Bioprinting of Small-Diameter Blood Vessels

Xia Cao , Sushila Maharjan , Ramla Ashfaq , Jane Shin , Yu Shrike Zhang

Engineering ›› 2021, Vol. 7 ›› Issue (6) : 832 -844.

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Engineering ›› 2021, Vol. 7 ›› Issue (6) :832 -844. DOI: 10.1016/j.eng.2020.03.019
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Bioprinting of Small-Diameter Blood Vessels
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Abstract

There has been an increasing demand for bioengineered blood vessels for utilization in both regenerative medicine and drug screening. However, the availability of a true bioengineered vascular graft remains limited. Three-dimensional (3D) bioprinting presents a potential approach for fabricating blood vessels or vascularized tissue constructs of various architectures and sizes for transplantation and regeneration.
In this review, we summarize the basic biology of different blood vessels, as well as 3D bioprinting approaches and bioink designs that have been applied to fabricate vascular and vascularized tissue constructs, with a focus on small-diameter blood vessels.

Keywords

Bioprinting / Small-diameter blood vessel / Bioink / Vascular engineering

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Xia Cao, Sushila Maharjan, Ramla Ashfaq, Jane Shin, Yu Shrike Zhang. Bioprinting of Small-Diameter Blood Vessels. Engineering, 2021, 7(6): 832-844 DOI:10.1016/j.eng.2020.03.019

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1. Introduction

Blood vessels are the conduits that transport oxygen and nutrients to the tissues and carry carbon dioxide and wastes from them. Arteries, capillaries, and veins are three main types of blood vessels, which differ in structural composition, dimension, and physiological function [1]. Based on the inner diameter (ID), blood vessels are broadly categorized into microvessels, having an ID less than 1 mm, small vessels, having an ID ranging from 1 to 6 mm, and large vessels, with an ID greater than 6 mm[2–3].

The aorta is the largest blood vessel in the human body; it transports oxygenated blood from the heart to different parts of the body. The aorta branches into arteries, which further divide into smaller vessels known as arterioles. The arterioles eventually branch into tiny capillaries; these are the smallest blood vessels, which allow the diffusion of oxygen, nutrients, carbon dioxide, and other metabolic wastes between the blood and the local tissues[4–5]. Capillaries form a network between the arterioles and venules, the latter of which are very small veins that collect deoxygenated blood from the capillaries and transport it into progressively larger veins until it re-enters the heart [6].

Although arteries and veins have different functions, their walls similarly consist of variable compositions of cells and extracellular matrix (ECM), which are grouped into three main layers: the innermost tunica intima, the middle tunica media, and the outer tunica adventitia (Fig. 1(a))[7–8]. Capillaries (ID < 10 μm) consist of a single layer of endothelial cells (ECs) forming the inner lining of the blood vessel (tunica intima), and are surrounded by basal lamina and pericytes (Fig. 1(b))[9,10]. Both arterioles and venules (ID = 10–100 μm) consist of an inner EC layer (tunica intima) with a surrounding thin smooth muscle cell (SMC) layer (tunica media) (Fig. 1(b)). Arteries and veins (ID > 100 μm) consist of an inner EC lining (tunica intima), a middle dense SMC layer (tunica media), and an outer layer of fibroblasts and ECM components (tunica adventitia) (Fig. 1(b))[10,11]. Different layers of blood vessels with their cell composition, diameter, ECM components, and functions are listed in Table 1[12,13].

Fig. 1. The general structure and types of blood vessels. (a) Schematics of different layers of an arterial and venous wall. Reproduced from Ref. [8] with permission of Mary Hammes, © 2015. (b) Schematics of different types of blood vessels showing cell composition, diameters, and thickness. The bar in the table represents the relative proportion of different layers of blood vessel. A capillary comprises an EC lining surrounded by pericytes. An arteriole/venule consists of an EC lining surrounded by a thin layer of smooth muscle cells (SMCs). An artery/vein consists of an EC lining surrounded by a dense SMC layer and fibroblasts. Reproduced from Ref. [10] with permission of Nature, © 2018; and Ref. [11] with permission of the American Physiological Society, Inc., © 1954.

Table 1 The general structures and functions of blood vessels[12,13].

ECs and vascular SMCs are the key cellular components of both arteries and veins. The interactions of these cell types play a critical role not only in the development of the vasculature, but also in maintaining vascular homeostasis [14]. The EC linings of blood vessels are significantly heterogeneous in phenotype and function, due to the varying microenvironments and different levels of pressures and shear stresses that they experience during blood flow [15]. The EC linings serve as tightly regulated dynamic barriers, preventing blood leakage into the surrounding tissue and interstitial space. They also play pivotal roles in several physiological processes, including regulation of blood flow and permeability, transendothelial migration of circulating immune cells at the site of inflammation, and maintenance of vascular tone and hemostasis[16,17]. While ECs are aligned longitudinally, SMCs are circumferentially aligned and interspaced by elastic fibers. SMCs possess the contractile phenotype and maintain vascular homeostasis through active contraction and relaxation. In addition, SMCs possess a secretory phenotype, which is responsible for the synthesis and repair of ECMs and thus regulates the structure of the vascular wall [18].

