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Experimental approaches to vascularisation within tissue engineering constructs a

Md. Sarker , X.B. Chen

ac

ab

& D.J. Schreyer

a

Division of Biomedical Engineering, College of Engineering, University of Saskatchewan, 1A26, 57 Campus Drive, Saskatoon, SK S7N 5A9, Canada b

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Department of Anatomy and Cell Biology, College of Medicine, University of Saskatchewan, 5800 Saskatoon City Hospital, Saskatoon, SK S7K 0M7, Canada c

Department of Mechanical Engineering, College of Engineering, University of Saskatchewan, 3B47, 57 Campus Drive, Saskatoon, SK S7N 5A9, Canada Accepted author version posted online: 08 Jun 2015.Published online: 10 Jul 2015.

To cite this article: Md. Sarker, X.B. Chen & D.J. Schreyer (2015) Experimental approaches to vascularisation within tissue engineering constructs, Journal of Biomaterials Science, Polymer Edition, 26:12, 683-734, DOI: 10.1080/09205063.2015.1059018 To link to this article: http://dx.doi.org/10.1080/09205063.2015.1059018

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Journal of Biomaterials Science, Polymer Edition, 2015 Vol. 26, No. 12, 683–734, http://dx.doi.org/10.1080/09205063.2015.1059018

REVIEW ARTICLE Experimental approaches to vascularisation within tissue engineering constructs Md. Sarkera*, X.B. Chena,c and D.J. Schreyera,b a Division of Biomedical Engineering, College of Engineering, University of Saskatchewan, 1A26, 57 Campus Drive, Saskatoon, SK, S7N 5A9, Canada; bDepartment of Anatomy and Cell Biology, College of Medicine, University of Saskatchewan, 5800 Saskatoon City Hospital, Saskatoon, SK, S7K 0M7, Canada; cDepartment of Mechanical Engineering, College of Engineering, University of Saskatchewan, 3B47, 57 Campus Drive, Saskatoon, SK, S7N 5A9, Canada

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(Received 26 March 2015; accepted 2 June 2015) Tissue engineering opens up a new area to restore the function of damaged tissue or replace a defective organ. Common strategies in tissue engineering to repair and form new tissue containing a functional vascular network include the use of cells, growth factors, extracellular matrix proteins, and biophysical stimuli. Yet, formation of well-distributed, interconnected, and stable vascular network still remains challenging. In addition, anastomoses with host vasculature upon implantation and long-time survival of the new blood vessel in vivo are other critical issues to be addressed. This paper presents a brief review of recent advances in vascularization in vitro as well as in vivo for tissue engineering, along with suggestions for future research. Keywords: vascularization; angiogenesis; vasculogenesis; biomaterial; growth factors; scaffold

1. Introduction A major challenge in the creation of new tissues by tissue engineering is the limitation of diffusional mass transfer into new tissue having a thickness of more than 100–200 μm,[1] causing ischemia and cell death. Vascularization overcomes the limit of diffusional mass transfer for delivery of various types of nutrients, oxygen, growth factors, biochemical signaling factors, and removal of carbon dioxide, and metabolic waste.[2] The 100–200 μm limit regulates the critical distance necessary between two successive capillaries to prevent an ischemic condition.[3] This issue must be considered in any experimental or therapeutic effort to regenerate functional tissue of appreciable size using a tissue engineering approach. Two mechanisms are responsible for the generation of a vascular network in vivo: vasculogenesis and angiogenesis. Vasculogenesis generally takes place at an embryonic stage, while angiogenesis occurs mostly in postnatal life.[4,5] The term vasculogenesis denotes the assembly of endothelial progenitor cells (EPCs) that eventually form capillaries and a vascular network [6] in response to various growth factors, then undergoes remodeling to form functional vasculature. Angiogenesis refers to the sprouting of new blood vessels from existing ones due to specific angiogenic signals.[4] Engineered *Corresponding author. Email: [email protected] © 2015 Taylor & Francis

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tissue can be vascularized by either process, but must ultimately result in anastomosis with host vasculature.

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2. Different types of blood vessels and their functions The vascular system is a geometrically complex network consisting of arteries that supply blood to tissues, and veins that remove it (Figure 1). Major arteries branch out into successively smaller arterioles, metarterioles, and arterial capillaries. Capillaries combine downstream to form venules, and then major veins. Arterioles typically have diameters in the range of 10–300 μm and are composed of concentric layers of smooth muscle cells (SMC), basement membrane and endothelial cells (ECs).[7] A metarteriole is a short vessel which branches out from an arteriole with a smaller diameter. Generally, a metarteriole consists of only two layers; SMCs surrounding ECs. At the point where capillaries branch out from metarterioles, there exists a loop of SMCs called the precapillary sphincter. The function of precapillary sphincter is to control the blood flow to the capillary bed and to ensure enough time to facilitate diffusion and other mass transfer mechanism between capillaries and nearby cells. To keep the blood flow continuous, a bypass vessel called the thoroughfare channel connects the metarteriole with the venule. The capillaries consist only of ECs having an outer layer of protein-based basement membrane. However, pericyte cells, which are responsible for paracrine signaling and capillary blood flow, can be found within the capillary basement membrane. The capillaries are also known as microvessels, and have a diameter of 4–10 μm. The transition from capillary to venule takes place gradually and the approximate diameter of post-capillary venule is 10–50 μm. Post-capillary venules then fuse with a connecting venule whose diameter is 50–300 μm.[8] Capillaries are the principle site of mass transfer. The difference between the pressure of blood within the capillaries and the interstitial fluid pressure causes the movement of fluid into or out of capillaries. Generally, three types of capillaries are

Figure 1.

Blood circulatory system inside the body (Reproduced from [288]).

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Figure 2.

685

Structure of different types of capillaries (Reproduced from [288]).

seen in the body: continuous, fenestrated and sinusoidal (Figure 2). Most capillaries within the body are the continuous type. These have tight junctions as well as intercellular clefts between the ECs, and are covered with basement membrane. Water, gas molecules, ions, and other water soluble molecules can pass in or out of continuous capillaries through diffusion, vesicles, and intercellular clefts. However, high molecular weight molecules are transported across ECs by pinocytosis. Fenestrated capillaries have 60–80 nm pores in their ECs and are covered with basement membrane. Fenestrated capillaries carry out mass transfer through diffusion, vesicles, intercellular clefts, and pores. This type of capillary is found within the endocrine glands, intestines, pancreas, and kidneys. Sinusoidal capillaries are leaky in nature and have 30–40 μm pores, much larger than those seen in fenestrated capillaries. Sinusoidal capillaries have incomplete basement membrane covering and more intercellular clefts compare to other capillaries. White and red blood cell and serum protein can pass through the walls of sinusoidal capillaries. Generally, sinusoidal capillaries are found in bone marrow, liver, spleen, and adrenal gland.[9] 3. Mechanisms of blood vessel formation in vivo 3.1. Vasculogenesis EPCs from bone marrow or blood are mainly responsible for pre- and post-natal vasculogenesis through a complex series of steps (Figure 3). At the initial stage, EPCs move from their source and enter unvascularized tissue in response to chemo-attractant factors, such as granulocyte–monocyte colony stimulating factor, granulocyte colony stimulating factor, vascular endothelial growth factor (VEGF), basic fibroblast growth factor (bFGF), placental growth factor, erythropoietin and stromal cell-derived factor-1 (SDF-1).[10] Moreover, factors secreted from ischemic tissue cause nitric oxide (NO) production that activates extracellular proteases, especially matrix metalloproteases-9 (MMP-9), and facilitates EPC migration.[11] Homing of EPCs into ischemic tissue site requires chemotaxis, adhesion, and transendothelial migration. After moving out from bone marrow traveling through blood vessels, EPCs undergo chemotaxis toward a particular ischemic region in response to gradients of chemokines, such as SDF-1, interleukin-8 (IL-8), growth-regulated oncogene-α, and C–C chemokine.[12,13] The chemokines also stimulate EPCs to promote adhesion with the inner layer of blood vessels. EPCs pass through the endothelial lining

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Figure 3.

