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Tissue Engineering of the Microvasculature

Joe Tien

10.1002/cphy.c180037

Source: Volume 9, Issue 3, July 2019

Published online: June 2019

Full Article on Wiley Online Library



ABSTRACT

The ability to generate new microvessels in desired numbers and at desired locations has been a long‐sought goal in vascular medicine, engineering, and biology. Historically, the need to revascularize ischemic tissues nonsurgically (so‐called therapeutic vascularization) served as the main driving force for the development of new methods of vascular growth. More recently, vascularization of engineered tissues and the generation of vascularized microphysiological systems have provided additional targets for these methods, and have required adaptation of therapeutic vascularization to biomaterial scaffolds and to microscale devices. Three complementary strategies have been investigated to engineer microvasculature: angiogenesis (the sprouting of existing vessels), vasculogenesis (the coalescence of adult or progenitor cells into vessels), and microfluidics (the vascularization of scaffolds that possess the open geometry of microvascular networks). Over the past several decades, vascularization techniques have grown tremendously in sophistication, from the crude implantation of arteries into myocardial tunnels by Vineberg in the 1940s, to the current use of micropatterning techniques to control the exact shape and placement of vessels within a scaffold. This review provides a broad historical view of methods to engineer the microvasculature, and offers a common framework for organizing and analyzing the numerous studies in this area of tissue engineering and regenerative medicine. © 2019 American Physiological Society. Compr Physiol 9:1155‐1212, 2019.

