Adapted from: 

J Rouwkema, A Khademhosseini, Vascularization and angiogenesis in tissue engineering: beyond creating static networks, Trends in Biotechnology, 34(9); 733-745.


Importance of vascularization in tissue engineering

One of the goals of tissue engineering is to generate tissues that can be used as alternatives for donor material to repair or replace damaged tissues or organs [1]. Tissues generated for this purpose will generally be of a size larger than the diffusional limit for nutrients and oxygen [2]. Therefore, a need for a system to distribute nutrients within the tissue is apparent. During culture in the lab this distribution can be facilitated by using, for instance, perfusion bioreactors, but after implantation the tissue will have to rely on a vascular network to supply nutrients to all cells within the tissue. As part of the foreign body response, a vascular network will generally invade an implant. However, this is a process that takes days or weeks, meaning that cells in the middle of the tissue will be starved of nutrients for a considerable amount of time [3], resulting in suboptimal tissue integration or cell death.

In order to decrease the time that is needed to vascularize an engineered tissue, researchers have been exploring the possibility to add a vascular network before implantation. This network has the potential to connect to the vasculature of the patient, resulting in a much faster perfusion of the implant [4]. Theoretically, the perfusion will be instantaneous if the pre-engineered network in the implant is sufficiently organized, and can be microsurgically connected to the patient during the implantation procedure.

An optimal vascular network in an engineered tissue needs to possess several characteristics. One of the key tasks of a vascular network is to supply all cells in a tissue with sufficient nutrients. This means that all the cells need to be within a distance of 200 µm from a vessel, which is generally regarded as the diffusion limit of oxygen and nutrients within a tissue [5]. To reach this fine distribution while minimizing the pressure that is needed for the blood flow, the vascular network should be organized as a vascular tree, where larger vessels branch into smaller vessels, which ultimately branch into capillaries that are distributed throughout the tissue volume. Apart from that, the vascular network should act as a barrier that selectively controls the passage of materials from the vessels to the surrounding tissue to prevent excessive outflow of fluid leading to tissue edema. Finally, in order to supply the tissue with nutrients shortly after implantation, it should be possible for the network to easily connect to the vasculature of the patient, or to be microsurgically connected to it. For the latter, vascular structures with a diameter of several hundred micrometers are needed.

Vascularization is currently regarded as one of the main hurdles that need to be taken to translate tissue engineering to clinical applications at a large scale [6]. As such, it is one of the main research topics in the tissue engineering community. This review will give an overview of current strategies that have been explored in the past to prevascularize engineered tissues, and will propose future perspectives to engineer the optimal vascular network within an engineered tissue.


Angiogenesis and vascular remodeling

To include an organized vascular network in an engineered tissue, it is important to understand the process of vascular formation and remodeling. A starting point for this is to look at the formation of the vascular network during embryonic development and growth. Two processes can be distinguished during the formation of a natural vascular network; vasculogenesis and angiogenesis. Vasculogenesis is the process that takes place during early embryonic development where angioblasts (see Glossary) differentiate into endothelial cells, and proliferate within a previously avascular tissue to form a primitive capillary network [7]. Vasculogenesis can also occur in adults to revascularize a tissue following extensive damage or during tumor growth [8, 9]. During these so-called postnatal vasculogenesis processes, bone marrow derived endothelial progenitor cells are mobilized into the circulation, home to the tissue repair site, and differentiate into mature endothelial cells to form a primitive vascular network [8]Most of the processes involved in vascular organization and remodeling, both in embryonic development and afterwards, are angiogenic processes. Angiogenesis is defined as the formation of new vessels from an existing vascular network. These vessels can be formed by sprouting angiogenesis, where endothelial cells form sprouts starting from pre-existing vessels, or intussusceptive angiogenesis, where tissue pillars are inserted within existing capillaries to split the vessels [10]. Angiogenesis is mainly driven by the need to supply tissues with sufficient nutrients and oxygen. As such, angiogenesis is regulated to a large extent by oxygen levels within tissues. Hypoxic tissues secrete growth factors and chemokines that activate vascular growth and remodeling [11]. Endothelial cells are stimulated to break out of their stable position in the vessel wall and jointly coordinate sprouting, branching, and new lumenized network formation [12]. When perfusion of the tissue has increased and the supply of oxygen meets the demand, quiescence can be re-established resulting in a stable vascular network.  The process of angiogenesis initially leads to the development of capillaries. This can be followed by a process called arteriogenesis, where the vascular structures further mature and increase in diameter and the vascular wall thickness [13]. Mural cells proliferate, and acquire specialized characteristics, including contractile components [14]. The mechanisms governing arteriogenesis are not yet completely understood, but the remodeling process is mainly mediated by fluid flow shear stresses [15].