Md. Sarker et al.

Blood vessel formation by vasculogenesis (Reproduced from [289]).

of blood vessels by a specific transendothelial migration mechanism. Once at the basement membrane, EPCs initiate their invasion mechanism by rupturing it and remodeling basement membrane proteins, as well as the extracellular matrix (ECM) of the interstitial space, using extracellular proteases, such as MMP-9, cathepsin L, urokinase-type plasminogen activator, and tissue-type plasminogen activator, to reach the ischemic tissue site.[14,15] Then EPCs accumulate at the ischemic site; form a vascular pattern by differentiating into ECs and interacting with existing ECs and ECM. EPCs attached to the ECM proliferate under the influence of VEGF, immunoglobulin, and epidermal growth factor (EGF),[16,17] and differentiate due to monocyte chemoattractant protein1, insulin-like growth factor-1 (IGF-1), SDF-1, VEGF, and platelet-derived growth factor (PDGF).[18,19] 3.2. Angiogenesis During angiogenesis, new capillaries sprout from existing blood vessels in response to various biochemical signals and expand into ischemic tissue following tracks, gradients, or attractive or repulsive signals. Any unnecessary vascular plexus caused by random sprouting undergoes remodeling according to the requirements of specific tissue.

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Functional non-uniformity of ECs along the blood vessel wall caused by VEGF gradients plays a critical role in angiogenic sprouting. Mechanistically, specific ECs respond to a chemo attractant by extending as tip cell, then propagating in different directions by forming stalk-like structure, which eventually acquire a lumen.[20,21] To communicate with the surroundings, tip cells extend multiple filopoida which are sensitive to various guidance molecules and specific growth factors, such as VEGF-A, FGF, angiopoietin-1 (Ang-1), EGF, and IL-8. The spatial as well as temporal distributions of these factors are very crucial for guiding the growth of new capillaries within the ischemic tissue.[22] For example, a VEGF-A gradient in tissue plays an important role for filopodia extension from the tip cell, and the corresponding receptor for VEGF-A is VEGFR-2, which is expressed by the tip cell.[23] Angiogenesis in ischemic or newly formed tissue is a complicated process which can be demonstrated through some steps (Figure 4). At the beginning, ischemic tissue secretes VEGF which dilates the existing blood vessel through the redistribution of intercellular adhesion molecules as well as modification of the cell membrane structure. It also increases vascular permeability with the help of NO [24,25] and assists the relocation of plasma protein to form temporary structures to facilitate EC migration. Besides VEGF, Ang-2 contributes to angiogenic sprouting through the degradation of ECM protein and the removal of SMCs from the outer surfaces of capillaries.[25,26] Degradation of basement membrane and ECM are vital for EC migration, while microvascular ECs themselves secrete some matrix metalloproteases, such as MMP-2, MMP-3, MMP-9, along with protease inhibitor mainly tissue inhibitor, of metalloproteinase-2, which coordinately regulate basement membrane and ECM degradation [27] by causing various spatial and temporal factors to

Figure 4.

Blood vessel formation by angiogenesis in brain cancer (Reproduced from [290]).

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generate non-uniformity along the wall of blood vessel.[28,29] In addition, another important feature of extracellular proteolytic degradation is the release of some growth factors, such as IGF-1, VEGF, and FGF-2, which also facilitate angiogenic sprouting.[30] The progression step is associated with the migration and proliferation of ECs due to the influence of VEGF, angiopoietins (Angs), FGFs, and PDGF. Among them, Ang1 regulates the interaction between ECs and periendothelial cells during migration, [25,31] FGF facilitates EC growth via the recruitment of mesenchymal cells,[32] and PDGF enhances EC growth through the recruitment of pericytes and SMCs.[33] In addition, other factors involved in EC proliferation and migration include neuropeptides, IGF-1, erythropoietin, hepatocyte, and interleukins.[34,35] After migration and proliferation of EC into ECM, ECs eventually differentiate to form cord-like structure and a lumen starts developing due to the influence of VEGF and Ang. Interestingly, Ang-1, VEGF121, and VEGF165 help to increase lumen diameter, while VEGF189 decreases lumen diameter.[36] 4. Tissue engineering approaches to blood vessel formation Well-distributed and interconnected vascular network is required to maintain the viability of large cell population in engineered tissue. While tissue engineering is eventually emerging as a potential approach to regenerate tissue and organ, formation of 3D vascular network within the neotissue still remains challenging. Current strategies to form complex vascular channel within engineering construct or growing tissue can be categorized as scaffold-based and scaffold-free approaches (Figure 5). Strategies (Table 1) associated with scaffold-based approaches are, construction of 3D matrix with appropriate biomaterial by biofabrication technique or stacking multiple micropatterened thin planar surfaces or micro tissue modules, engineering new tissue with decellularized matrix, functionalization of scaffolds with cells, angiogenic factors (AFs) or proteins, transfection of seeded cells by gene therapy to obtain sustained release of AFs and prevascularisation to maintain viability of the seeded cells. In contrast, in scaffold-free approach cell sheets are assembled to promote vascularized tissue. 4.1. Scaffold-based aproaches 4.1.1. Scaffold fabrication biomaterial The choice of matrix biomaterial has profound impact on scaffold vascularization in tissue engineering. Like natural ECM, the biomaterial should facilitate vascular cell– biomaterial interactions, which would activate numerous signaling pathways to promote vascular cell survival, proliferation, differentiation, and migration. A tissue scaffold experiences different types of force upon implantation in vivo, such as compression, shear, torsion, and tensile force. Therefore, the biomaterial should have sufficient mechanical strength to sustain those forces to keep the scaffold architecture intact. Moreover, controlled biodegradability is desired so that over time, the scaffold could maintain its mechanical strength while the biomaterial is replaced with a vascular network and newly formed tissue. Furthermore, cytotoxicity has to be considered for its impact on implanted or host cell survival in vivo or during preparation in vitro. Generally, polymers are used as biomaterials for the fabrication of scaffolds.[37] Biopolymers can be classified into three categories in terms of preparation or source: synthetic, natural, and hybrid.

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Figure 5.

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Different approaches for vascular network formation.

