Recent advances in three‐dimensional bioprinting of stem cells

In spite of being a new field, three‐dimensional (3D) bioprinting has undergone rapid growth in the recent years. Bioprinting methods offer a unique opportunity for stem cell distribution, positioning, and differentiation at the microscale to make the differentiated architecture of any tissue while maintaining precision and control over the cellular microenvironment. Bioprinting introduces a wide array of approaches to modify stem cell fate. This review discusses these methodologies of 3D bioprinting stem cells. Fabricating a fully operational tissue or organ construct with a long life will be the most significant challenge of 3D bioprinting. Once this is achieved, a whole human organ can be fabricated for the defect place at the site of surgery.     

Effects of 3-dimensional Bioprinting Alginate/Gelatin Hydrogel Scaffold Extract on Proliferation and Differentiation of Human Dental Pulp Stem Cells

IntroductionAlginate/gelatin hydrogel (Alg-Gel) scaffold has been applied in tissue engineering, but the research on its application in dental tissues regeneration is still lacking. We investigated the effect of this scaffold on human dental pulp stem cells (hDPSCs).MethodshDPSCs were cultured in both Alg-Gel and 3D-printed Alg-Gel scaffolds. Cell growth and adhesion were compared using fluorescein isothiocyanate–phalloidin staining and scanning electron microscopic micrographs. Changes in the proliferation in hDPSCs cultured in the complete culture medium containing aqueous extracts of the Alg-Gel or 3D-printed Alg-Gel scaffolds were examined using Cell Counting Kit-8 assay and flow cytometry analysis. Cells were cultured in the mineralization medium containing aqueous extracts of the Alg-Gel or 3D-printed Alg-Gel scaffolds for 7 or 14 days, and the differentiation of cells was shown by alizarin red S staining and alkaline phosphatase staining. The messenger RNA and protein expression of mineralization-related genes were detected with real-time polymerase chain reaction and Western blotting. Elemental analysis was used to test the material extract composition.ResultsMore cells were grown and adhered to the 3D-printed Alg-Gel scaffolds than the Alg-Gel scaffolds. The aqueous extracts of 3D-printed scaffolds can promote cell proliferation, and compared with Alg-Gel scaffolds, the extracts of 3D-printed scaffolds were more effective. Compared with the negative control group, 3D-printed Alg-Gel scaffold and Alg-Gel scaffold aqueous extracts promoted osteogenic/odontoblastic differentiation of hDPSCs with the enhanced formation of bone-like nodules and the alkaline phosphatase staining. The expression of mineralization-related genes was also up-regulated. 3D-printed scaffold aqueous extract contained more calcium and phosphorus ions than the Alg-Gel scaffold.ConclusionsThese findings suggest that compared with the Alg-Gel scaffold, 3D-printed Alg-Gel is more suitable for the growth of hDPSCs, and the scaffold extracts can better promote cell proliferation and differentiation.     