Natural organization of endothelial cells in engineered tissues

Many research efforts in prevascularized tissue engineering have relied on the spontaneous organization of endothelial cells to form vascular networks on scaffolds [16-18], in extracellular matrix analogues [4, 19], or in cellular aggregates with other cells [20, 21]. These studies show that endothelial cells are capable of forming vascular networks, often without the addition of specific cues or growth factors. First, the endothelial cells form a primitive network within a previously avascular, which is similar to vasculogenesis. Subsequently, this network can organize further in a process that is similar to angiogenesis and arteriogenesis. Depending on the culture conditions and the cell type with which the endothelial cells are combined, the morphology of the vascular networks ranges from immature cord-like networks with a limited amount of lumen [21, 22], to more mature networks containing well-developed lumen [4, 19] (see figure 1A). The addition of mural precursor cells such as embryonic fibroblasts or mesenchymal stem cells (MSC) results in maturation and stabilization of the vascular structures, indicated by an increase in the amount of lumen [4]. This is beneficial for the amount of blood that can be distributed in the tissue, since luminal structures are needed for the transport of fluids. Apart from that, mural cells play a role in the regulation of vascular permeability, which means that stabilized vessels will leak less fluid into the tissue, resulting in lower interstitial fluid pressure [23]. Upon implantation, the vascular structures can connect to the vasculature of the patient, thus contributing to implant perfusion and survival [4, 24]. Using intravital microscopy, Koike et al showed that vascular networks formed in fibronectin-collagen hydrogels can remain functional and transport blood for one year in vivo when stabilized by mural precursor cells, even though perfusion of the network was only apparent after 7 days of implantation [25].   

These studies are successful in demonstrating the potential of adding a vascular network to engineered tissues, but relying on uncontrolled angiogenesis in engineered tissues results in a random organization of the vascular network. Even though the networks will remodel after implantation, histology shows that the spacing between vascular structures is initially often larger than 200 µm, which means that the random organization is unlikely to be fully able to supply all cells of the engineered tissue with sufficient nutrients shortly after implantation. Apart from that, the random network does not offer clear locations for natural or surgical anastomosis, which can result in a delay of the network perfusion. 


Patterning of endothelial cells in engineered tissues

Figure 1: Fabrication of vascular networks.  Panel A shows co-cultures of HUVEC and MSC, where endothelial cells spontaneously organize into vascular networks (red) which are stabilized by mural cells (green) originating from the MSC. Panel B shows the organization of HUVEC in 2D by patterning both cell-adhesive and non-cell-adhesive regions on a 2D surface (left). By stacking multiple layers, a 3D networks is acquired (middle) which further remodels during culture (right). Panel C shows a strategy where channels are prepared in a collagen hydrogel, which are subsequently seeded with vascular cells. This results in a perfusable endothelial lined network (top). Depending on culture conditions and the presence of mural cells, vascular structures will sprout into the hydrogel, and will adapt their barrier function as illustrated using FITC-dextran perfusion (bottom). This approach can be extended to 3D by printing a sacrificial lattice network that is subsequently embedded in hydrogel. After removal of the sacrificial structure and seeding with vascular cells, a 3D perfusable network is acquired as shown in panel D. Finally, vascular cells can be patterned directly in 3D using bioprinting. By printing either hydrogels containing cells or cellular aggregates as shown in panel E, complex and well controlled vascular organizations can be achieved. Panels A, B, C, D, and E have been adapted with permission from [19], [43], [35], [34], and [50] respectively.


Many studies have focused on the active patterning of vascular networks within engineered tissues to closer resemble the natural organization of a vascular tree.  Using novel fabrication technologies, the initial organization of vascular cells can be designed and controlled. This approach offers the clear advantage that the resulting network can be designed such that all cells in the tissue are within 200 micrometer from a vessel, and provides clear locations for vascular anastomosis.

One strategy used to pattern vascular structures is to prepare hollow channels within scaffolds or hydrogel matrices, which can be seeded with vascular cells to form a predesigned pattern of vascular structures. Multiple methods have been reported to create hollow channels in polymeric biomaterial scaffolds, including the use of multi-material 3D fiber deposition [26, 27], electrospinning [28], the casting of scaffold material around sacrificial materials [29, 30], laser drilling [31], and the use of silicon molds [32, 33] to replicate patterns. Even though these approaches have been successful in generating well organized endothelialized vascular networks, these structures are generally bordered by a dense, impenetrable layer of biomaterial, limiting further vascular remodeling and nutrient transport. Apart from that, the resolution of these methods is generally insufficient to attain a complex network with a highly organized capillary bed.

Channels prepared in cell permissive hydrogels offer a more flexible environment, where endothelial cells can sprout into the matrix [34, 35], thus further remodeling the initial patterned vascular network. Individual channels [36-38] and interconnected channel networks [34, 35, 39, 40] have been fabricated in hydrogels using sacrificial materials such as gelatin [39] or glass filaments [34] (see figure 1D), structures such as hypodermic needles that can be removed after gelation [37], or polydimethylsiloxane (PDMS) molds [35]. Perfusion of these channels with culture medium results in the active transport of nutrients within engineered tissues, resulting in increased cell survival in vitro [41]. A recent study shows that channels as narrow as 20 µm can be seeded successfully with endothelial cells, resulting in millimeter-long perfusable capillaries [42]. This development enables the creation of an engineered tissue with a vascular-tree like network, including a highly organized capillary bed, where all cells are within 200 µm from a vascular structure. Apart from an approach to vascularize large tissue-engineered constructs, the resemblance of these vascular structures with natural vessels makes them a good research platform to test the effect of compounds on vascular structures by investigating, for instance, how readily fluid passes through the vessel wall [35, 40] (see figure 1C).