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Figure 1. Figure 1. Vascularization strategies based on (A) angiogenesis into a graft, (B) vasculogenesis within a graft, and (C) direct seeding and perfusion of a microfluidic scaffold. GF, growth factor; ECM, extracellular matrix. Adapted, with permission, from (584).
Figure 2. Figure 2. Quantitative metrics of microvascular physiology. (A) Calculation of endothelial hydraulic conductivity relies on measurement of filtration speed as a function of vascular pressure. Reprinted with permission from (313). (B) Calculation of solute permeability relies on measurement of solute accumulation over time. Reproduced, with permission, from (227).
Figure 3. Figure 3. Binding of VEGFs to their receptors, the VEGFRs and neuropilins. Reproduced, with permission, from (247).
Figure 4. Figure 4. Diagram of the Vineberg procedure for vascularizing ischemic myocardium, using a carotid artery implant. RV, right ventricle; LV, left ventricle. Reproduced, with permission, from (479).
Figure 5. Figure 5. Dose‐dependent collateral growth 2 months after implantation of FGF2‐loaded heparin/alginate beads around occluded coronary arteries in the ischemic pig heart. bFGF, basic fibroblast growth factor (FGF2). Reproduced, with permission, from (350).
Figure 6. Figure 6. Density of vascular ingrowth into polyHEMA implants after 1 month. Reproduced, with permission, from (361).
Figure 7. Figure 7. Cellularity of fibrovascular ingrowth into porous PTFE chambers in the presence of various growth factors after 10 days. Reproduced, with permission, from (541).
Figure 8. Figure 8. Number of viable hepatocytes in PLGA sponge implants as a function of time. Hepatocytes were added directly to scaffolds before implantation; scaffolds were not prevascularized. Reproduced, with permission, from (388).
Figure 9. Figure 9. Strategy for vascularization of grafts from a surgically constructed arteriovenous (AV) loop. Angiogenesis from the loop invades into overlying tissue. A, artery; V, vein. Reproduced, with permission, from (137).
Figure 10. Figure 10. Generation of centimeter‐scale tissue with an arteriovenous (AV) loop in a polycarbonate chamber. (A) AV loop overlaid on a polycarbonate base. (B) Addition of rat cardiomyocytes and Matrigel around the AV loop. (C) Explant of pedicled myocardium that formed by 4 weeks. Scale bar refers to 1 cm. Reproduced, with permission, from (394).
Figure 11. Figure 11. VEGF‐driven angiogenesis into collagen gels in microfluidic devices. Treatment of devices with poly‐D‐lysine (PDL) enabled ECs to sprout from a monolayer toward a VEGF source. Arrows denote the direction of VEGF transport. Scale bars refer to 100 μm. EGM2mv, endothelial cell growth media. Reproduced, with permission, from (92).
Figure 12. Figure 12. Derivation and characterization of EPCs. (A) Spindle‐shaped EPCs that were derived from 1‐week‐old cultures of CD34‐enriched peripheral blood‐derived mononuclear cells. Reproduced with permission from (23). (B) Flow cytometry of EPCs and monocytes. EPCs express blood cell markers, including CD45 and the monocyte activation marker CD11c. Adapted, with permission, from (459).
Figure 13. Figure 13. Derivation and characterization of EOCs. (Left) Colony of cord blood‐derived EOCs that emerged after mononuclear cells were cultured for 9 days. (Right) Flow cytometry of EOCs. EOCs express EC markers, such as CD31 and CD144 (VE‐cadherin), but do not express markers of blood cells, such as CD45 and the macrophage/monocyte marker CD14. Adapted, with permission, from (232).
Figure 14. Figure 14. Incorporation of bone marrow‐derived, lacZ‐expressing EPCs at sites of ischemia. Sections were costained by X‐gal (blue) and for lectin (A) and CD31 (B). Scale bar refers to 25 μm. Reproduced, with permission, from (22).
Figure 15. Figure 15. Vascular mimicry in uveal melanoma. Red blood cells (top, arrowheads) fill vessel‐like structures that do not appear to be lined by ECs. Reproduced, with permission, from (358).
Figure 16. Figure 16. Fine structure of vessels that formed from normal and Bcl2‐expressing HUVECs in collagen‐fibronectin gels 1 month after implantation. (A) Implants of normal HUVECs. (B, C) Implants of Bcl2‐expressing HUVECs. The lumens are perfused with red blood cells (RBC); asterisks denote mural cells. Reproduced, with permission, from (494).
Figure 17. Figure 17. Organization of human microvascular ECs into blood and lymphatic vessels in a dermal graft after 2 weeks. Immunostains are shown for CD31, blood vessel marker laminin (Lam1,2), and lymphatic markers LYVE‐1 and podoplanin (hPDN). Scale bars refer to 100 μm. Adapted, with permission, from (360).
Figure 18. Figure 18. Transformation of endothelial aggregates into perfused vessels. (A) Random self‐organized HUVEC networks and patterned HUVEC cords, before and after implantation. (B) Two‐week‐old implants after perfusion with species‐specific lectins. Vessels in implants consist of ECs that are derived from graft (human, red) and host (mouse, green). Scale bars refer to 250 μm (A, upper left), 500 μm (A, upper middle and upper right), 100 μm (A, lower), and 150 μm (B). Reproduced, with permission, from (38).
Figure 19. Figure 19. Increase in perfused vascular density over time in subcutaneous polymer implants that contained adipose‐derived microvascular fragments (black circles) or that did not (clear circles). (Left) Vascular density near the surface of implants. (Right) Vascular density at the center of implants. Adapted, with permission, from (316).
Figure 20. Figure 20. Vascularized myocardial constructs that contained human cardiomyocytes, HUVECs, and mouse fibroblasts in a PLA/PLGA sponge after 2 weeks. (Left) Immunostain for human CD31 (red) and von Willebrand factor (green). (Right) Immunostain for human CD31 (brown). Reproduced, with permission, from (332).
Figure 21. Figure 21. Perfused vascular networks in microscale fibrin gels within microfluidic devices. Fibroblasts, in the same or separate gel, were included to promote vasculogenesis. In (B), the resulting networks were perfused with a solution of fluorescent dextran. Reproduced, with permission, from (277,395).
Figure 22. Figure 22. Branching vascular‐like pattern that was etched into a silicon wafer. Such patterns could be seeded with ECs to generate patterned cultures. Feature widths were on the order of ∼10 μm. Adapted, with permission, from (249).
Figure 23. Figure 23. Vascularization of a microfluidic type I collagen scaffold that was formed by combining a micropatterned and planar gel. Both en face and reconstructed 3D views of HUVEC‐seeded structures are shown. Scale bar refers to 100 μm. Reproduced, with permission, from (677).
Figure 24. Figure 24. Vascularization of a microfluidic type I collagen scaffold that was formed around a removable needle. (Top) Unseeded scaffold. (Bottom) Scaffold that was seeded with HUVECs. Insets show cross‐sectional views. Scale bar refers to 100 μm. Reproduced, with permission, from (90).
Figure 25. Figure 25. Vascularization of a microfluidic fibrin scaffold that was formed around a sacrificial 3D‐printed network of sugar‐based fibers. Channels were seeded with HUVECs (red), and the scaffold bulk contained 10T1/2 mouse fibroblasts (green). Scale bar refers to 1 mm. Reproduced, with permission, from (382).
Figure 26. Figure 26. Vascularization of a microfluidic polyethylene glycol gel that was patterned by photodegradation. (Left) Visualization of interconnected channels by perfusion with fluorescent dextran. (Right) 3D confocal reconstruction of a vascularized channel that was stained for ZO‐1 (green). Reproduced, with permission, from (196).
Figure 27. Figure 27. Perfusion‐decellularization of whole hearts preserves the vascular architecture. (A) Corrosion casts of native and decellularized hearts. Scale bars refer to 1 mm (top) and 250 μm (bottom). (B) Transplanted decellularized heart before and after reestablishment of blood flow. The recipient animal was heparinized to minimize thrombosis. Reproduced, with permission, from (424).
Figure 28. Figure 28. Physical mechanisms for stable vascularization of micro‐fluidic scaffolds. Maintenance of vascular adhesion can be viewed as (A) a balance of outward (stabilizing) and inward (destabilizing) stresses, including pressures, contractile stress, and adhesion stress, or as (B) a balance of adhesion (stabilizing) and stored elastic (destabilizing) energies. Adapted, with permission, from (643).