Coronary heart disease and peripheral arterial disease represent the leading causes of mortality and morbidity globally, with an annual mortality incidence estimated to increase to 23.3 million by 2030 [19]. Angioplasty, stent implantation, and surgical bypass grafting are the current commonly used revascularization approaches [20]. The use of autologous arteries or veins, mostly the internal thoracic artery or saphenous vein, remains the gold standard for bypass grafts for the treatment of vascular diseases such as coronary artery and peripheral vascular diseases [20]. However, healthy autologous vessels are not always available, require invasive harvesting, and have poor long-term patency rates[21,22]. Synthetic vascular grafts, which are mostly made from polyethylene terephthalate or polytetrafluoroethylene, have been used as alternatives to autologous vessels for the successful replacement of medium and large vessels[23,24]. Although these synthetic grafts have proven effective for medium and large vessels with long-term patency, their adoption has been limited for small vessels due to mismatched biomechanical properties, poor biocompatibility, thrombogenicity, microbiological contamination, and poor patency rates[25–27]. Considering the limitations of current vascular conduits and the pathological events in smalldiameter blood vessels including coronary arteries and peripheral arteries, there is a persistent clinical demand for bioengineered vascular conduits with the potential to reduce the morbidity associated with small-diameter vascular disease [28].

With the progress that has been achieved in vascular bioengineering, three-dimensional (3D) bioprinting has emerged as a potential approach for generating blood vessels with the ability to grow, remodel, and repair vascular disease in vivo. Different 3D bioprinting techniques and strategies have been investigated and established over the past years to fabricate biomimetic blood vessels and are constantly being refined and improved[29–32]. In this review article, we summarize different bioprinting techniques, including extrusion bioprinting, inkjet bioprinting, laser-assisted bioprinting, and vat polymerization-based bioprinting, applied to the engineering of small-diameter blood vessels, including small vessels and microvessels, along with their strengths and limitations. We also discuss different biomaterials and their properties relating to biocompatibility, printability, and mechanical properties, as well as the challenges involved in the formulation of appropriate bioinks that satisfy the critical requirements for the 3D bioprinting of small-diameter blood vessels. We conclude with potential future directions.

2. 3D bioprinting

3D bioprinting has significantly advanced in the field of vascular tissue engineering. The essential requirements for engineering blood vessels with long-term patency include appropriate mechanical properties, high compliance, remodeling capability, and anti-thrombogenicity, in order to achieve successful integration with the native blood vessels [33]. The requirements for ideal bioengineered vessels—particularly small-diameter blood vessels— are listed in Table 2 [25]. 3D bioprinting permits the precise deposition of biomaterials and living cells (together termed as ‘‘bioink”) in well-defined spatial patterns that closely mimic the structures of blood vessels [34]. Vascular tissue engineering through bioprinting has been accomplished via various modalities and strategies, along with a wide range of bioinks [24]. The most commonly used bioprinting modalities include inkjet [35], extrusion[36,37], laserassisted [38], and vat polymerization-based bioprinting[39,40]. The inkjet method, which is a non-contact bioprinting technique, deposits bioink droplets of a predetermined size from a cartridge to generate pseudo-3D or 3D structures (Fig. 2(a))[31,41]. Extrusion bioprinting is the most commonly used bioprinting method, in which the bioink is dispensed continuously in the form of filaments through a nozzle (Fig. 2(b))[31,42]. Laser-assisted bioprinting is comparatively less common than inkjet and extrusion and uses laser pulses to deposit the bioink from the donor slide onto the receiver substrate (Fig. 2(c))[31,43,44]. Vat polymerization-based bioprinting, typically in the form of stereolithography or digital light processing, uses precisely controlled patterns of light to polymerize photosensitive polymers on a vertically movable platform to create 3D constructs with desired architectures (Fig. 2(d))[45,46].

Table 2 Requirements for an ideal bioengineered small-diameter blood vessel. Reproduced from Ref. [25] with permission of Valentina Catto et al., © 2014.

Each modality has its strengths and limitations and differs in several parameters such as choice of bioinks and substrates utilized, complexity of patterns, and yield of fabrication [47]. Inkjet bioprinting offers several advantages including rapid bioink deposition and high resolution; however, it often has limited capacity due to instability of the dispensed layers[48,49]. Extrusion bioprinting is a cost-effective method with ease of control. However, it is a comparatively slow method and has lower resolution [50]. Laser-assisted bioprinting provides rapid fabrication, high resolution, and biocompatibility but at high instrument cost [51]. Vat polymerization-based bioprinting also offers high resolution and rapid fabrication, with instrumentation of medium complexity [52]. Depending on the varying shapes and sizes of blood vessels, different 3D bioprinting modalities have been applied to fabricate vasculature with varied degrees of function in vitro (Fig. 2(e)) [5].

Fig. 2. Representative 3D bioprinting approaches. (a) Schematic of inkjet bioprinting method. (b) Schematic of extrusion bioprinting method. (c) Schematic of laser-assisted bioprinting method. (a–c) are reproduced from Ref. [44] with permission of Wiley, © 2013. (d) Schematic of vat polymerization-based bioprinting method. Reproduced from Ref. [46] with permission of Xuanyi Ma et al., ©2016. (e) Schematic showing the hierarchy of blood vessels and the current bioprinting techniques that have been employed to fabricate the range of different blood vessels. Reproduced from Ref. [5] with permission of Elsevier, © 2019.