Next to the preparation of channels that can be seeded with endothelial cells and mural cells, vascular structures with a controlled geometry can be obtained by the direct patterning of vascular cells. Structures where vascular cells are patterned in two dimensions (2D) have been prepared using cell sheet technology [43], PDMS molds [44, 45], or photo-patterning [46]. These approaches all offer good control over the initial organization of the vascular network and the diameter of the vascular structures. By stacking these 2D constructs, more complex, three dimensional (3D) vascular networks can be obtained [43] (see figure 1B). Complex 3D vascular networks can also be prepared directly using technology such as bioprinting [47-49] which allows for the placement of either cellular aggregates [50] (see figure 1E) or biomaterials containing cells [51] at a specific location with a high level of spatial control.

The patterning of endothelial cells, either by preparing channels that are subsequently seeded with vascular cells, or by the direct patterning of these cells, enables the creation of complex vascular networks over multiple diameter scales. A study by Chaturvedi et al using patterned vascular structures shows that after implantation, vascular cords ranging in diameter from 25 to 250 micrometer can anastomose to the mouse vasculature and become functional, perfused vessels [44]. However, vessels remodeled in vivo, resulting in vessels with a diameter in the 10-15 micrometer range for all starting diameters. This study shows that even though the initial organization and diameter of vascular structures can be controlled, vascular remodeling will change this organization, either during in vitro culture or after implantation. Therefore, without further cues to control vascular remodeling, a carefully designed vascular network may not be sufficient to offer long term functionality.     


Guiding organization of endothelial cells in engineered tissues

An alternative approach to control the architecture of vascular structures in engineered tissues is to include local cues to guide vascular organization and remodeling. The adaptation of local microenvironments offers the possibility to engineer a complex, predictable vascular organization, starting from an initial random distribution of vascular cells. Even though this approach is less straightforward than the direct patterning of cells, guided morphogenesis may result in a better control of vascular organization on the long term due to an inherent control over the remodeling process. However, due to the complexity of all environmental factors that are involved in vascular organization and remodeling, an exact prediction and design of the resulting vascular network geometry will be challenging.   


Growth factors and chemical functionalities

As has been mentioned before, vascular organization and angiogenesis are controlled by growth factors. As such, many approaches to control vascular organization in engineered tissues are based on the local availability of these compounds. Since vascular endothelial growth factor (VEGF) is the factor that is involved in most angiogenic processes, it has often been the factor of choice to control vascular organization in a tissue engineering setting. It has been shown that in order to control vascular organization, it is not so much the availability of this factor, but the presence of gradients that controls vascular migration [52]. By creating distinct patterns of VEGF onto scaffolds or within hydrogels, gradients will be instituted, resulting in spatially driven endothelial cell elongation and branching [53, 54] (see figure 2A).

Angiogenesis and vascular organization are processes that consist of multiple phases. After the initiation of vascular network formation, a maturation process where endothelial cells are stabilized and mural cells are recruited is important for vascular function. In order to control both processes, patterning of a single growth factor will be insufficient. As such, researchers have combined the inclusion of VEGF with other factors governing maturation and mural cell recruitment such as platelet-derived growth factor (PDGF) [55, 56] and Angiopoietin 1 (Ang1) [57, 58], resulting in an increase in vascular structure formation and maturation compared to VEGF alone (see figure 2C).

A major challenge in the use of growth factors to control vascular organization in engineered tissues is the correct patterning of the multiple involved factors in space and time. In order to cope with this challenge, several approaches have been investigated where an indirect chemical signal is used. Since in vivo angiogenesis is largely governed by oxygen levels, oxygen gradients have been included in engineered tissues [59, 60], resulting in local differences in the expression of VEGF by fibroblasts present in the tissue [59]. Similarly, researchers have included the morphogen Sonic Hedgehog (Shh) to co-cultures of endothelial cells and MSC for a bone tissue engineering application [61]. This approach results in an increase of vascular structure formation and maturation, by modulating the expression of multiple angiogenic genes including VEGF, angiopoietins, laminins, and integrins. This result shows that indirect factors have the potential to tune the angiogenic environment of an engineered tissue, by stimulating the resident cells to secrete angiogenic factors. The advantage of this approach is that the factor secretion will be biologically regulated similar to in vivo angiogenic processes, where the cells will secrete factors based on the current need.   

Apart from using diffusible factors, chemical functionalities have been included in cellular environments to control vascular organization. Using two-photon laser photolithography, complex 3D patterns of chemical functionalities can be included in hydrogels [62, 63]. Hahn et al patterned the cell binding motive Arg-Gly-Asp-Ser (RGDS) in bio-inert polyethylene glycol (PEG) hydrogels functionalized with the peptide sequence GGPGQGILQGGK, which makes the PEG degradable to matrix metalloproteinase [63]. These enzymes are secreted by invading endothelial cells to degrade extracellular matrix. The study shows that invading cells are limited to the regions containing the RGDS pattern, thus resulting in the patterning of cellular ingrowth (see figure 2B). Similarly, by locally incorporating RGDS in PEG hydrogels, Culver et al could control the organization of human umbilical vein endothelial cells (HUVEC) and mural precursor cells to resemble the pre-scanned vasculature from the cerebral cortex of a mouse [62] (see figure 2D). The patterning of chemical functionalities thus allows for the creation of a vascular network with a designable organization, even when the vascular cells are allowed to organize themselves.