Figure 1. Vascularization strategies based on (A) angiogenesis into a graft, (B) vasculogenesis within a graft, and (C) direct seeding and perfusion of a microfluidic scaffold. GF, growth factor; ECM, extracellular matrix. Adapted, with permission, from (584).


Figure 2. Quantitative metrics of microvascular physiology. (A) Calculation of endothelial hydraulic conductivity relies on measurement of filtration speed as a function of vascular pressure. Reprinted with permission from (313). (B) Calculation of solute permeability relies on measurement of solute accumulation over time. Reproduced, with permission, from (227).


Figure 3. Binding of VEGFs to their receptors, the VEGFRs and neuropilins. Reproduced, with permission, from (247).


Figure 4. Diagram of the Vineberg procedure for vascularizing ischemic myocardium, using a carotid artery implant. RV, right ventricle; LV, left ventricle. Reproduced, with permission, from (479).


Figure 5. Dose‐dependent collateral growth 2 months after implantation of FGF2‐loaded heparin/alginate beads around occluded coronary arteries in the ischemic pig heart. bFGF, basic fibroblast growth factor (FGF2). Reproduced, with permission, from (350).


Figure 6. Density of vascular ingrowth into polyHEMA implants after 1 month. Reproduced, with permission, from (361).


Figure 7. Cellularity of fibrovascular ingrowth into porous PTFE chambers in the presence of various growth factors after 10 days. Reproduced, with permission, from (541).


Figure 8. Number of viable hepatocytes in PLGA sponge implants as a function of time. Hepatocytes were added directly to scaffolds before implantation; scaffolds were not prevascularized. Reproduced, with permission, from (388).


Figure 9. Strategy for vascularization of grafts from a surgically constructed arteriovenous (AV) loop. Angiogenesis from the loop invades into overlying tissue. A, artery; V, vein. Reproduced, with permission, from (137).