3. Bioink designs

To be functional, the structure and integrity of bioengineered vasculature must resemble those of native blood vessels, both of which are highly dependent on mechanical and chemical cues from the surrounding ECM [53]. The ECM serves as a reservoir and modulator for various cytokines and growth factors, as well as providing structural support for EC organization and stabilization during blood vessel formation [54]. Thus, ECM properties such as density, heterogeneity, and stiffness play critical roles in regulating vascular morphogenesis, capillary network formation, and barrier integrity[55,56].

The major ECM components that comprise different layers of blood vessels are shown in Table 1[12,13]. Vessel lumens are lined with ECs, resting on a basement membrane that is largely composed of collagens, laminins, fibronectin, nidogen, heparin sulfate proteoglycans, entactin, and other macromolecules[54,57]. Tunica media, the middle SMC layer, is composed of collagenous ECM containing fibrillar collagens I, III, and V; laminin; elastin; fibronectin; hyaluronan; and decorin[58–60]. Tunica intima and tunica media are separated by the internal elastic lamina. The tunica adventitia, the outermost layer of connective tissue and fibroblasts, consists of elastin, collagen types I and IV, and versican. Tunica media and tunica adventitia are separated by the external elastic lamina [57,59–61].

To this end, the successful in vitro fabrication of blood vessels, particularly those with small diameters, requires synergy between higher-resolution bioprinting approaches and adequate bioink formulations that can provide structural and mechanical support to maintain cell viability while facilitating grafting or neovascularization [62]. A bioink is a combination of biomaterial(s), cells, nutrients, and/or growth factors that should mimic the optimal ECM environment of the tissue [63]. The design of a bioink should consider major ECM components, and is critical for the 3D bioprinting of vessels as it determines not only the bioprinting resolution but also vessel integrity and other critical biological properties[64–66].

The most popular formulation of bioink for 3D bioprinting of blood vessels is based on hydrogel precursors, due to their excellent biocompatibility, tunable stiffness and porosity, and ability to mimic native ECM, as well as their compatibility with different bioprinting modalities [67]. Several natural and synthetic biomaterials have been investigated, alone or in combination, for designing bioink that resembles the physicochemical properties of the ECM. Natural biomaterials that have been widely used for in vitro vascular tissue engineering include gelatin[68–71], collagen[72–77], elastin[78–81], fibrin[82–85], hyaluronic acid[68,86,87], agarose[49,88], alginate[89,90], and Matrigel[91–93], among others. Natural biomaterial-based hydrogel provides a suitable microenvironment for cell adhesion, growth, and proliferation; most importantly, it does not typically cause chronic inflammation or toxicity to the host [74]. However, the weak mechanical strength of naturally derived hydrogels fails to withstand physiological pressure and thus limits their usage in vascular tissue engineering [94]. Therefore, various synthetic polymers have been investigated, in combination with natural biomaterials, for such applications because of their precisely controlled mechanical properties, porosity, reproducibility, structural diversity, stiffness, and biodegradation [95]. Commonly used synthetic polymers include poly(ethylene glycol) (PEG)[96,97], poly(hydroxyethyl methacrylate) [98], and poly(vinyl alcohol) (PVA) [99,100]. These composite materials can provide good mechanical support and increase the mechanical strength of printed structures. However, the application of synthetic polymers is limited due to their lack of biocompatibility, poor cell adhesion, release of toxic byproducts, and decrease in mechanical properties due to the degradation process [47].

To overcome these limitations and obtain bioinks with desirable properties, functionalization has been explored using different methods based on chemical, mechanical, physical, and biological modifications. One of the most commonly adopted functionalization methods is the introduction of methacryloyl groups to polymers such as gelatin [101–105], collagen [106,107], and hyaluronic acid [108,109]. This method results in photopolymerizable polymers that can form mechanically stable constructs when needed. The mechanical strength of these polymers depends largely on the degree of methacryloyl modification and light exposure, with higher degree of methacryloyl modification and longer light exposure leading to increased stiffness of the constructs and decreased degradation [110]. The application of two or more crosslinking mechanisms contributes significantly to increased structural integrity and mechanical strength [110,111]. For example, improved mechanical properties were achieved when a decellularized ECM (dECM) bioink was covalently crosslinked using vitamin B2 (riboflavin) [112], followed by thermally triggered physical crosslinking via collagen fibrillogenesis [113]. This twostep crosslinking mechanism demonstrated a superior storage modulus compared with constructs crosslinked separately, indicating that each crosslinking mechanism contributes to the overall mechanical properties of the constructs [113]. On the other hand, combining bioink materials may achieve the same phenomenon. For example, the addition of gelatin to gelatin methacryloyl (GelMA) increased the degree of crosslinking and thus the mechanical properties [114]. In addition, when other polymers, such as methylcellulose [115], 4-arm PEG-tetra-acrylate (PEGTA) [110], 8-arm PEG-acrylate with a tripentaerythritol core (PEGOA) [116], hyaluronic acid [117], PVA [118], or hydroxyapatite [119], are incorporated, they can enhance printability and/or improve shape fidelity post-bioprinting.