Figure 2: Guiding vascular organization via chemical signals. Panel A shows three situations where channels have been microfabricated in collagen hydrogels (top). The upper channel is seeded with HUVEC, while the lower channel is perfused with VEGF. By varying the distance between the channels, the local growth factor gradients vary, resulting in differences in endothelial sprouting and invasion (bottom). Panel C shows the effect of VEGF gradients on vascular invasion into a collagen gel either in the presence or absence of Ang1. When the cells are stimulated with both factors, more sprouts are formed, and the tip cells of the sprouts stay better attached to the stalk cells. This clearly indicates that the delivery of a single growth factor is often not enough to stimulate the formation of a well-organized vascular network.  Apart from controlling the local availability of growth factors, other chemical functionalities can be patterned to control vascular organization. Panel B shows the patterning of the cell binding motive RGDS into bio-inert PEG hydrogels using two-photon laser photolithography (top left). Since cells can only attach to the RGDS patterns, cell invasion is limited to these domains (top right and bottom). Panel D shows that by patterning RGDS domains in an organization resembling the vasculature of a mouse cerebral cortex (left), this approach can be used to replicate a natural vascular organization by controlling the invasion of HUVEC (green) and fibroblasts (red) (right). Panels A, B, C, and D have been adapted with permission from [54], [63], [58], and [62] respectively.



Cells respond to the mechanical properties of their environment. Mesenchymal stem cells for instance, adapt their differentiation profile to the local stiffness [64]. Several studies have shown that vascular organization is also dependent on the mechanical properties of the matrix in which the endothelial cells reside. A study by Santos et al demonstrates that HUVEC grown on collagen coated polyacrylamide hydrogels of low (3 kPa) and high (30 kPa) stiffness display similar proliferation and gene transcription levels, but show a lower expression of the functional VEGF receptor-2 protein on the stiffer substrate [65], which means that the extent to which endothelial cells are responsive to angiogenetic processes depends on the mechanical environment. In a different study, Mason et al used a system where the stiffness of a collagen hydrogel can be changed without significant changes to the collagen density and architecture. Increasing the stiffness from 175 Pa to 515 Pa results in a dramatic increase in the length and number of angiogenic sprouts growing from multicellular spheroids [66] (see figure 3A). Another study by Shamloo and Heilshorn points out that the matrix density and mechanical properties do not only have a direct effect on endothelial cells, but can also alter the response of endothelial cells to angiogenic growth factors such as VEGF. The number of sprouts, the sprout aspect ratios, and the direction of sprouts in relation to a VEGF gradient, all depend on the density of the collagen matrix [67].

Several studies have been reported where gradients and patterns of different mechanical properties are used to direct vascular organization within matrices. In a study by He et al, a stiffness gradient in an RGD-functionalized PEG hydrogel was formed using microfluidic mixing. This results in morphological differences of HUVEC seeded on the hydrogel, with cells maintaining a round morphology in the softer region, and spreading out in the stiffer region [68]. In a different study, Turturro et al report a gradient RGD-functionalized PEG hydrogel prepared using perfusion-based frontal photopolymerization, with the elastic modulus decreasing from 3.2 to 0.62 kPa over a span of 10 mm [69]. The gradient results in a decrease in the overall amount of vascular invasion from co-culture aggregates of endothelial cells and smooth muscle cells, but in a clear increase in the anisotropy of the vascular structures, with an alignment parallel to the direction of the gradient (see figure 3B).

Cellular responses to differences in matrix mechanical properties are mediated by the tension that cells can exert on the matrix upon cellular contraction [70]. Apart from locally changing the mechanical properties of a matrix, this phenomenon provides another opportunity to control vascular organization in engineered tissues. Using a matrigel assay in PDMS microwells of different shapes including circles, squares, triangles, and stars, Sun et al show that the HUVEC network near the boundary of the shapes has significantly higher densities and shorter mean cord length compared to the center regions [71]. Finite element analysis, points out that physical confinement can tune gel displacement due to cellular contraction, resulting in variations in cellular tension. Similarly, Rivron et al used a shaped microwell system to make biomaterial-free MSC-HUVEC co-culture microtissues of different shapes including circles, squares and triangles [72]. Local tissue compaction depends on the distance from the microtissue periphery, and the angles of the different shapes, resulting in patterns of vascular structures perpendicular to the strain direction in regions of high deformations. This corresponds with the institution of a long-range VEGF gradient in the interstitial cells, indicating that local tension can shape the angiogenic microenvironment in tissues.   

Fluid flow is an important factor for vascular organization and remodeling. During embryonic development of a chick embryo, the vascular network remodels from a largely random vascular plexus to an organized vascular-tree within 26 hours after the onset of perfusion [73] (see figure 3C). Researchers have used flow to guide vascular organization [74-76] and maturation [77] (see figure 3D). Apart from a direct response of vascular cells to fluid flow shear stresses, interstitial flow results in the institution of gradients of growth factors like VEGF, which further affects vascular organization [78].