4.1.1.1. Synthetic polymer. Synthetic polymers used for tissue engineering offer scaffold fabrication flexibility while maintaining desired mechanical properties of the engineering construct. However, lack of cell adhesion peptides in the molecular structure of synthetic materials limits their application for vascular tissue engineering. A number of biodegradable, biocompatible, and non-toxic synthetic polymers have shown potential for tissue engineering applications. Among them, polyglycolic acid (PGA), polylactic acid (PLA), polylactic-co-glycolic acid (PLGA), poly-L-lactic acid (PLLA), poly-ε-caprolactone (PCL), polyethylene glycol (PEG), and polyhydroxyalkanoates (PHAs) have frequently been explored in different studies of tissue-engineered graft vascularization.[38] When exposed to pulsatile conditions, PGA mesh tube scaffolds seeded with porcine carotid SMCs were eventually replaced by blood vessel-like structures while maintaining uniform SMCs distribution and collagen content close to that of native vessels.[39] In vivo experiments have also shown PGA scaffolds as promoters of vascularization. PGA scaffolds seeded with bovine artery cells degraded completely after 11 weeks of implantation in the pulmonary artery of sheep and were replaced by new tissue having 73.9% collagen content with respect to native tissue.[40] Similarly, in an in vivo study, PEG diacrylate hydrogel was implantated subfascially into SpragueDawley rats and kept in vivo for three weeks. Interestingly, even in the absence of incorporated growth factors, vascularized tissue ingrowth was reported throughout the entire PEG gels having pore size 50–150 μm.[41] Derivatives of PLA (polymer of lactide) have also shown their angiogenic potential in different studies. For example, in vivo implantation of tubular scaffold fabricated

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Table 1.

Tissue engineering approaches to blood vessel formation.

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Approaches

Description

A. Scaffold based approach A.1. Scaffold fabrication Synthetic polymers (e.g. PGA, PLA, PLGA, PLLA, PCL, PEG, biomaterial and PHAs); natural polymers (e.g. collagen, fibronectin, fibrin, elastin, silk fibroin, HA, alginate, chitosan); decellularized matrix (e.g. allogenic or xenogenic tissue or organ graft) A.2. 3D printing 3D Fabrication (e.g. 3D plotting, inkjet printing, laser-based approaches biofabrication and stereolithography) technique to fabricate complex vascular network; microfabrication (e.g. microfluidics, micromolding, lithographic, direct laser write) technique to indent micro pattern on polymer or cell seeded hydrogel to allow complex network formation at micro level; modular assembly of patterned micro modules generated from self-assembled aggregation, microfabrication of cell seeded hydrogels, or direct printing method to form macro tissue assembled by random packing, stacking of layers or directed assembly A.3. Addition of cells Autologus mature ECs (e.g. human umbilical vein, human dermal microvascular, human vascular), stem cells (e.g. embryonic stem cell, MSC), EPCs (e.g. peripheral blood derived, umbilical cord blood derived), vascular wall cells (e.g. pericytes, vascular SMCs, pulmonary artery SMCs) A.4. Addition of growth VEGF, FGFs, PDGF, Angs, TGF-β factors A.5. Gene therapy Viral vectors (e.g. retrovirus, lentivirus, adeno-virus and adenoassociated virus) and nonviral vectors (e.g. plasmid DNA) A.6. Prevascularisation In vitro or in vivo culture prior to implant to form vasculature and maintain cell viability B. B.1.

Scaffold free approach Cell sheet technique Temperature controlled cell attachment, culture and detachment followed by stacking multiple layers

with porous films of poly-D, L-lactic-co-glycolic acid facilitated the ingrowth of fibrovascular tissue, which eventually formed a vascularized, tubular tissue.[42] To investigate the combined effect of PLLA and collagen gel on vascularization, porous PLLA scaffold was exposed to in vitro condition, where the pores were filled with the mixture of human aortic SMCs and collagen gel, resulting in the rapid formation of a smooth inner layer around the vascular graft.[43] Another composite polymer of lactide PLGA was explored to study the release profile of VEGF. VEGF was directly incorporated into PLGA scaffolds and pre-encapsulated in PLGA microspheres. It was reported that direct incorporation of VEGF resulted in 40–60% release within five days in vitro, while encapsulation of VEGF within microspheres resulted in prolonged release. When implanted into subcutaneous pockets of SCID mice, VEGF released from PLGA scaffolds significantly enhanced local angiogenesis.[44] Like PLA, composites and derivatives of PCL have been identified as a vasculature enhancer in several studies. In a comprehensive review, Woodruff et al. presented the attractive features of PCL (e.g. inexpensive, FDA-approved, and superior rheological and viscoelastic properties) to demonstrate its applicability for the fabrication of longer term degradable implants suitable for a specific anatomical site.[45] In vivo implantation of PCL/PLA tubular scaffolds seeded with mixed cells obtained from femoral

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veins of a mongrel dog showed no evidence of stenosis, thrombus, occlusion, or aneurysmal formation within the tubular graft. Moreover, the vascular graft was replaced by native tissue containing a large amount of ECM, where the ECs were found to organize themselves in a linear way on the luminal surface of scaffold.[46] Besides animal model study, in May 2000 a clinical trial was done on a four-year-old girl to implant a pulmonary bypass graft made of PCL/PLA copolymer seeded with autologous vein cells. This implantation was a milestone in pediatric cardiovascular surgery as the patient was doing well even after seven months of implantation and no postoperative complications were reported.[47] Another derivative of PCL called poly (lactide-co-caprolactone) (PLCL) was studied for tissue vascularization due to its biocompatibility, elasticity, and slow degradation property. In a study, PLCL vascular grafts seeded with rabbit aortic SMCs were exposed to pulsatile flow in a perfusion bioreactor for eight weeks. It was reported that the PLCL graft significantly regulated the proliferation of SMCs, increased collagen production, and promoted cell alignment similar to that of native vascular smooth muscle tissues.[48] The angiogenic potential of PHA derivatives and its composites were demonstrated in different investigations. In a study, PHA–PGA tubular scaffolds seeded with ovine carotid artery cells were implanted into abdominal aortic segments of sheep for five months. No aneurysms were observed in the tubular graft; however, the percentage of collagen, DNA content, as well as the mechanical strength of grown tissue, reached close to that of native vessels.[49] Another research group maintained static/pulsatile condition for PGA/PHA derivative-fabricated vascular grafts seeded with ovine vascular myofibroblasts and ECs. After 28 days of culture, it was reported that all samples gained viable, dense, and confluent smooth tissue with high collagen content.[50] 4.1.1.2. Natural polymer. Biopolymer or naturally derived polymers can be classified into two categories, such as; protein-based polymers and polysaccharidic polymers. In tissue engineering, natural polymers are more advantageous over synthetic polymers in many aspects, however, are not free from shortcomings.Some advantages and drawbacks of natural polymers have been presented in Table 2. Protein-based polymers, particularly collagen, fibrin, fibronectin, elastin, and silk fibroin, have frequently been explored in different studies for vascularization application. For example, it has been reported that freeze-dried collagen scaffold seeded with dermal fibroblasts and bone marrow-derived stem cells enhance the proliferation of EC.[51] Further, immobilized VEGF on collagen scaffolds promotes EC invasion and proliferation.[52] Another group incorporated modified VEGF into collagen scaffolds implanted subcutaneously into mice, resulting in an improved vascular network within the implanted scaffold.[53] Besides, Fibrin tissue engineering scaffolds have also presented promising results for promoting blood vessel formation. Human microvascular ECs were seeded into a 90% fibrin, 10% collagen matrix with combinations of added growth factors; FGF-2 or VEGF, with TNF-α.[54] This investigation suggests that fibrin matrix facilitates the formation of a vascular capillary network through matrix fibrinolysis by ECs, which occurs from the activities of plasmin and metalloproteinases. In addition to providing specific cell adhesion sequences, a fibrin scaffold can store and release angiogenic growth factors, such as VEGF-A165 and FGF-2, as well as plasmid DNA to improve angiogenesis.[55] Another ECM protein fibronectin has been considered as a vasculature promoter due to its enormous contribution in vascularization. In a study, fibronectin-coated collagen modules increased human vascular EC survival and vessel formation when implanted in immunodeficient mice compared to collagen