Furthermore, various bioactive materials such as fibrin (or fibrinogen and thrombin) and polylysine have been added to bioinks, for the induction or enhancement of their bioactivity. Fibrin, a polymer of fibrinogen that forms fibrous, viscoelastic, and porous hydrogels in the presence of thrombin, was shown to improve the bioadhesiveness and bioactivity of bioink [120]. Positively charged polylysine has also been used to enhance cell adhesion by increasing the electrostatic interactions with negatively charged cell membranes [121].

Similarly, the biochemical properties of non-adhesive hydrogels have been improved by their modification with adhesive peptide ligands. The arginine-glycine-aspartic acid (RGD) peptide is the most commonly used peptide ligand that plays a pivotal role in cell adhesion, spreading, and migration [122]. For example, PEGdiacrylate (PEGDA) hydrogels conjugated with RGD peptides enhanced the adhesion of cells as compared with those without RGD modifications [123]. It was further demonstrated that the inclusion of RGD peptides on implanted PEGDA hydrogels regulated cell adhesion, inflammation, fibrous encapsulation, and vascularization, in vivo [123]. RGD peptides have also been introduced to various other hydrogel-forming polymers, such as hyaluronic acid [123], alginate [124], chitosan [125], and PEG [126] hydrogels, to improve cell behaviors.

Several growth factors such as fibroblast growth factor (FGF), vascular endothelial growth factor (VEGF), and transforming growth factor-α (TGF-α) and TGF-β have been identified as the inducers of angiogenesis. For example, FGF-2 [127] and VEGF [128] are the key mediators of blood vessel formation that stimulate the proliferation, motility, and differentiation of ECs [129]. VEGF was physically entrapped within PEG hydrogels for release in response to proteases secreted by the cells, leading to enhanced vessel formation in both in vitro and in vivo studies [130]. Moreover, several growth factor-binding sites have been discovered within ECM proteins such as fibronectin [131], fibrinogen [132], tenascin C [133], and vitronectin [134]. VEGF bound to these ECM proteins has been shown to improve healing of diabetic wounds and bone defects [135]. While some of these strategies may have not been combined with bioinks for vascular bioprinting, their adaptation would seem to be natural.

More recently, several studies have shown the effect of altered substrate viscoelasticity on cell behaviors, including tuning the loss modulus of acrylamide hydrogels by altering the covalent crosslinking and/or concentration of the polymer [136,137], modulating stress relaxation in PEG hydrogels by altering covalent crosslinkers with different affinities [138], or using the covalent and/or physical crosslinking of alginate hydrogels [139]. These studies showed that an improved loss modulus and substrate creep enhanced the cell spreading and osteogenic differentiation of MSCs in twodimensional (2D) culture [136,137], and that enhanced substrate stress relaxation improved the spreading and proliferation of cells in 2D [139] and allowed cells to adapt to physiologically relevant morphologies in 3D culture [138]. Similarly, it was demonstrated that cells behaved differently in elastic and viscoelastic hydrogels and that the relaxation behaviors of the biomaterial changed with the interaction of cells, altering cell behaviors including spreading, proliferation, and differentiation [140]. MSC differentiation was found to be dependent on the initial elastic modulus of the hydrogel in 3D, whereas it lost sensitivity to hydrogel stiffness in elastic hydrogels, indicating the significance of stress relaxation in cells with respect to mechanical cues in the ECM [141]. These insights may be helpful in vascular bioink designs for improving cell behaviors and vessel functions.

4. Current advances in the bioprinting of small-diameter vessels

Small vessels include the arteriolar and venules. The flow of blood through the capillaries is regulated by the arteriolar lumen. Venules are the smallest veins that receive blood from capillaries and play an important role in the exchange of oxygen and nutrients between the blood and tissue [4]. It is typically difficult to construct small-diameter blood vessels, including small vessels and microvessels, using a manual approach. However, with the advantages of bioprinting, more sophisticated and physiologically relevant structures of small-diameter blood vessels may be obtained [46,142].

4.1. Extrusion bioprinting

Extrusion bioprinting is a cost-effective and commonly used method of bioprinting, which dispenses the bioink from a printhead in a way that is synchronized with the printhead’s spatial movement in a layer-by-layer manner [47,143,144]. Extrusion bioprinters dispense bioinks continuously through the nozzle pneumatically or mechanically, and are compatible with a wide range of bioinks with high viscosities [145]. Alginate has been extensively used in the extrusion bioprinting of vascular tissue constructs. For example, alginate can be physically crosslinked with simultaneously delivered calcium chloride (CaCl2) solution in the core, to ensure the structural fidelity of the bioprinted hollow constructs [89,146]. Perfusable vasculature conduits were bioprinted with primary human umbilical vein SMCs (HUVSMCs) encapsulated in sodium alginate solution (3%, 4%, and 5% w/v in deionized water) using a coaxial nozzle, fabricated with an outer needle (ID = 540 μm) and a inner needle (ID = 250 μm) [147]. The average vascular conduit and lumen diameters bioprinted with sodium alginate solution (5% w/v) were (1449 ± 27) and (990 ± 16) μm, respectively. Similarly, 3D bioprinting of hollow calcium alginate fibers of uniform diameter with high mechanical strength and structural integrity was achieved with L929 mouse fibroblastsencapsulated sodium alginate (2%–5% w/v) and CaCl2 (2%–5% w/v) solutions, using the coaxial nozzle-assisted extrusion bioprinting method [148]. The coaxial assembly consisted of an outer needle with an ID of 1600 μm and three different inner needles of 510, 410, or 330 μm in ID. The size of the bioprinted hollow fibers varied depending on the size of the inner needles as well as the flow rate and concentration of sodium alginate and CaCl2. The average inner and outer diameters of the bioprinted alginate hollow fibers were found to be 892 and 1192 μm, respectively, when sodium alginate solution (2% w/v at 1 mL·min-1 flow rate) and calcium chloride (4% w/v at 2 mL·min-1 flow rate) were used.