Combined, these studies on the effect of mechanical signals show that vascular organization can be controlled. However, it should be noted that it is often not possible to adjust the mechanical environment within an engineered tissue with a high spatial resolution, which means that these strategies by themselves will most likely not enable the creation of a vascular network with designable features on the capillary scale. As such, mechanobiology should be regarded as a complementary tool to better control vascular organization, or as a method to stabilize an already organized vascular network. 

Figure 3: Guiding vascular organization via mechanical signals. Panel A shows a study where the mechanical properties of collagen hydrogels were adapted independently of collagen concentration. Changes in the compressive modulus of the hydrogel resulted in a clear effect on endothelial sprouting into the matrix, with stiffer gels resulting in an increase in the number and length of the sprouts. Panel B shows a similar approach where a gradient of RGD-functionalized PEG hydrogel is prepared using perfusion-based frontal photopolymerization, with the elastic modulus decreasing from 3.2 to 0.62 kPa over a span of 10 mm. Compared to a hydrogel with a constant bulk stiffness (left), the gradient hydrogel shows a clear increase in the anisotropy of vascular structure formation, with an alignment parallel to the direction of the gradient (right). Panel C demonstrates the importance of fluid flow during embryonic development. Just after the onset of perfusion, the vasculature in a chicken embryo yolk sac is poorly organized (top). There are no separate vessels carrying blood away from the heart (red arrows) or back toward the heart (blue arrows). Triggered by fluid flow, the same embryo shows an advanced state of vascular organization just 26 hours later. A clear distinction between arteries (red arrows) and veins (blue arrows) can now be made. Panel D shows an example of the use of fluid flow to affect the organization of an engineered vascular network. A perfusable network was formed in a microfluidic device, as evidenced by the perfusion of fluorescent microbeads (top left) and FITC-dextran (top right). Compared to static conditions, endothelial cells respond to luminal fluid flow with cytoskeletal reorganization (bottom left) and an increase in nitric oxide (NO) synthesis (bottom right). Panels A, B, C, and D have been adapted with permission from [66], [69], [73], and [74] respectively. 


Conclusions and future perspectives

When regarding vascular networks for engineered tissues, it is important to realize that quality is more important than quantity. It is not about the number of vascular structures in a given volume of tissue, but about the amount of blood that is perfused through the vascular network and the distribution of this blood over the tissue volume. Therefore, it is important that the vascular network is well organized and matured. In studies where angiogenesis is overstimulated resulting in excessive amount of vessels, tracer perfusion experiments show that the vessels are poorly perfused [79]. Studies that stimulate vascular maturation and stabilization on the other hand result in reduced vascular branching and density, but enhanced vessel diameter and perfusion [80]. The optimal vascular network will need to be highly organized, including venules, capillaries, and arterioles, in order to supply all cells with sufficient nutrients. However, organization is not the only characteristic that determines the success of engineered vessels. When assessing the quality of an organized vascular network, it is important to also take into account functional parameters such as perfusability and barrier function, something which is currently often lacking in studies that focus on the patterning of vascular networks. The presence of macrovascular structures that can be microsurgically anastomosed to the patient is desirable for direct perfusion after implantation. In cases where microsurgical anastomosis is not achievable, both the angiogenic activity of the implanted vascular structures and the host tissue can be optimized to enhance inosculation of the two vascular networks [81].

It is clear that approaches that focus on the active patterning of vascular cells within engineered tissues provide the highest level of control over the initial organization of vascular structures, and therefore have the potential to result in the most naturally organized vascular networks at the initial stage of tissue culture. However, given the activity and mobility of endothelial cells, these networks will remodel during in vitro culture and after implantation of the engineered tissue. When there are no additional cues to guide this remodeling process, it is likely that what starts out as a well-organized network will soon revert to a random organization of the vascular structures. As such, simply patterning vascular cells in engineered tissues may not be sufficient to ensure a good vascular organization on the long term.

Even though the requirements for vascular networks to be included in engineered tissues are generally well defined, it is not yet clear how all of these requirements can be fulfilled (see Outstanding Questions). As is often the case, an optimal protocol to add a well-organized vascular network will likely combine multiple of the approaches delineated above. Patterning of endothelial cells will provide a good starting situation, but one or more methods to control vascular remodeling and maturation will need to be included to ensure long-term functionality. Finally, it is important to realize that vascular networks will generally be engineered within a base tissue like muscle or bone. Strategies that are designed to direct vascular organization, such as growth factor localization or the patterning of mechanical signals, will often have an (unwanted) effect on the development of this tissue as well. As such, the strategies depicted in this review may not be useable in all situations, thus making it unlikely that the future will provide us with one single optimal method to add a vascular network to engineered tissues.