Figure 10. Generation of centimeter‐scale tissue with an arteriovenous (AV) loop in a polycarbonate chamber. (A) AV loop overlaid on a polycarbonate base. (B) Addition of rat cardiomyocytes and Matrigel around the AV loop. (C) Explant of pedicled myocardium that formed by 4 weeks. Scale bar refers to 1 cm. Reproduced, with permission, from (394).


Figure 11. VEGF‐driven angiogenesis into collagen gels in microfluidic devices. Treatment of devices with poly‐D‐lysine (PDL) enabled ECs to sprout from a monolayer toward a VEGF source. Arrows denote the direction of VEGF transport. Scale bars refer to 100 μm. EGM2mv, endothelial cell growth media. Reproduced, with permission, from (92).


Figure 12. Derivation and characterization of EPCs. (A) Spindle‐shaped EPCs that were derived from 1‐week‐old cultures of CD34‐enriched peripheral blood‐derived mononuclear cells. Reproduced with permission from (23). (B) Flow cytometry of EPCs and monocytes. EPCs express blood cell markers, including CD45 and the monocyte activation marker CD11c. Adapted, with permission, from (459).


Figure 13. Derivation and characterization of EOCs. (Left) Colony of cord blood‐derived EOCs that emerged after mononuclear cells were cultured for 9 days. (Right) Flow cytometry of EOCs. EOCs express EC markers, such as CD31 and CD144 (VE‐cadherin), but do not express markers of blood cells, such as CD45 and the macrophage/monocyte marker CD14. Adapted, with permission, from (232).


Figure 14. Incorporation of bone marrow‐derived, lacZ‐expressing EPCs at sites of ischemia. Sections were costained by X‐gal (blue) and for lectin (A) and CD31 (B). Scale bar refers to 25 μm. Reproduced, with permission, from (22).


Figure 15. Vascular mimicry in uveal melanoma. Red blood cells (top, arrowheads) fill vessel‐like structures that do not appear to be lined by ECs. Reproduced, with permission, from (358).


Figure 16. Fine structure of vessels that formed from normal and Bcl2‐expressing HUVECs in collagen‐fibronectin gels 1 month after implantation. (A) Implants of normal HUVECs. (B, C) Implants of Bcl2‐expressing HUVECs. The lumens are perfused with red blood cells (RBC); asterisks denote mural cells. Reproduced, with permission, from (494).


Figure 17. Organization of human microvascular ECs into blood and lymphatic vessels in a dermal graft after 2 weeks. Immunostains are shown for CD31, blood vessel marker laminin (Lam1,2), and lymphatic markers LYVE‐1 and podoplanin (hPDN). Scale bars refer to 100 μm. Adapted, with permission, from (360).


Figure 18. Transformation of endothelial aggregates into perfused vessels. (A) Random self‐organized HUVEC networks and patterned HUVEC cords, before and after implantation. (B) Two‐week‐old implants after perfusion with species‐specific lectins. Vessels in implants consist of ECs that are derived from graft (human, red) and host (mouse, green). Scale bars refer to 250 μm (A, upper left), 500 μm (A, upper middle and upper right), 100 μm (A, lower), and 150 μm (B). Reproduced, with permission, from (38).


Figure 19. Increase in perfused vascular density over time in subcutaneous polymer implants that contained adipose‐derived microvascular fragments (black circles) or that did not (clear circles). (Left) Vascular density near the surface of implants. (Right) Vascular density at the center of implants. Adapted, with permission, from (316).


Figure 20. Vascularized myocardial constructs that contained human cardiomyocytes, HUVECs, and mouse fibroblasts in a PLA/PLGA sponge after 2 weeks. (Left) Immunostain for human CD31 (red) and von Willebrand factor (green). (Right) Immunostain for human CD31 (brown). Reproduced, with permission, from (332).