In another study, perusable vascular tissue constructs were fabricated using a multilayered coaxial nozzle with concentric needles, and a human umbilical vein ECs (HUVECs)- and human mesenchymal stem cells (hMSCs)-encapsulated bioink [110]. The bioink in this study was composed of sodium alginate (2% or 3% w/v), PEGTA (2% or 3% w/v), and GelMA (5% or 7% w/v) prepolymer solutions. PEGTA was added to enhance the mechanical properties of the bioprinted vascular tissue constructs. Depending on the size of the inner (ID = 0.159–0.210 mm) and outer (ID = 0.153–0.838 mm) needles used, different-sized hollow tubes were bioprinted with the optimized blend bioink (3% w/v alginate, 2% w/v PEGTA, and 7% w/v GelMA). The average outer diameter (OD) of the bioprinted hollow tubes ranged approximately from 500 to 1500 μm, the ID ranged from 400 to 1000 μm, and the wall thickness ranged from 60 to 280 μm. The blended bioink was shown to support the spreading and proliferation of encapsulated HUVECs and MSCs in the bioprinted vascular constructs, thus forming physiologically relevant vascular tissue constructs (Fig. 3(a)) [110]. Recently, multilayered hollow vascular tissues were further fabricated using a multichannel coaxial extrusion system. The system consisted of a coaxial nozzle with three concentric needles (a needle in the core (ID = 0.210 mm), a needle in the middle layer (ID = 0.686 mm), and a needle in the outer layer (ID = 1.600 mm)); a bioink composed of GelMA (5% or 7% w/v), sodium alginate (2% or 3% w/v), and PEGOA (1% or 2% w/v); HUVECs to be delivered in the middle layer; and SMCs to be delivered in the outer layer [117]. When a blended bioink (7% w/v GelMA, 2% w/v alginate, and 2% w/v PEGOA) with optimal printability was used, the average diameter of the inner lumen was measured to be (663 ± 52) μm while that of the outer lumen was (977 ± 63) μm. The inner and outer wall thicknesses were found to be (62 ± 10) and (94 ± 10) μm, respectively (Fig. 3(b)) [116]. The bioprinted vascular tissues exhibited cellular heterogeneity of different layers and respective phenotypes, indicating the formation of blood vessel-like tissues.

Fig. 3. (a) Coaxial bioprinting of small-diameter blood vessels. (I) Multilayered coaxial nozzles and schematics demonstrating the fabrication of various hollow tubes. (II) Schematics of the bioprinting process and fabrication of a vascular bed. (III) Schematics and fluorescence micrographs showing bioprinted tubes with varying diameters. (IV) Fluorescence photographs of bioprinted tubes before (inset) and after the injection of red fluorescent microbeads. (V) Confocal images of a vascular tubular structure showing the expression of smooth muscle α-actin (α-SMA) by MSCs and the expression of cluster of differentiation 31 (CD31) by HUVECs after 14 d (upper row) and 21 d (lower row) of culture. Nuclei are stained with 4' ,6-diamidino-2-phenylindole (DAPI). (b) (I) Schematics showing a multichannel coaxial extrusion system for the bioprinting of a multilayered hollow tube. (II) Representative fluorescence micrographs of double-layered hollow tubes (upper row) and a cross-sectional view (lower row) of single- and double-layered hollow tubes. (III) Fluorescence micrographs of a bioprinted tube consisting of inner human urothelial cells labeled with green cell tracker and outer human bladder SMCs labeled with red cell tracker on Day 4. (IV) Confocal micrographs of immunostained vascular tubes exhibiting expression of CD31 (green) by HUVECs and a-SMA (red) by SMCs. (a) Reproduced from Ref. [110] with permission of Elsevier, © 2016; (b) reproduced from Ref. [116] with permission of Wiley, © 2018.