 1 Khademhosseini, A., et al. (2009) Progress in tissue engineering. Sci Am 300, 64-71

 2 Rouwkema, J., et al. (2008) Vascularization in tissue engineering. Trends Biotechnol 26, 434-441

 3 Butt, O.I., et al. (2007) Stimulation of peri-implant vascularization with bone marrow-derived progenitor cells: monitoring by in vivo EPR oximetry. Tissue Eng 13, 2053-2061

 4 Levenberg, S., et al. (2005) Engineering vascularized skeletal muscle tissue. Nat Biotechnol 23, 879-884

 5 Jain, R.K., et al. (2005) Engineering vascularized tissue. Nat Biotechnol 23, 821-823

 6 Jaklenec, A., et al. (2012) Progress in the tissue engineering and stem cell industry "are we there yet?". Tissue Eng Part B Rev 18, 155-166

 7 Risau, W. and Flamme, I. (1995) Vasculogenesis. Annu Rev Cell Dev Biol 11, 73-91

 8 Balaji, S., et al. (2013) The Role of Endothelial Progenitor Cells in Postnatal Vasculogenesis: Implications for Therapeutic Neovascularization and Wound Healing. Adv Wound Care (New Rochelle) 2, 283-295

9 Brown, J.M. (2014) Vasculogenesis: a crucial player in the resistance of solid tumours to radiotherapy. Br J Radiol 87, 20130686

10 Patel-Hett, S. and D'Amore, P.A. (2011) Signal transduction in vasculogenesis and developmental angiogenesis. Int J Dev Biol 55, 353-363

11 Fraisl, P., et al. (2009) Regulation of angiogenesis by oxygen and metabolism. Dev Cell 16, 167-179

12 Phng, L.K. and Gerhardt, H. (2009) Angiogenesis: a team effort coordinated by notch. Dev Cell 16, 196-208

13 Simons, M. (2005) Angiogenesis: where do we stand now? Circulation 111, 1556-1566

14 Carmeliet, P. (2000) Mechanisms of angiogenesis and arteriogenesis. Nat Med 6, 389-395

15 Helisch, A. and Schaper, W. (2003) Arteriogenesis: the development and growth of collateral arteries. Microcirculation 10, 83-97

16 Santos, M.I., et al. (2007) Response of micro- and macrovascular endothelial cells to starch-based fiber meshes for bone tissue engineering. Biomaterials 28, 240-248

17 Unger, R.E., et al. (2015) Improving vascularization of engineered bone through the generation of pro-angiogenic effects in co-culture systems. Adv Drug Deliv Rev

18 Unger, R.E., et al. (2007) Tissue-like self-assembly in cocultures of endothelial cells and osteoblasts and the formation of microcapillary-like structures on three-dimensional porous biomaterials. Biomaterials 28, 3965-3976

19 Chen, Y.C., et al. (2012) Functional Human Vascular Network Generated in Photocrosslinkable Gelatin Methacrylate Hydrogels. Adv Funct Mater 22, 2027-2039

20 Fuchs, S., et al. (2007) Microvessel-like structures from outgrowth endothelial cells from human peripheral blood in 2-dimensional and 3-dimensional co-cultures with osteoblastic lineage cells. Tissue Eng 13, 2577-2588

21 Rouwkema, J., et al. (2009) The use of endothelial progenitor cells for prevascularized bone tissue engineering. Tissue Eng Part A 15, 2015-2027

22 Kunz-Schughart, L.A., et al. (2006) Potential of fibroblasts to regulate the formation of three-dimensional vessel-like structures from endothelial cells in vitro. Am J Physiol Cell Physiol 290, C1385-1398

23 Goel, S., et al. (2011) Normalization of the vasculature for treatment of cancer and other diseases. Physiol Rev 91, 1071-1121

24 McFadden, T.M., et al. (2013) The delayed addition of human mesenchymal stem cells to pre-formed endothelial cell networks results in functional vascularization of a collagen-glycosaminoglycan scaffold in vivo. Acta Biomater 9, 9303-9316

25 Koike, N., et al. (2004) Tissue engineering: creation of long-lasting blood vessels. Nature 428, 138-139

26 Moroni, L., et al. (2006) Polymer hollow fiber three-dimensional matrices with controllable cavity and shell thickness. Biomaterials 27, 5918-5926

27 Luo, Y., et al. (2013) Direct plotting of three-dimensional hollow fiber scaffolds based on concentrated alginate pastes for tissue engineering. Adv Healthc Mater 2, 777-783

28 Sun, B., et al. (2015) Electrospun anisotropic architectures and porous structures for tissue engineering. Journal of Materials Chemistry B 3, 5389-5410

29 Wray, L.S., et al. (2012) A silk-based scaffold platform with tunable architecture for engineering critically-sized tissue constructs. Biomaterials 33, 9214-9224

30 Tocchio, A., et al. (2015) Versatile fabrication of vascularizable scaffolds for large tissue engineering in bioreactor. Biomaterials 45, 124-131

31 Malinauskas, M., et al. (2014) 3D Microporous Scaffolds Manufactured via Combination of Fused Filament Fabrication and Direct Laser Writing Ablation. Micromachines 5, 839

32 Borenstein, J., et al. (2002) Microfabrication Technology for Vascularized Tissue Engineering. Biomedical Microdevices 4, 167-175

33 Ye, X., et al. (2013) A biodegradable microvessel scaffold as a framework to enable vascular support of engineered tissues. Biomaterials 34, 10007-10015

34 Miller, J.S., et al. (2012) Rapid casting of patterned vascular networks for perfusable engineered three-dimensional tissues. Nat Mater 11, 768-774