692 Table 2.

Md. Sarker et al. Natural biomaterials for scaffold fabrication.

Biomaterial Collagen

Advantages (1)

(2)

Biocompatible, biodegradable, ease availability and modifiability, nonantigenic and controlled protein release ability.[291] Hydrophilic and enhance cell interactions.[292]

Disadvanages (1)

(2)

(3)

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Hyaluronic acid

(1)

(2)

Fibrin

(1)

(2) (3)

Fibronectin

(1) (2) (3)

Alginate

(1)

Easily modifiable, hydrophilic, nonadhesive, biodegradable,[297] nonantigenic and noninflammatory.[298] Regulate cellular migration, interaction, and differentiation.[299] Non toxic, non inflammatory, [302] non allergenic,[303] non immunogenic[304], economical,[305] biodegradable [306] and easily processable.[307] Facilitates synthesis of collagen [308] and promotes angiogenesis.[309] Enhances attachment, migration and proliferation of smooth muscle [310] and ECs.[311]

(1) (2)

(1) (2) (3)

Poor mechanical strength, [293] deformability, flexibility and tensile strength.[294] High cost of purification, handling complexity,[295] rapid degradation, little compression force resistivity.[296] Sterilization causes alteration of protein structure.[291] Requires purification to avoid disease transmission.[300] Rapid degradation rate, poor mechanical strength,[301] and forms scar tissue in vivo.[294]

Inconsistency in polymerization, fragile and stress squeezable. Structural weakness.[312] Upon implantation, solubility increases and structural integrity decreases over time.[313]

Critical role for angiogenesis [314] and scaffold vascularisation.[315] Facilitates cell adhesion,[316] migration,[317] proliferation and alignment.[318] Can be used to coat artificial biomaterials to support cell adhesion, spreading [319] and proliferation.[320]

(1) (2)

High cost.[321] Excessive mechanical stress interrupts formation of vasculature.[322]

Non inflammatory,[321] biocompatible, non toxic, cheap and potential for cell encapsulation and drug delivery.[323,324]

(1)

Uncontrollable degradation kinetics due to loss of divalent ion and burst release of protein drugs at higher pH.[325] (Continued)

Journal of Biomaterials Science, Polymer Edition Table 2.

(Continued).

Biomaterial

Advantages (2)

pH sensitive behavior facilitates drug delivery.[325]

Disadvanages (2) (3) (4)

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Poor mechanical strength.[326] Hydrophilic character prevents it from adsorption of cell adhesive protein.[327] Lack of cell adhesion.[323]

modules without fibronectin coating.[56] Similarly, implantation of a fibronectin/ collagen I gel seeded with EC and mesenchymal cell facilitated the formation of wellperfused and long-lasting vascular network after implantation into mice.[57] The role of structural protein elastin in vascularization is very significant as genetically modified mice lacking elastin died within four days of birth from arterial obstruction resulting from subendothelial cell proliferation and reorganization of smooth muscle into the lumen of artery.[58] Cross-linked soluble alpha-elastin discs were found to facilitate vascular SMC adhesion,[59] whereas tropoelastin-coated surface was reported as a promoter of EC attachment and proliferation.[60] Elastin composites have also been investigated to vascularize tissue engineering grafts. For instance, in a study, human coronary artery SMCs were cultured for seven days on aligned nanofibrous polyurethane scaffolds blended with the mixture of elastin and collagen. It was concluded that the growth of SMCs on elastin/collagen-blended scaffold increased by 224% compared to that of polyurethane.[61] Besides animal body-derived protein, silkworm-secreted protein silk fibroin showed promising results in several studies in tissue vascularization. In a study, co-culture of human dermal microvascular endothelial cells (HDMECs) and primary human osteoblast cells (HOCs) in 3D silk fibroin nets caused ECs to form intertwined microcapillary-like structures.[62] In another study, 3D silk fibroin scaffolds pre-cultured with HDMEC and HOC were implanted into immunodeficient mice. After 14 days, it was found that the microcapillary structures pre-formed in vitro, perfused well with the host vasculature.[63] Apart from protein-based polymers, polysaccharide polymers, mostly hyaluronic acid (HA), alginate, and chitosan have been studied for tissue vascularization. For example, subcutaneously injected HA combined with recombinant gelatin facilitates the formation of vascular network as well as the deposition of ECM components in rats.[64] Another group achieved controlled release of two angiogenic growth factors (VEGF165 and PDGF-BB) from a hybrid mesh consisting of poly (3-caprolactone)collagen blend and HA hydrogel. They reported that the growth factor-loaded hybrid mesh enhances cellular attachment and formation of vascular capillary network within the tissue engineering construct during the culture of ECs and fibroblasts in a 3D model.[65] However, HA has limitations in tissue engineering applications as it exhibits poor cell attachment and mechanical properties. Seaweed-derived polysaccharide alginate does not have EC adhesion properties, but can be introduced through adsorption of proteins,[66] or by covalent incorporation of different functional groups into the sugar backbone.[67] To facilitate vascularization within alginate scaffolds, internal pores should be well interconnected and large enough to allow ingrowth of blood vessels upon implantation in vivo.[68] For example, an