In a more recent study, small-diameter blood vascular constructs comprising endothelial and muscular cell layers were fabricated with extrusion bioprinting, using a triple-layered coaxial nozzle (ID = 2.159/2.906/3.810 mm) with HUVECs and human aortic SMCs (HAoSMCs) encapsulated in vascular tissue-derived ECM (VdECM, 3% w/v)/alginate (2% w/v) hydrogel as the bioink [149]. The resultant tube of approximately 2 mm in ID consisted of a thin HUVEC layer (approximately 50 μm) surrounded by a thicker HAoSMC layer (800–1000 μm). The prematured vessel was then implanted into the abdominal aorta of rat for 21 d. The vascular graft showed smooth muscle layer remodeling and integration into the host tissue, along with good patency and an intact endothelium. Similarly, using a 3D-printed microfluidic coextrusion device, long permeable alginate tubes of (265 ± 11), (360 ± 11), or (448 ± 12) μm ODs were produced, which were subsequently seeded with HUVECs and vascular SMCs (vSMCs) co-encapsulated in Matrigel (35% v/v) [150]. The resulting vascular tubes possessed the correct configuration of lumen with an inner endothelial lining and an outer sheath of SMCs. These vascular tubes exhibited functional properties of vessels, including quiescence, perfusability, and contractility in response to vasoconstrictor agents. Another study presented the coaxial extrusion printing of small-diameter blood vessels, where human coronary artery SMCs (HCASMCs) were encapsulated in a catecholfunctionalized GelMA (GelMA/C, 20 wt%) bioink [151]. The GelMA/C bioink with HCASMCs was extruded through an external larger needle (diameter: 840 μm), whereas the HUVECs in Pluronic F127 (30 wt%) and sodium periodate (23.4 nmol·L–1) were delivered through a smaller internal needle (diameter: 406 μm). Depending on the coaxial nozzles used, the IDs of the bioprinted tubes were found to be 500–1500 μm and the wall thickness ranged from 100 to 300 μm. The bioprinted vascular constructs were further implanted into nonobese diabetic-severe combined immunodeficiency (NOD-SCID) interleukin-2 (IL-2) receptor gamma null (NSG) mice, and suggested autonomous integration and functional vessel formation in vivo.

4.2. Inkjet bioprinting

Inkjet bioprinting is a fabrication method in which bioink droplets are deposited at predefined positions on a substrate. Formation of droplets may be induced by thermal or piezoelectric forces [48,152]. Alginate-based 3D tubular tissue constructs were fabricated with the sudden gelation of alginate beads in a CaCl2 solution, thus forming microgel droplets as building blocks [153]. Specifically, sodium alginate solution (0.8% w/v) was ejected into a CaCl2 solution (2% w/v) through an ink jet nozzle to form circular microgel beads of approximately 40 lm in diameter. Alginate tubes of 200 μm in diameter were fabricated by moving the inkjet printhead in a circular pattern, wherein droplets of the alginate gel were bioprinted in a bath of CaCl2.

3D straight and zigzag cellular tubes of approximately 3 mm in diameter with overhung structures were successfully fabricated by layer-by-layer deposition with a 3D inkjet bioprinting system using NIH/3T3 fibroblasts encapsulated in sodium alginate (1% w/v) as the bioink [154]. The bioink droplets were ejected through a piezoelectric nozzle dispenser (ID = 120 μm) into a CaCl2 (2% w/v) bath, forming a gelled cell-calcium alginate layer. The presented fabrication approach could potentially be applied for the bioprinting of blood vessels with complex geometries and structures. Similarly, vascular-like structures with bifurcations were generated by means of a liquid support-based inkjet bioprinting method. The bioink droplets, which were composed of sodium alginate solution (1% w/v) with or without NIH/3T3 fibroblasts, were ejected using a piezoelectric inkjet printhead (ID = 120 μm); CaCl2 solution (2% w/v) was used as the crosslinking agent. The vascular constructs were bioprinted in either a horizontal manner (Figs. 4 (a-I), 4(a-II), 4(b-I), and 4(b-II)) or a vertical manner (Figs. 4(a-III), 4(a-IV), 4(b-III), and 4(b-IV)) into a bath of CaCl2 solution (2% w/v). The mean diameter of the bifurcated vascular structure was 3 mm and the thickness of the wall was approximately 1 mm [35]. In another example, a bioink of thrombin (50 U·mL-1 ) and CaCl2 (80 mmol·L–1) was mixed with human microvascular ECs (HMVECs) and deposited into a fibrinogen substrate (60 mg·mL-1 ; so-called ‘‘bio-paper”) to form a fibrin channel of < 100 μm in diameter mimicking the microvascular structure. The bioprinted cells were aligned inside the fibrin channels and were ready for proliferation, and the channel structure of the bioprinted microvasculature were observed to remain stable for 21 d [155].

Fig. 4. Inkjet bioprinting of small-diameter blood vessels. (a) Schematics showing the inkjet bioprinting of (I, II) horizontal and (III, IV) vertical bifurcated tubular constructs in which sodium alginate droplets (1% w/v), with or without cells, were deposited on the platform and then crosslinked with CaCl2 solution. (b) (I) Top view and (II) global view of bifurcated alginate tubes generated by horizontal printing; (III) front view and (IV) global view of bifurcated alginate tubes generated by vertical printing. Reproduced from Ref. [35] with permission of Wiley, © 2015.

