac

regeneration. Circulation 110(8):962–968.50. Borenstein JT, et al. (2010) Functional endothelialized microvascular networks with

circular cross-sections in a tissue culture substrate. Biomed Microdevices 12(1):71–79.51. Bettinger CJ, et al. (2007) Silk fibroin microfluidic devices. Advanced Materials 19(5):

2847–2850.52. Price GM, et al. (2010) Effect of mechanical factors on the function of engineered

human blood microvessels in microfluidic collagen gels. Biomaterials 31(24):6182–6189.53. Gillette BM, et al. (2008) In situ collagen assembly for integrating microfabricated

three-dimensional cell-seeded matrices. Nat Mater 7(8):636–640.54. Choi NW, et al. (2007) Microfluidic scaffolds for tissue engineering. Nat Mater 6(11):

908–915.55. Lee KH, Shin SJ, Park Y, Lee SH (2009) Synthesis of cell-laden alginate hollow fibers

using microfluidic chips and microvascularized tissue-engineering applications. Small

5(11):1264–1268.56. Wang Y, et al. (2010) Ephrin-B2 controls VEGF-induced angiogenesis and lymphan-

giogenesis. Nature 465(7297):483–486.57. Kamei M, et al. (2006) Endothelial tubes assemble from intracellular vacuoles in vivo.

Nature 442(7101):453–456.58. Lubarsky B, Krasnow MA (2003) Tube morphogenesis: Making and shaping biological

tubes. Cell 112(1):19–28.59. Khademhosseini A, et al. (2007) Microfluidic patterning for fabrication of contractile

cardiac organoids. Biomed Microdevices 9(2):149–157.

Chiu et al. PNAS | Published online November 26, 2012 | E3423

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