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alginate scaffold with 90% porosity and pore sizes ranging from 50 to 200 mm facilitates human embryonic stem cell aggregation and the formation of void and tubelike structures in vivo.[69] Furthermore, growth factors can be incorporated in alginate scaffold to enhance vascularization, for instance incorporation of VEGF in alginate hydrogels enhance neovascularization into the matrix.[70,71] Due to remarkable wound healing capability, chitosan and its composites were explored for tissue engineering graft vascularization in different studies.[72] In a study, porous chitosan, glycosaminoglycans (GAGs)-chitosan and dextran sulfate (DS)-chitosan scaffolds were implanted in dorsal subcutaneous pockets of male Sprague-Dawley rats. The rats were sacrificed after two weeks and it was noticed that chitosan, heparin– chitosan, and DS-chitosan scaffolds promoted cell proliferation, tissue ingrowth, and formation of vascularized granulation tissue. Chitosan composites can also be used for the controlled release of growth factors. To study the release profile of an AF from chitosan complexes, FGF-2 incorporated chitosan/heparin hydrogel was subcutaneously injected into the back of male Sprague-Dawley rats. Controlled release of FGF-2 molecules for 20 days caused micro blood vessel and fibrous tissue formation around the injected site.[73] 4.1.1.3. Decellularized matrix. Decellularization of tissue or whole organ causes the formation of ECM protein-enriched three-dimensional bioscaffold. The acellular matrix contains unique, tissue-specific, structural and functional molecules that regulate cellular phenotype and function, mechano-transduction, signaling, cell–matrix interactions as well as tissue homeostasis.[74,75] Therefore, matrix decellularization followed by recellularization with autologous cell could be a promising approach in tissue engineering. The technical difficulty of removing all cellular remnants limits the application of decellularized matrix due to the risk of a host immune response after transplantation. Moreover, preserving the native properties of ECM, for example, three-dimensional ultrastructure, surface topography, composition, bioactivity, and density of ligand distribution as well as internal network, such as nervous, vascular, and lymphatic networks, presents another technical challenge.[76] One of the important features of using decellularized matrix is that it helps to rebuild the intricate vascular network within the organ or tissue when the intact vascular spaces within the matrix are repopulated with ECs. Tissue decellularization can be accomplished by chemical, biologic, or physical agents. Selection of an agent largely depends on tissue size, thickness, density, cellularity, and lipid content. Chemical agents, for example, acids and bases, hypotonic and hypertonic solutions, ionic, non-ionic, and zwitterionic detergents, or solvents, can be used to decellularize tissue based on the mechanism of solubilizing cytoplasmic components, denaturing proteins, dehydrating cells, disrupting nucleic acids, lipids, and proteins. However, chemical agents may cause some disruption of the ultrastructure of ECM as well as damage of collagen, GAGs, and growth factors.[77,78] Biologic agents, such as nucleases, trypsin, or dispase, are basically enzymes that work by catalyzing the hydrolysis of RNA and DNA chains or cleaving peptide bonds. The use of biologic agents can also result in complications, for example, it can cause an immune response, can disrupt ECM ultrastructure, or can remove collagen, laminin, fibronectin, elastin, and GAGs after prolonged exposure.[77,79] Physical techniques based on temperature, pressure, direct force, electroporation, perfusion, agitation, pressure gradient, or super critical fluid can also be used to remove cellular material from tissue. Physical

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agents work by facilitating chemical exposure, bursting cells, or disrupting cell membranes, but can cause disruption or damage of ECM due to physical force.[77] Repopulation of a decullarized matrix with appropriate cells is a major engineering challenge to be solved. The parenchymal cells, for example, hepatocytes, cardiomyocytes, epithelium, etc., are responsible for the specific functions of the organ, while nonparenchymal cells, such as fibroblasts and ECs, help to maintain the functional phenotype of the parenchymal cells and the cellular architecture of the tissue.[80,81] Moreover, ECs eliminate the thrombotic barrier within decellularized matrix and protect the parenchymal cells from the shear stress by maintaining blood flow within the vascular network.[82,83] Therefore, the success of decellularized matrix as a scaffold mostly depends on re-endothelialization of the intact vascular space. Autologus, allogenic, progenitor, or multipotent stem cells are frequently investigated for their ability to repopulate decullarized tissues or organs. Non-immunogenic autologous cells are the best choice because no immunosuppressive drug is required. However, harvesting complexity and inadequate proliferation capability limit the use of autologous cells to repopulate acellular matrix. By some criteria, for example, required cell population, urgency of implantation, simplicity of cell harvesting and expansion, and capability of differentiation into desired cell types, allogenic cells could be more useful [75] if immunogenic reaction and the use of immunosuppressive drugs can be tolerated. Multipotent stem cells are a third option to repopulate decellularized matrix, although challenges exist here in controlling or directing differentiation along organ-specific or tissue-specific cell lineages.[76] Bioreactors with appropriate biophysical stimuli might be used to assist in differentiation of stem cells. A number of studies suggest a promising future for organ or tissue vascularization using decellularized matrix. For example, rat hearts decellularized by coronary perfusion and reseeded with cardiac or ECs were kept in a bioreactor, where a simulated cardiac physiology was maintained for 28 days. In this study, EC formed single layers in both larger and smaller coronary vessels after day seven, and recellularized heart exhibited pump function by day eight under physiological load with electrical stimulation.[84] Similarly, lungs decellularized by detergent perfusion, reseeded with epithelial or EC, and then perfused with blood and ventilated using physiologic pressures resulted in gas exchange similar to that of isolated native lungs by day five. Moreover, when these reseeded lung constructs were reimplanted into an orthotopic position, they generated gas exchange for up to 6 h after extubation.[85] 4.1.2. 3D printing approaches to create vascular networks Vascular cell survival, growth, signaling, gene expression, and phenotype during matrix tissue culture is significantly affected by the pore architecture, pore interconnectivity and mechanical stiffness of the scaffold.[86–88] The porous architecture of a scaffold includes pore size, shape, porosity, and surface topography of the pores. Proper porosity allows cell migration, proliferation, and interaction that facilitate the formation of a vascular network. In addition, porosity assists to transport nutrients and oxygen gas to cells and remove metabolic waste from cell surroundings. However, excessive porous structure might result in poor mechanical strength. In bone tissue engineering, higher porosity and larger pore size of the scaffold enhance vascularization and tissue growth. For example, a six-day in vitro investigation on the proliferation of goat bone marrow stromal cells suggests that scaffolds having