Figure 21. Perfused vascular networks in microscale fibrin gels within microfluidic devices. Fibroblasts, in the same or separate gel, were included to promote vasculogenesis. In (B), the resulting networks were perfused with a solution of fluorescent dextran. Reproduced, with permission, from (277,395).


Figure 22. Branching vascular‐like pattern that was etched into a silicon wafer. Such patterns could be seeded with ECs to generate patterned cultures. Feature widths were on the order of ∼10 μm. Adapted, with permission, from (249).


Figure 23. Vascularization of a microfluidic type I collagen scaffold that was formed by combining a micropatterned and planar gel. Both en face and reconstructed 3D views of HUVEC‐seeded structures are shown. Scale bar refers to 100 μm. Reproduced, with permission, from (677).


Figure 24. Vascularization of a microfluidic type I collagen scaffold that was formed around a removable needle. (Top) Unseeded scaffold. (Bottom) Scaffold that was seeded with HUVECs. Insets show cross‐sectional views. Scale bar refers to 100 μm. Reproduced, with permission, from (90).


Figure 25. Vascularization of a microfluidic fibrin scaffold that was formed around a sacrificial 3D‐printed network of sugar‐based fibers. Channels were seeded with HUVECs (red), and the scaffold bulk contained 10T1/2 mouse fibroblasts (green). Scale bar refers to 1 mm. Reproduced, with permission, from (382).


Figure 26. Vascularization of a microfluidic polyethylene glycol gel that was patterned by photodegradation. (Left) Visualization of interconnected channels by perfusion with fluorescent dextran. (Right) 3D confocal reconstruction of a vascularized channel that was stained for ZO‐1 (green). Reproduced, with permission, from (196).


Figure 27. Perfusion‐decellularization of whole hearts preserves the vascular architecture. (A) Corrosion casts of native and decellularized hearts. Scale bars refer to 1 mm (top) and 250 μm (bottom). (B) Transplanted decellularized heart before and after reestablishment of blood flow. The recipient animal was heparinized to minimize thrombosis. Reproduced, with permission, from (424).


Figure 28. Physical mechanisms for stable vascularization of micro‐fluidic scaffolds. Maintenance of vascular adhesion can be viewed as (A) a balance of outward (stabilizing) and inward (destabilizing) stresses, including pressures, contractile stress, and adhesion stress, or as (B) a balance of adhesion (stabilizing) and stored elastic (destabilizing) energies. Adapted, with permission, from (643).
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FURTHER READING

Briquez PS, Clegg LE, Martino MM, Mac Gabhann F, Hubbell JA. Design principles for therapeutic angiogenic materials. Nat Rev Mater 1: 1-15, 2016.

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Tien J. Microfluidic approaches for engineering vasculature. Curr Opin Chem Eng 3: 36-41, 2014.

Yap KK, Yeoh GC, Morrison WA, Mitchell GM. The vascularised chamber as an in vivo bioreactor. Trends Biotechnol 36: 1011-1024, 2018.

Zheng W, Aspelund A, Alitalo K. Lymphangiogenic factors, mechanisms, and applications. J Clin Invest 124: 878-887, 2014.


 

 

 

 

Teaching Material

 

 

J. Tien. Tissue Engineering of the Microvasculature. Compr Physiol 9: 2019, 1153-1210.

Didactic Synopsis

Major Teaching Points:

  1. The ability to engineer microvasculature is crucial to the success of therapeutic vascularization and tissue engineering.
  2. Currently, the main strategies for engineering microvasculature are based on angiogenesis, vasculogenesis, and microfluidics.
  3. Engineered vasculature can be formed in an intrinsic or extrinsic manner.
  4. Local and sustained delivery of vascularizing signals usually leads to more robust formation of vascular networks, compared to systemic and transient delivery of the same signals.
  5. Biomaterial scaffolds can provide a means for local, controlled release of vascularizing signals and a space for vascular ingrowth.
  6. Widespread success of vascularization methods in laboratory and animal studies has yet to lead to clinically relevant therapies in humans.