35 Zheng, Y., et al. (2012) In vitro microvessels for the study of angiogenesis and thrombosis. Proc Natl Acad Sci U S A 109, 9342-9347

36 Zhao, L., et al. (2012) The integration of 3-D cell printing and mesoscopic fluorescence molecular tomography of vascular constructs within thick hydrogel scaffolds. Biomaterials 33, 5325-5332

37 Nichol, J.W., et al. (2010) Cell-laden microengineered gelatin methacrylate hydrogels. Biomaterials 31, 5536-5544

38 Sadr, N., et al. (2011) SAM-based cell transfer to photopatterned hydrogels for microengineering vascular-like structures. Biomaterials 32, 7479-7490

39 Golden, A.P. and Tien, J. (2007) Fabrication of microfluidic hydrogels using molded gelatin as a sacrificial element. Lab Chip 7, 720-725

40 Yoshida, H., et al. (2013) Multilayered Blood Capillary Analogs in Biodegradable Hydrogels for In Vitro Drug Permeability Assays. Advanced Functional Materials 23, 1736-1742

41 Lee, W., et al. (2010) On-demand three-dimensional freeform fabrication of multi-layered hydrogel scaffold with fluidic channels. Biotechnol Bioeng 105, 1178-1186

42 Linville, R.M., et al. (2016) Physical and Chemical Signals That Promote Vascularization of Capillary-Scale Channels. Cellular and Molecular Bioengineering, 1-12

43 Tsuda, Y., et al. (2007) Cellular control of tissue architectures using a three-dimensional tissue fabrication technique. Biomaterials 28, 4939-4946

44 Chaturvedi, R.R., et al. (2015) Patterning vascular networks in vivo for tissue engineering applications. Tissue Eng Part C Methods 21, 509-517

45 Raghavan, S., et al. (2010) Geometrically controlled endothelial tubulogenesis in micropatterned gels. Tissue Eng Part A 16, 2255-2263

46 Nikkhah, M., et al. (2012) Directed endothelial cell morphogenesis in micropatterned gelatin methacrylate hydrogels. Biomaterials 33, 9009-9018

47 Hoch, E., et al. (2014) Bioprinting of artificial blood vessels: current approaches towards a demanding goal. Eur J Cardiothorac Surg 46, 767-778

48 Villar, G., et al. (2013) A tissue-like printed material. Science 340, 48-52

49 Ozbolat, I.T. and Yu, Y. (2013) Bioprinting toward organ fabrication: challenges and future trends. IEEE Trans Biomed Eng 60, 691-699

50 Norotte, C., et al. (2009) Scaffold-free vascular tissue engineering using bioprinting. Biomaterials 30, 5910-5917

51 Bertassoni, L.E., et al. (2014) Hydrogel bioprinted microchannel networks for vascularization of tissue engineering constructs. Lab Chip 14, 2202-2211

52 Odedra, D., et al. (2011) Endothelial cells guided by immobilized gradients of vascular endothelial growth factor on porous collagen scaffolds. Acta Biomater 7, 3027-3035

53 Alsop, A.T., et al. (2014) Photopatterning of vascular endothelial growth factor within collagen-glycosaminoglycan scaffolds can induce a spatially confined response in human umbilical vein endothelial cells. Acta Biomater 10, 4715-4722

54 Baker, B.M., et al. (2013) Microfluidics embedded within extracellular matrix to define vascular architectures and pattern diffusive gradients. Lab Chip 13, 3246-3252

55 Richardson, T.P., et al. (2001) Polymeric system for dual growth factor delivery. Nat Biotechnol 19, 1029-1034

56 Chen, R.R., et al. (2007) Spatio-temporal VEGF and PDGF delivery patterns blood vessel formation and maturation. Pharm Res 24, 258-264

57 Chiu, L.L. and Radisic, M. (2010) Scaffolds with covalently immobilized VEGF and Angiopoietin-1 for vascularization of engineered tissues. Biomaterials 31, 226-241

58 Shin, Y., et al. (2011) In vitro 3D collective sprouting angiogenesis under orchestrated ANG-1 and VEGF gradients. Lab Chip 11, 2175-2181

59 Cheema, U., et al. (2008) Spatially defined oxygen gradients and vascular endothelial growth factor expression in an engineered 3D cell model. Cell Mol Life Sci 65, 177-186

60 Moore, M., et al. (2013) Directed oxygen gradients initiate a robust early remodeling response in engineered vascular grafts. Tissue Eng Part A 19, 2005-2013

61 Rivron, N.C., et al. (2012) Sonic Hedgehog-activated engineered blood vessels enhance bone tissue formation. Proc Natl Acad Sci U S A 109, 4413-4418

62 Culver, J.C., et al. (2012) Three-dimensional biomimetic patterning in hydrogels to guide cellular organization. Adv Mater 24, 2344-2348

63 Hahn, M.S., et al. (2006) Three-Dimensional Biochemical and Biomechanical Patterning of Hydrogels for Guiding Cell Behavior. Advanced Materials 18, 2679-2684

64 Wen, J.H., et al. (2014) Interplay of matrix stiffness and protein tethering in stem cell differentiation. Nat Mater 13, 979-987