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70% porosity and 800 μm average pore size enhance stromal cell proliferation compared to scaffolds possessing 60% porosity and 700 μm average pore size.[89] Like porosity, the average pore size of a scaffold has a profound effect on tissue growth and vascularization. A number of studies suggest that variation of pore size affects cell functionality. For example, it was reported that implanted polytetrafluoroethylene membrane having pore size 5 μm under rat skin significantly facilitates the capillary blood vessel formation across the membrane–tissue interface.[90] When fibronectin-coated porous silicon nitride substrates seeded with 3T3 fibroblasts and bovine aortic ECs were kept in ex vivo condition for five days, EC covered the scaffold pores having diameter below 80 μm. Interestingly, pore size under 30 μm didn’t display any effect on EC coverage. Besides, fibroblast could cover pores of 50 μm and less.[91] Further, disc-shaped porous PLA scaffolds with pore size in the range of 63–150 μm facilitated vascular SMC proliferation and matrix deposition during a four-weeks culture period.[92] Therefore, a potential strategy would be to fabricate scaffolds with targeted pore size that would promote the desired cellular activity, for example, tissue ingrowth as well as blood vessel formation. The surface area of a scaffold is also an important parameter to be considered as there exists a relationship between pore size and available surface area for cell attachment where an RGD binding sequence motif prevails. In general, small pore size increases specific surface area, which ensures minimal ligand density for the attachment of a critical number of cells.[93,94] On the other hand, large pore size facilitates cell migration but reduces cell density. Finally, pore interconnectivity within the scaffold can have a profound influence on neovascularization and tissue growth. The diffusional mass transfer of nutrient and oxygen gas and removal of waste metabolites can be interrupted due to improper pore interconnectivity.[95] A β-tricalcium phosphate scaffold fabricated by the casting technique was used in a rabbit model to investigate the effect of pore parameters on vascularized bone tissue formation. An increase in pore size increases the size of blood vessels, but the increase in pore interconnectivity results in an increase of size as well as the number of blood vessels.[96] As well, pore sizes greater than 400 μm do not support vascularization at all.[96] Scaffolds fabricated with PLGA biomaterial having a small inter-pore distance along with minute pore size in the range of 5–20 μm enhances EC growth significantly.[97] Physical stiffness of a scaffold can enhance formation of focal adhesions and cytoskeletal reorganization in ECs which regulate cell migration, proliferation, and differentiation, as well as cell–cell and cell–scaffold adhesion.[98,99] A number of studies suggest that the spreading area of EC during focal adhesion increases with the increase of biomaterial’s stiffness,[100,101] however, in a stiffer biomaterial, ECs like to make cell–biomaterial interactions, rather than cell–cell interaction that cause failure to form vascular network. On the other hand, ECs take an elongated morphology in compliant material whose Young’s modulus is around 1 kPa, augment cell–cell interactions instead of cell–biomaterial interaction [102], which result in the formation of self-assembled vascular network.[100] 4.1.2.1. Rapid protyping technique and miscellaneous. To promote vascularization within tissue engineering construct, two major factors are to be considered during scaffold fabrication, one is well-interconnected microchannels or micropores, and the other is controlled positioning of vascular cells. Some studies suggest that incorporation of channel or grooves into scaffold can promote the formation of vascular network as ECs require aligned organization to change their function and phenotype to form

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Journal of Biomaterials Science, Polymer Edition

697

capillaries.[103,104] More specifically, the depth and topography of microgrooves has profound effect on cell alignment and functionality. Uttayarat et al. reported that maximum alignment of bovine aortic ECs can be obtained on fibronectin-coated 1 μm deep microgrooved surface, while Hu and colleagues concluded that microgrooves having wavy surface display better cell attachment, alignment, and survival compared to those on the rectangularshaped microgrooved surface.[104,105] To construct functional complex tissue with vasculature, different types of cells should be seeded in an organized way as per their need and function, rather than in a random fashion. Scaffolds fabricated with conventional techniques, such as gas foaming, electrospinning, salt-leaching, porogen melting, molding, fiber deposition, and freeze-drying, display uneven, uncontrolled, and undesired pore size, shape, wall thickness, pore interconnectivity, and morphology.[106] To fabricate the complex vascular network layer by layer in a controlled and precise manner, CAD-based rapid prototyping (RP) techniques, such as 3D bioplotting, inkjet printing, laser-based biofabrication, and stereolithography, can be used. However, in many cases, acellular structures were seen to be fabricated by RP technique to keep cells free from the effect of unfavorable fabrication conditions (e.g. heat, pressure, light, adhesives, cytotoxic solvents, or molding). One major flaw of acellular approach is the inefficient and inhomogeneous postfabrication cell incorporation. Yet, a number of studies have been accomplished to fabricate intricate microvascular pattern with RP technique. Of several approaches, one promising approach was to incorporate sacrificial filaments into scaffold by 3D bioplotter to get complex and well-interconnected vascular pattern (Figure 6). Lewis group demonstrated the effectiveness of fugitive ink to create 3D microvascular network by direct write assembly method.[107] In another study, they created vascular pattern by direct write method using fugitive ink and encapsulated the whole 3D network into gelatin methacrylate gel (GelMA). Then HUVECs were injected after evacuation of fugitive ink. HUVECs were found to attach, proliferate, and differentiate to form vascular network.[108] To avoid cytotoxic organic solvents, and sacrificial template-removal associated extreme temperature or pressure, Wang et al. encapsulated sodium alginatebased sacrificial vascular networks fabricated with 3D bioplotter into gelatin hydrogels and then dissolved the sacrificial pattern with EDTA solution. HUVECs were injected into evacuated channel and found to form vascular lumens and network.[109] Another research group, Chen and colleagues fabricated a patterned 3D network composed of sacrificial carbohydrate glass filament by bioplotter, dispensed ECM (e.g. fibrin gels, matrigel) encapsulated cells and mixture of poly (ethylene glycol) diacrylate and

Figure 6.

3D approaches for vascular channel formation.

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acrylate-PEG-RGDS into a rectangular mold containing the lattice filament, photo-crosslinked the PEG hydrogel to encapsulate the 3D network as well as ECM embedded cells, and then dissolved the lattice with cell culture media to obtain the interconnected empty vascular lumen. HUVECs were then seeded in the vascular lumen, and perfused with blood under high-pressure pulsatile flow. Incorporated vascular pattern was able to sustain the metabolic function of primary rat hepatocytes in the core of a thick, densely populated tissue constructs.[110] In another study, to access the effect of complex vascular geometry on diffusion gradient, they prepared vascular channel structure in gelatin gel by photolithography, encapsulated the microfluidic network and therapeutic cells within collagen gel, flushed gelatin gel by external flow, and then injected HUVECs into the vascular lumen. Injected HUVECs formed confluent monolayers within the open microfluidic channels after attachment, alignment, and proliferation. In addition, the relationship between the geometry of vasculature and spatial diffusive gradients which is responsible for angiogenic sprouting was also demonstrated.[111] To avoid the necrosis of large cell population in a thick tissue, vascular pattern design parameter (e.g. vessel alignment, branching frequency and angles, and tortuosity) is required to be optimized to ensure efficient convective diffusional mass transfer (e.g. nutrients, metabolic wastes, gases, AFs, etc.) between cells and culture medium or blood during in vitro or in vivo culture.[112] Therefore, several mathematical modeling and computational studies have been accomplished to optimize the architecture of vasculature considering the diffusional mass transfer, microfluidic behavior, and physiological data of human microvasculature.[113] Controlled positioning of vascular cell into scaffold can be achieved through the deposition of cell-laden hydrogels. Polymer hydrogels offer structural support and 3D hydrated environment similar to in vivo condition for EC attachment, proliferation, and differentiation as well as keep ECs free from fabrication-induced high shear forces.[114] To maintain EC viability and function in culture, hydrogel crosslinking and scaffold fabrication process should be biocompatible. Several studies frequently applied stereolithography,[115] inkjet printing [116,117], and 3D bioplotting [118,119] techniques to prepare cell (e.g. ECs, SMCs) laden hydrogel to fabricate vascular pattern. Besides, precise cell positioning, microchannel or interconnected porous structure is required to avoid necrosis. To incorporate of EC-laden gel into microchannel, in a study, liquid collagen containing EC suspension was distributed above micropatterned poly (dimethylsiloxane) PDMS templates, the templates were then centrifuged to move ECs into the channels, and then collagen was crosslinked to form gel. It was reported that EC formed capillary tubes after culture with VEGF and bFGF over 24–48 h.[120] While microfabrication offers many promising features to pattern microcapillary network, time delay, use of nondegradable hydrogel, and handling complexity limits its application. Therefore,an alternative way, such as mechanically removable spacer (e.g. aligned array of wires), was investigated to create microchannel into hyrogel matrix. Wray et al. used mechanical spacer to generate unbranched microchannel ranging from 152 to 787 μm in diameter into silk scaffolds, and then they seeded human arterial endothelial cells (hAECs) into the hollow channels along with bioactive agents. The formation of a contiguous layer of hAECs around the channel wall was reported seven days after cell seeding.[121] In addition, to ensure uniform cell seeding into incorporated microchannels, spacers can also be used to transfer self-assembled cell layer into hydrogel matrix. In a study, casted GelMA over EC-coated micrometric gold rods positioned in a culture chamber was photo-crosslinked, an electrical potential was applied to transfer the self-assembled layer, and then rods were removed to leave