4.3. Laser-assisted and vat polymerization-based bioprinting

Laser-assisted bioprinting is a nozzle-free bioprinting approach that utilizes laser pulses to deposit the bioink from the donor slide onto the receiver [156]. A study demonstrated the potential of laser-assisted bioprinting for fabricating straight and Y-shaped tubular constructs using alginate (8% w/v) or alginate (2% w/v) containing mouse fibroblasts as bioinks. The average cellular Y-shaped constructs were 5 mm in diameter when optimized parameters were applied (laser fluence: 1445 mJ·cm-2 , repetition rate: 10 Hz, substrate velocity: 80 mm·min-1 , downward movement step size: 25 μm) [38]. Simple structures that mimicked the vascular networks in natural tissue were fabricated with HUVECs and HUVSMCs using laser-assisted bioprinting. Bioprinted droplets of approximately 50 μm in diameter were deposited 50–150 μm away from each other when a laser energy of 0.5–1.5 μJ·pulse–1 was used [157].

Dynamic digital micro-mirror device (DMD)-based vat polymerization bioprinting is another fabrication method with high resolution [158]. This technique enables the fabrication of highly complex tissue constructs with micrometer resolution. Volumetric tissue constructs with embedded vasculature were fabricated using hydrogels encapsulating relevant cells [159]. For example, a 3D vascularized hepatic model was fabricated with a DMDbased stereolithographic bioprinting approach in which hepatic lobules were fabricated with human induced pluripotent stem cell (hiPSC)-derived hepatic cells encapsulated in GelMA (5% w/v), followed by the fabrication of vasculature using HUVECs and adipose-derived stem cells encapsulated in glycidal methacrylatehyaluronic acid (1% w/v) and GelMA (2.5% w/v) [46]. These 3D-bioprinted liver construct, measuring 3 mm × 3 mm with a thickness of approximately 200 μm, consisted of a hexagonal array of hiPSC-derived hepatic cell lobules with surrounding layers of HUVECs and adipose-derived stem cells. This vascularized liver model provides a platform for pathophysiological and drug screening studies [46]. In another study, a pre-vascularized tissue construct were created following the same approach; three digital masks were employed to fabricate the patterned tissue constructs, with uniform (Fig. 5) [43] or gradient vascular channels forming the vasculature network [142]. The size of the vascular channels ranged from 50 to 250 μm. While the base layer of the vascularized tissue constructs consisted of hexagonal regions which was fabricated with HepG2 cells encapsulated in GelMA (5% w/v) using a digital mask with hexagonal patterns, the vascular channel region was fabricated with HUVECs and 10T1/2 murine embryo fibroblasts encapsulated in the hydrogel (2.5% w/v GelMA + 1% w/v glycidal methacrylatehyaluronic acid) using the vascular channel mask. Finally, the vascular network was covered on the top with GelMA (5% w/v) using a slab mask, thus generating a vascularized tissue construct of 4 mm × 5 mm in size with a thickness of 600 μm. The HUVECs in the bioprinted tissue construct spontaneously formed lumen-like structures in vitro. Furthermore, the bioprinted pre-vascularized and non-pre-vascularized (without cells) tissue constructs were implanted under the dorsal skin of severe combined immunodeficiency mice for 14 d and demonstrated endothelialization of the pre-vascularized tissue constructs [142].

Fig. 5. Vat polymerization-based bioprinting of microvascularized tissues. (a) Schematics showing the fabrication of pre-vascularized tissue constructs; (b) fluorescence micrograph showing bioprinted tissue constructs consisting of HUVECs (red) in the channels and HepG2 cells (green) in the surrounding hydrogel matrix; (c) 3D confocal reconstruction micrograph showing ECs in microchannels, labeled with red cell tracker and stained for CD31 in green. Reproduced from Ref. [43] with permission of Elsevier, © 2013.

4.4. Other bioprinting strategies

Besides the abovementioned bioprinting modalities for fabricating vasculature structures, sacrificial bioprinting is a classical indirect (bio)printing method. The sacrificial bioprinting of vascularized tissue constructs utilizes fugitive bioinks that are removed after the completion of the bioprinting, usually by physical extraction or changing the temperature, leaving behind the perfusable channels [62]. Cylindrical agarose fibers were bioprinted within cell-laden GelMA hydrogels by the extrusion bioprinting method, using agarose as the sacrificial template [88,160]. The agarose fibers were then removed, resulting in perfusable vessels within GelMA. The ID of these vessels ranged from 800 μm to 2 mm, while the wall thickness ranged from 50 to 100 μm. These perfusable vessels allowed increased mass transport in thick hydrogels and supported the formation of endothelial monolayers. A similar bioprinting approach was employed for the generation of hollow vasculature within GelMA hydrogels using Pluronic F127 [161]. HUVECs were perfused into the hollow channel to form EC monolayers.