65 Santos, L., et al. (2015) Extracellular Stiffness Modulates the Expression of Functional Proteins and Growth Factors in Endothelial Cells. Adv Healthc Mater

66 Mason, B.N., et al. (2013) Tuning three-dimensional collagen matrix stiffness independently of collagen concentration modulates endothelial cell behavior. Acta Biomater 9, 4635-4644

67 Shamloo, A. and Heilshorn, S.C. (2010) Matrix density mediates polarization and lumen formation of endothelial sprouts in VEGF gradients. Lab Chip 10, 3061-3068

68 He, J., et al. (2010) Rapid generation of biologically relevant hydrogels containing long-range chemical gradients. Adv Funct Mater 20, 131-137

69 Turturro, M.V., et al. (2013) MMP-sensitive PEG diacrylate hydrogels with spatial variations in matrix properties stimulate directional vascular sprout formation. PLoS One 8, e58897

70 Ventre, M., et al. (2012) Determinants of cell-material crosstalk at the interface: towards engineering of cell instructive materials. J R Soc Interface 9, 2017-2032

71 Sun, J., et al. (2014) Geometric control of capillary architecture via cell-matrix mechanical interactions. Biomaterials 35, 3273-3280

72 Rivron, N.C., et al. (2012) Tissue deformation spatially modulates VEGF signaling and angiogenesis. Proc Natl Acad Sci U S A 109, 6886-6891

73 le Noble, F., et al. (2004) Flow regulates arterial-venous differentiation in the chick embryo yolk sac. Development 131, 361-375

74 Kim, S., et al. (2013) Engineering of functional, perfusable 3D microvascular networks on a chip. Lab Chip 13, 1489-1500

75 Song, J.W. and Munn, L.L. (2011) Fluid forces control endothelial sprouting. Proc Natl Acad Sci U S A 108, 15342-15347

76 Hsu, Y.H., et al. (2013) A microfluidic platform for generating large-scale nearly identical human microphysiological vascularized tissue arrays. Lab Chip 13, 2990-2998

77 Price, G.M., et al. (2010) Effect of mechanical factors on the function of engineered human blood microvessels in microfluidic collagen gels. Biomaterials 31, 6182-6189

78 Helm, C.L., et al. (2005) Synergy between interstitial flow and VEGF directs capillary morphogenesis in vitro through a gradient amplification mechanism. Proc Natl Acad Sci U S A 102, 15779-15784

79 Thurston, G., et al. (2007) The Delta paradox: DLL4 blockade leads to more tumour vessels but less tumour growth. Nat Rev Cancer 7, 327-331

80 Noguera-Troise, I., et al. (2006) Blockade of Dll4 inhibits tumour growth by promoting non-productive angiogenesis. Nature 444, 1032-1037

81 Laschke, M.W. and Menger, M.D. (2012) Vascularization in tissue engineering: angiogenesis versus inosculation. Eur Surg Res 48, 85-92

82 Ruhrberg, C. (2003) Growing and shaping the vascular tree: multiple roles for VEGF. Bioessays 25, 1052-1060

83 Ferrara, N. (2001) Role of vascular endothelial growth factor in regulation of physiological angiogenesis. Am J Physiol Cell Physiol 280, C1358-1366

84 Gerhardt, H. (2008) VEGF and endothelial guidance in angiogenic sprouting. Organogenesis 4, 241-246

85 Iruela-Arispe, M.L. and Davis, G.E. (2009) Cellular and molecular mechanisms of vascular lumen formation. Dev Cell 16, 222-231

86 Cross, M.J. and Claesson-Welsh, L. (2001) FGF and VEGF function in angiogenesis: signalling pathways, biological responses and therapeutic inhibition. Trends Pharmacol Sci 22, 201-207

87 Khurana, R. and Simons, M. (2003) Insights from angiogenesis trials using fibroblast growth factor for advanced arteriosclerotic disease. Trends Cardiovasc Med 13, 116-122

88 Xue, Y., et al. (2012) PDGF-BB modulates hematopoiesis and tumor angiogenesis by inducing erythropoietin production in stromal cells. Nat Med 18, 100-110

89 Abramsson, A., et al. (2003) Endothelial and nonendothelial sources of PDGF-B regulate pericyte recruitment and influence vascular pattern formation in tumors. J Clin Invest 112, 1142-1151

90 De Marchis, F., et al. (2002) Platelet-derived growth factor inhibits basic fibroblast growth factor angiogenic properties in vitro and in vivo through its alpha receptor. Blood 99, 2045-2053

91 Danza, K., et al. (2013) Angiogenetic axis angiopoietins/Tie2 and VEGF in familial breast cancer. Eur J Hum Genet 21, 824-830

92 Folkman, J. (2007) Angiogenesis: an organizing principle for drug discovery? Nat Rev Drug Discov 6, 273-286

93 Sakurai, T. and Kudo, M. (2011) Signaling pathways governing tumor angiogenesis. Oncology 81 Suppl 1, 24-29

94 Ramsauer, M. and D'Amore, P.A. (2002) Getting Tie(2)d up in angiogenesis. J Clin Invest 110, 1615-1617