    7. Didactic Legends

    The figures—in a freely downloadable PowerPoint format—can be found on the Images tab along with the formal legends published in the article. The following legends to the same figures are written to be useful for teaching.

    Figure 1 summarizes the different strategies used in engineering the microvasculature.

    Figure 2 shows examples of physiological data that can be used to obtain quantitative measures of vascular permeability to water and solutes.

    Figure 3 summarizes the members of the VEGF family and their cognate receptors and their effects on angiogenesis and lymphangiogenesis.

    Figure 4 illustrates the Vineberg procedure of vascularization, which is based on implantation of an artery into the ischemic tissue (myocardium).

    Figure 5 shows the increase in collateral vascular growth with implantation of FGF2-releasing beads in the ischemic pig heart.

    Figure 6 shows how the degree of vascular ingrowth into a scaffold depends on the pore size of the scaffold.

    Figure 7 shows that fibrovascular ingrowth into a scaffold can be induced by a variety of growth factors.

    Figure 8 shows that hepatocytes exhibit extremely low viability when implanted within a PLGA sponge and then transplanted in vivo.

    Figure 9 illustrates how to vascularize an overlying tissue (a skin graft) with an arteriovenous (AV) loop.

    Figure 10 shows an example of a centimeter-scale vascularized myocardium that formed after implanting cardiomyocytes with Matrigel around an arteriovenous (AV) loop in a polycarbonate chamber.

    Figure 11 shows that angiogenesis in microfluidic devices exhibits distinct phenotypes that correspond to the culture conditions.

    Figure 12 shows an image of EPCs and their associated expression of macrophage/monocyte-specific markers.

    Figure 13 shows an image of EOCs, their associated expression of EC-specific markers, and their lack of expression of macrophage/monocyte-specific markers.

    Figure 14 shows the incorporation of lacZ-positive EPCs into ischemic tissues.

    Figure 15 shows an example of vascular mimicry, in which blood fills structures that appear to lack an endothelial lining.

    Figure 16 shows vessels that formed within implants of HUVECs in a collagen/fibronectin gel.

    Figure 17 shows the generation of distinct blood and lymphatic vessels from a mixture of blood- and lymphatic vessel-derived ECs in a dermal graft.

    Figure 18 shows the development of aligned endothelial cords in vitro and their transformation into perfused microvessels in vivo.

    Figure 19 shows how microvascular fragments can generate perfused microvessels within a polymer implant over time.

    Figure 20 shows the generation of vascularized myocardial tissue from an implant of cardiomyocytes, HUVECs, and fibroblasts in a polymer scaffold.

    Figure 21 shows examples of vasculogenesis within EC-containing hydrogels in microfluidic devices.

    Figure 22 shows a lithographic pattern that was designed to mimic a branching vascular architecture.

    Figure 23 shows perfused microvessels that were formed within a microfluidic collagen gel that combined two separate gels to define the channels.

    Figure 24 shows a perfused microvessel that was formed within a microfluidic collagen gel that used removal of a needle to define the channel.

    Figure 25 shows a “tissue” that was formed around printed sacrificial fibers that dissolved to allow vascularization.

    Figure 26 shows an example of a vessel that was formed within a microfluidic polymer gel that used photodegradation to define the channel.

    Figure 27 shows corrosion casts of native and decellularized hearts and photographs of decellularized hearts before and after perfusion with blood.

    Figure 28 illustrates two complementary ways, based on a balance of stresses or energies, to analyze the stability of EC adhesion to a microfluidic scaffold.

 

 

 

 


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Angiogenesis
Physiology and Pathobiology of Microvascular Endothelium
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Article Title: Tissue Engineering of the Microvasculature
 

How to Cite

Joe Tien. Tissue Engineering of the Microvasculature. Compr Physiol 2019, 9: 1155-1212. doi: 10.1002/cphy.c180037