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Journal of Biomaterials Science, Polymer Edition

699

behind the EC-assembled layer into the matrix. The same procedure was followed to prepare and transfer double-layered cell and gel mixture to combine microvessel stabilizing 3T3 fibroblast cells with HUVECs.[122] Beyond non-sacrificial filaments, microfluidic/micromolding techniques were also investigated to fabricate sacrificial pattern in a microscale. In a study, sacrificial gelatin meshes were prepared by micromolding in patterned PDMS stamp. Hydrogels (e.g. fibrinogen, Type I collagen, matrigel) were used to encapsulate the sacrificial pattern, and then successive melting and flushing were done to empty the interconnected channels. The viability of injected HDMECs into microchannel was excellent as HDMECs were seen to attach, spread, and proliferate.[123] Regardless of tremendous success with sacrificial templates, dissolutionassociated cytotoxic reaction limits their applications.[124] In an effort to fabricate sacrificial templates-free scaffold, Bertassoni and colleagues bioprinted agarose fibers, casted PEGDA hydrogels over the bioprinted templates, and then photo-polymerized them. Removal of bioprinted templates was easy, as agarose didn’t adhere with casted hydrogels. With this approach, a perfusable network could be incorporated into hydrogel matrix without producing any dissolution-related cytotoxicity.[125] Aside from sacrificial or removable filaments, laser scanning lithography (LSL) was investigated to create extremely precise micropattern on photo-sensitive hydrogel surface. In a study, West group covalently incorporated RGDs and VEGF on LSL-generated micropatterned poly (ethylene glycol) hydrogel surface. Tubules like formation in the micro-patterned area were observed within two days, whereas no tubules formation was recognized by day two for cells cultured on wide patterned lines.[126] As blood perfusion is very significant to regulate vascular pattern, West and colleagues combined microfabrication and self-assembly technique to study macro – micro-scale transport to achieve biomimetic perfusion in vitro. They prepared PDMS housing with fluid access ports by soft lithographic techniques, injected PEG hydrogel containing the HUVEC-10T1/2 cell mixture into the PDMS housing, photolithographically crosslinked the cell-laden hydrogel, and then initiated channel perfusion with normal physiologic flow rates. The perfusion between the fabricated microchannel and self-assembled vascular network significantly facilitated the convective diffusional mass transport.[127] Moreover, significant success for in vivo vascularization was reported while VEGF-mimetic peptide-bound PEGDA matrices were implanted into mice.[128] Besides microfabrication, Garicia and colleagues studied functionalized hydrogel to promote vascularization in vivo. They engineered photopolymerized PEG diacrylate hydrogel matrices containing protease-degradable sites, cell-adhesion motifs (e.g. arginine–glycine–aspartic acid), and growth factors (e.g. VEGF) and implanted subcutaneously in male Lewis rats. They reported that a significant number of vessels formed into the matrices at two weeks and the network became more densed at four weeks due to cell-demanded sustained release of VEGF over two weeks.[129] As hydrogel density regulates the mechanical strength, Putnam and colleagues investigated the effect of elevated fibrin gel density on vasculature formation in vitro and in vivo. Their findings suggest that inhibition of vasculature formation in vivo due to elevated fibrin matrix density can be partially lessened by co-delivery of mesenchymal stem cells (MSCs) with human umbilical vein endothelial cells (HUVECs) into hydrogel.[130] 4.1.2.2. Microfabrication technique. Implanted, biodegradable tissue engineering scaffolds containing cells and growth factors successfully vascularize thin tissue (1–2 mm). However, vascularization of thick tissue constructs (>2 mm) is still challenging as it takes time for blood vessels to form and anastomose with the host blood supply. To

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solve this issue, a microvascular network can be established by microfabrication within the implantable scaffold in vitro.[131] Microfabrication and bio-microelectromechanical techniques, are attracting attention due to their spatial resolutions of less than 10 μm, [132] a substantial improvement over conventional scaffold fabrication techniques such as solvent casting and porogen leaching,[133] gas foaming [134], and three-dimensional printing.[135] However, some parameters, such as selection of fabrication material, oxygen concentrations in the microenvironment [136], and fluid shear stresses within the microfluidic scaffold [137,138], significantly affect the success and applicability of microfabrication techniques. Microfabrication generally involves preparation of lithographic mask to pattern the blood vessel network according to a model,[113] formation of a master mold by thick photoresist or plasma etching, casting the desired biopolymer into the master mold, incorporation of a sacrificial layer to facilitate film removal from the master mold, and curing the films under vacuum (Figure 7). Single-layer microfluidic networks are then stacked together to form 3D scaffolds with complex vascular microchannels before seeding with ECs in a bioreactor.[132] Real vascular networks consist of blood vessels having various diameters. The direct-write laser technique can facilitate the precise fabrication of multi-depth channels mimicking the in vivo structure.[139] Synthetic nondegradable materials, such as poly dimethyl siloxane or silicon, are often employed for microfabrication.[132] However, biodegradation and biocompatibility are important issues related to the integration of the vascular construct into the host tissue as well as the potential for toxicity or inflammatory responses. The biodegradable polymers PLGA,[140] poly glycerol sebacate (PGS) [141], and silk fibroin [142] were investigated for their suitability in constructing microfluidic networks by microfabrication. PLGA exhibits rigid mechanical properties,[134] undesirable bulk degradation kinetics,[143] limited biocompatibility [144], and cytotoxic degradation byproducts, [145] all of which limit the applicability of PLGA to microfluidic scaffolds. PGS offers superior mechanical properties as well as better cellular response and morphology compared to PLGA.[143,144] PGS is inexpensive and attractive in terms of synthesizing and processing simplicity to form layers up to 100 μm.[146] A third possibility,

Figure 7.

Microfabrication technique for vascular network formation.