As an alternative, a 3D vascularized network was created in which an open and interconnected sacrificial carbohydrate glass lattice was printed at 110 ºC using a carbohydrate glass formulation (a mixture of 25 g of glucose, 53 g of sucrose, and 10 g of dextran in 50 mL of reverse-osmosis water), through a steel nozzle (ID = 1.2 or 0.84 mm) [162]. The printed carbohydrate glass lattice was then immersed in a poly(d-lactide-co-glycolide) solution after vitrification at 50 ºC for 5 min, which prevented osmotic damage of the encapsulated cells due to carbohydrate dissolution. Various cell-laden hydrogels, including human embryonic kidney (HEK) 293 cell-laden PEG hydrogel (comprising PEG-diacrylate (5%, 10%, or 20% w/w) and 1 mmol·L–1 acrylate– PEG–arginine-glycine-aspartic acid-serine (RGDS) and Irgacure 2959 (0.05% w/v), 10T1/2 cell-laden fibrin gel (10 mg·mL–1 , comprising fibrinogen and thrombin), 10T1/2 cell-laden alginate gel (2% w/v), primary rat hepatocyte- and stromal fibroblast-loaded agarose gels (2% w/v), and 10T1/2 cell-laden Matrigel, were used to encapsulate the 3D-printed glass lattices. Following ECM crosslinking, the sacrificial carbohydrate glass was removed by immersing the whole structure in medium, leaving behind open channels within the ECM hydrogel. These vascular channels were lined with HUVECs, followed by perfusion with blood under highpressure pulsatile flows. It was revealed that these vascular channels were able to support the metabolic function of primary rat hepatocytes in the 3D bioprinted tissue constructs. Following the same approach, sacrificial carbohydrate glass filaments were printed using 100 g of isomalt and 10 g of dextran in 60 mL of reverse-osmosis water, through a steel nozzle (ID = 0.84 mm), which was encapsulated within a fibrin gel (10 mg·mL–1 ) [163]. The sacrificial carbohydrate filaments were removed by washing with phosphate-buffered saline (PBS), followed by the injection of HUVECS into the hollow channels. These 3D-printed vascular patches were implanted in a mouse model of hind limb ischemia, and demonstrated enhanced angiogenesis and the integration of engineered vessels with the host vasculature, resulting in perfusion of distal ischemic tissues. Similarly, when applied in a myocardial infarction mouse model, these bioprinted vascular patches were demonstrated to partially rescue cardiac function, with an ejection fraction and a cardiac output similar to those in the healthy controls.

In another study, two fluidic vascular channels were created within a 3D collagen I matrix (3 mg·mL–1 ) using gelatin (10% w/v) as a sacrificial material; the collagen precursor was first bioprinted on a flow chamber, followed by the bioprinting of two gelatin fibers [164]. After the deposition of HUVECs and normal human lung fibroblasts encapsulated in a fibrin gel in between two gelatin fibers, several layers of collagen were bioprinted on top to cover the entire structure. The sacrificial gelatin was then removed and HUVECs were injected into the channels, which were later allowed to flow with medium. The study demonstrated the formation of a microvasculature network by embedding HUVECs and fibroblasts within the fibrin gel in between the two vascular channels.

Recently, a drop-on-demand bioprinting method for producing blood vessel constructs was demonstrated using a bioprinter with three printheads[10,11]. Gelatin (5% w/v) or HUVEC-loaded gelatin (3% w/v) was bioprinted as a sacrificial core using a printhead of 150 μm in diameter. Two other printheads of 0.3 mm in diameter were used to fabricate the muscle layer; one printhead dispensed droplets of crosslinker solution, containing thrombin, transglutaminase, and CaCl2, while the other printhead dispensed the droplets of fibrinogen (2.5% w/v)-human umbilical artery SMC suspension. Finally, after the completion of the bioprinting process, the outer layer of the blood vessel was formed by casting a fibrinogen (1.25% w/v)– collagen (0.18% v/v) blend encapsulating normal human dermal fibroblasts around the muscle layer. Thus, the resulting perfusable vascular channel, with a diameter of 1 mm and a wall thickness of up to 425 μm, consisted of an endothelium, an SMC layer, and a surrounding matrix of fibroblasts.

5. Future directions

Advances in bioprinting technology have led to the bioengineering of perfusable blood vessel structures and vascularized tissue constructs with different shapes, sizes, and functions (Table 3) [110,116,148–151,153]. However, most bioprinted vascular tissues still lack the key structural components, mechanical properties, and physiological features of native blood vessels. Each bioprinting modality has its own advantages and disadvantages. An ideal modality should have maximized capacities such as resolution and bioprinting speed, and should be applicable to a wide range of bioinks with optimal vascularization potential in a vessel typespecific manner. Most previous studies have used HUVECs as a model cell type. Nevertheless, in order to achieve clinically relevant vascular tissue formation, bioprinted vascular constructs should be able to undergo remodeling and functional transition with relevant cell sources. Therefore, more efforts are required to explore improved bioprinting processes, as well as the development of advanced bioinks for fabricating functional smalldiameter vascular grafts and vascularized tissue grafts with appropriate mechanical behaviors and favorable biological performances. Some of these biological performances may be facilitated further by incorporating dynamic culture conditions, such as perfusion bioreactors, to enhance maturation. Finally, a continued understanding of vascular biology, including the interactions between different layers of blood vessels such as ECs, SMCs, surrounding supporting cells, and the ECM, will provide better insights toward improved engineering designs.

Table 3 Representative parameters and properties of bioprinted small-diameter blood vessels.

HSFs: human skin fibroblasts.

Acknowledgements

This work was supported by funding from the US National Institutes of Health (R00CA201603, R21EB025270, R21EB026175, and R01EB028143) and the Brigham Research Institute.

Compliance with ethics guidelines

Xia Cao, Sushila Maharjan, Ramla Ashfaq, Jane Shin, and Yu Shrike Zhang declare that they have no conflict of interest or financial conflicts to disclose.

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