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chemically functionalized silk fibroin, is a promising biopolymer for microfabrication of vascular networks [142] because it offers improved cellular adhesion, controllable degradation, and enhanced mechanical property compared to PGS films. Alternative to multiple-layer stacking techniques, lithographic processes and sacrificial gel encapsulation methods can be applied to generate 3D microfluidic networks into hydrogels (alginate, collagen, fibrin, etc.) that can later be seeded with ECs for vascular tissue engineering. A lithographic process has been used to form microfluidic channels within an alginate scaffold seeded with chondrocytes, and these channels appear to support long-term cell survival and convective diffusion of soluble factors.[147] Examples of sacrificial gel techniques include a study where Type I collagen was used to encapsulate patterned Matrigel and defined shaped microcavities were achieved within collagen after the digestion of matrigel by enzymes.[148] Similarly, microfluidic channels as narrow as 6 μm were formed in collagen and fibrin gels using embedded sacrificial gelatin meshes. The channels generated in this way facilitate the transport of macromolecules into the channels and from channels into the gel matrix.[123] 4.1.2.3. Modular assembly. This sophisticated tissue engineering approach achieves formation of macro tissue constructs through the assembly of smaller modules. Micro modules can be prepared by self-assembled aggregation,[149] microfabrication of cellseeded hydrogels, [150]creation of cell sheets [151], or direct printing.[152] To promote vascularization, micro modules containing target tissue cells have been coated with ECs and perivascular cells prior to assembly.[153] The small size of the micro modules facilitates the transport of oxygen and metabolites even if they are seeded with high cell density.[154] The surface of the modules can also be modified by coating with ECM proteins. For example, fibronectin coating of collagen modules implanted in immunodeficient mice increases human vascular EC survival as well as blood vessel formation.[56] To form macro tissue, a number of methods can be adapted to assemble the micromodules, such as random packing,[155] stacking of layers,[156] or directed assembly.[157] In the random packing approach, EC-coated modules are packed together into a larger chamber and perfused with blood or culture medium which facilitates the formation of interconnected channels among the interstitial spaces between modules (Figure 8). Antithrombogenic ECs assemble to form vascularized tissue, and promote functional perfusion by delaying clotting time and inhibiting loss of platelets.[153] Incorporation of stem and progenitor cells along with ECs can significantly affect the stabilization of blood vessels that develop when using the modular approach. For example, implanted collagen gel modules coated with rat aortic ECs into Sprague-Dawley rats caused the formation of blood vessels within the first seven days. The nascent blood vessels eventually became mature and then lasted at least for 60 days. In these simple constructs, the new blood vessels connected with the host vasculature, yet some of them were leaky.[158] Interestingly, addition of bone marrow-derived stem cells to the EC coated modules lead to formation of blood vessels having similar density, but less leakiness.[159] However, the random-packed modular approach causes lack of mechanical integrity in the resultant macro tissue and limits the use of secondary cells other than ECs. Sequential assembly is a more directed method used to construct large tissue from modular tissues with a specific microarchitecture.[160] In this approach, microgels containing specific architectural designs are assembled in a controlled fashion to connect

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Figure 8.

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Modular assembly technique for vascular network formation.

the micro channels of each microgel, resulting in the formation of a bifurcating and interconnected network.[112] Thus, sequential assembly provides better control over the relative spatial arrangement of the building blocks. For example, concentric PEG microgel building blocks embedded with network of microchannels were assembled sequentially by photo-crosslinking to achieve a natural blood vessel-like structure. The inner layer was seeded with HUVECs, and the outer layer was coated with SMCs to achieve a tube-like structure using a mineral oil immersion technique.[1] Formation of vascular network by 3D fabrication technique has been summarized in Table 3. 4.1.2.4. Comparison among different fabrication techniques. It is well established that oriented pores or microchannels are required to maintain cell viability in tissue engineering construct. In previous section, it was already stated that 3D fabrication techniques are associated with unfavorable conditions which led researchers to fabricate acellular construct. However, post-fabrication cell seeding into scaffold demonstrates many complexities in terms of efficient and homogeneous cell positioning. In contrast, fabrication of cellular scaffold facilitates the controlled and homogeneous cell distribution. Compared to other fabrication material, hydrogels are more suitable to fabricate cellular scaffold as it can ensure cell viability by providing in vivo like environment. However, lack of mechanical strength limits the applications of hydrogel scaffold. In addition, free radical photo polymerization or chemical crosslinking to polymerize hydrogels can negatively affect the viability of encapsulated cells. Channel, which is the key parameter to promote mass transport and directional growth of EC, can be introduced into the 3D scaffold by using sacrificial and non-sacrificial filaments.

Sacrificial filaments/pluronic F127 Mechanical spacer

3D Bioplotter

5% gelatin filament, channel 400–700 μm Micropatterned cells

3D Robotic dispensing Laser-guided direct writing Photolithography and and assembly of structures

Multi-level interconnected lumens

Chemical

Non sacrificial agarose filaments Hexagonal/woodpile micropattern

3D Micromolding

Stereolithography

Photo, thermal Photo

Sacrificial lattice /sodium alginate

Bulk polymerization

Photo

Thermal

Ionic, photo, thermal, enzymatic Thermal, ionic

Sacrificial filaments/carbohydrate glass

Bulk polymerization

Thermal

Agarose spacer

PEGDA

3–9 mg/mL collagen Matrigel

10 or 15% GelMA

10% GelMA

Gelatin, agarose and collagen

Multicellular spheroids, diameter 300– 500 μm Alginate, fibrin, pegda, matrigel

GelMA

GelMA

Photo Photo

Scaffold material

Gelation method

3D Bioplotter

Bulk polymerization

Channel/pore/pattern/ spacer

Scaffold fabrication techniques for vascularisation.

Fabrication technique

Table 3.

In vitro

HUVECs

HUVECs, SMCs/ encapsulation

HUVECs, VEGF

HUVECs/ immersion and agitation HUVECs/mixing

HUVECs

In vitro

In vitro

In vitro

In vitro

In vitro

In vitro

In vitro

CHO, HUVSMCs, HSFs

HUVECs

In vitro

In vitro

HUVECs

HUVECs

Additives/addition Model/target technique site

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[1]

[331]

[330]

[329]

[125]

[109]

[110]

[328]

[122]

[108]

Ref

(Continued)

90% cell viability after two days, in vivo like vessel formation

Confluent layer on 3 mg/ml collagen after three days Elongated structures after 24 h

Confluent layer by day four

Confluent layer on gelatin and collagen, spheroids on agarose after 3–4 days Confluent layer by day seven

Confluent layer within one day

Confluent lumen of 10 mm long and 618 ± 15 μm diameter, stable geometry for 15 days Spheroids fused within 5–7 days to form vascular tubes of 0.9– 2.5 mm diameter

Formed confluent layer, viability >95% after 48 h

Channel confluence/lumen formation

Journal of Biomaterials Science, Polymer Edition 703

Photo, thermal Photo Photo Ionic

Micropatterned

Microchannels 50– 1000 and 15–50 μm

200 μm diameter channels

LSL

Soft lithography and photolithography CO2 laser engraving system

Gelation method

Photolithography Micropatterned (microfabrication)

Channel/pore/pattern/ spacer

(Continued).

Fabrication technique

Table 3.

Alginate, alginate-sulfate

PDMS, PEG

PEG, PEGDA

PDMS/2.4 mg/ mL collagen gel

Scaffold material HUVECs, bFGF, VEGF/immerse and centrifuge HUVECs, RGDS and VEGF RGDS, HUVEC and 10T1/2, injection HUVECs, RGD/ HBP Mice

In vitro

In vitro

In vitro

Additives/addition Model/target technique site

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High vessel density after eight weeks

lumens form in two days for