Three-dimensional bioprinting of thick vascularized tissues

The advancement of tissue and, ultimately, organ engineering requires the ability to pattern human tissues composed of cells, extracellular matrix, and vasculature with controlled microenvironments that can be sustained over prolonged time periods. To date, bioprinting methods have yielded thin tissues that only survive for short durations. To improve their physiological relevance, we report a method for bioprinting 3D cell-laden, vascularized tissues that exceed 1 cm in thickness and can be perfused on chip for long time periods (>6 wk). Specifically, we integrate parenchyma, stroma, and endothelium into a single thick tissue by coprinting multiple inks composed of human mesenchymal stem cells (hMSCs) and human neonatal dermal fibroblasts (hNDFs) within a customized extracellular matrix alongside embedded vasculature, which is subsequently lined with human umbilical vein endothelial cells (HUVECs). These thick vascularized tissues are actively perfused with growth factors to differentiate hMSCs toward an osteogenic lineage in situ. This longitudinal study of emergent biological phenomena in complex microenvironments represents a foundational step in human tissue generation.The ability to manufacture human tissues that replicate the essential spatial (1), mechanochemical (2, 3), and temporal aspects of biological tissues (4) would enable myriad applications, including 3D cell culture (5), drug screening (6, 7), disease modeling (8), and tissue repair and regeneration (9, 10). Three-dimensional bioprinting is an emerging approach for creating complex tissue architectures (10, 11), including those with embedded vasculature (12–15), that may address the unmet needs of tissue manufacturing. Recently, Miller et al. (15) reported an elegant method for creating vascularized tissues, in which a sacrificial carbohydrate glass is printed at elevated temperature (>100 °C), protectively coated, and then removed, before introducing a homogeneous cell-laden matrix. Kolesky et al. (14) developed an alternate approach, in which multiple cell-laden, fugitive (vasculature), and extracellular matrix (ECM) inks are coprinted under ambient conditions. However, in both cases, the inability to directly perfuse these vascularized tissues limited their thickness (1–2 mm) and culture times (<14 d). Here, we report a route for creating thick vascularized tissues (≥1 cm) within 3D perfusion chips that provides unprecedented control over tissue composition, architecture, and microenvironment over several weeks (>6 wk). This longitudinal study of emergent biological phenomena in complex microenvironments represents a foundational step in human tissue generation.Central to the fabrication of thick vascularized tissues is the design of biological, fugitive, and elastomeric inks for multimaterial 3D bioprinting. To satisfy the concomitant requirements of processability, heterogeneous integration, biocompatibility, and long-term stability, we first developed printable cell-laden inks and castable ECM based on a gelatin and fibrinogen blend (16). Specifically, these materials form a gelatin–fibrin matrix cross-linked by a dual-enzymatic, thrombin and transglutaminase (TG), strategy (Fig. 1 and SI Appendix, Fig. S1). The cell-laden inks must facilitate printing of self-supporting filamentary features under ambient conditions as well as subsequent infilling of the printed tissue architectures by casting without dissolving or distorting the patterned construct (Fig. 1A). The thermally reversible gelation of the gelatin–fibrinogen network enables its use in both printing and casting, where gel and fluid states are required, respectively (SI Appendix, Fig. S2). Thrombin is used to rapidly polymerize fibrinogen (17), whereas TG is a slow-acting Ca²⁺-dependent enzymatic cross-linker that imparts the mechanical and thermal stability (18) needed for long-term perfusion. Notably, the cell-laden ink does not contain either enzyme to prevent polymerization during printing. However, the castable matrix contains both thrombin and TG, which diffuse into adjacent printed filaments, forming a continuous, interpenetrating polymer network, in which the native fibrillar structure of fibrin is preserved (SI Appendix, Fig. S3). Importantly, our approach allows arbitrarily thick tissues to be fabricated, because the matrix does not require UV curing (19), which has a low penetration depth in tissue (20) and can be readily expanded to other biomaterials, including fibrin and hyaluronic acid (SI Appendix, Fig. S4).Open in a separate windowFig. 1.Three-dimensional vascularized tissue fabrication. (A) Schematic illustration of the tissue manufacturing process. (i) Fugitive (vascular) ink, which contains pluronic and thrombin, and cell-laden inks, which contain gelatin, fibrinogen, and cells, are printed within a 3D perfusion chip. (ii) ECM material, which contains gelatin, fibrinogen, cells, thrombin, and TG, is then cast over the printed inks. After casting, thrombin induces fibrinogen cleavage and rapid polymerization into fibrin in both the cast matrix, and through diffusion, in the printed cell ink. Similarly, TG diffuses from the molten casting matrix and slowly cross-links the gelatin and fibrin. (iii) Upon cooling, the fugitive ink liquefies and is evacuated, leaving behind a pervasive vascular network, which is (iv) endothelialized and perfused via an external pump. (B) HUVECs growing on top of the matrix in 2D, (C) HNDFs growing inside the matrix in 3D, and (D) hMSCs growing on top of the matrix in 2D. (Scale bar: 50 µm.) (E and F) Images of printed hMSC-laden ink prepared using gelatin preprocessed at 95 °C before ink formation (E) as printed and (F) after 3 d in the 3D printed filament where actin (green) and nuclei (blue) are stained. (G) Gelatin preprocessing temperature affects the plateau modulus and cell viability after printing. Higher temperatures lead to lower modulus and higher HNDF viability postprinting. (H) Photographs of interpenetrated sacrificial (red) and cell inks (green) as printed on chip. (Scale bar: 2 mm.) (I) Top-down bright-field image of sacrificial and cell inks. (Scale bar: 50 µm.). (J–L) Photograph of a printed tissue construct housed within a perfusion chamber (J) and corresponding cross-sections (K and L). (Scale bars: 5 mm.)The gelatin–fibrin matrix supports multiple cell types of interest to both 2D and 3D culture conditions, including human umbilical vein endothelial cells (HUVECs), human neonatal dermal fibroblasts (HNDFs), and human bone marrow-derived mesenchymal stem cells (hMSCs) (Fig. 1 B–D and SI Appendix, Fig. S5). We find that endothelial cells express vascular endothelial-cadherin (VE-Cad) (Fig. 1B), and HNDFs (Fig. 1C) and hMSCs (Fig. 1D) proliferate and spread on this matrix surface and in bulk. Moreover, the printed cell viability can be as high as 95%, depending on how gelatin is processed before ink formulation. At higher processing temperatures, the average molecular weight of gelatin is reduced from 69 kDa at 70 °C to 32 kDa at 95 °C processing, resulting in softer gels with lower viscosity, shear yield stress, and shear elastic modulus. These cell-laden inks can be printed with ease and accommodate cell densities ranging from 0.1 million per mL to 10 million cells per mL (Fig. 1E and SI Appendix, Fig. S6). Upon printing, hMSCs within this soft gelatin–fibrinogen matrix continue to spread, proliferate, and contract into dense, cellular architectures that align along the printing direction (Fig. 1F), likely arising due to cellular confinement (21) and contraction via the Poisson effect (22).To construct thick, vascularized tissues within 3D perfusion chips, we coprinted cell-laden, fugitive, and silicone inks (Fig. 1 H and I). First, the silicone ink is printed on a glass substrate and cured to create customized perfusion chips (Movie S1 and SI Appendix, Fig. S1). Next, the cell-laden and fugitive inks are printed on chip, and then encapsulated with the castable ECM (Fig. 1 J–L and Movie S2). The fugitive ink, which defines the embedded vascular network, is composed of a triblock copolymer [i.e., polyethylene oxide (PEO)–polypropylene oxide (PPO)–PEO]. This ink can be removed from the fabricated tissue upon cooling to roughly 4 °C, where it undergoes a gel-to-fluid transition (14, 23). This process yields a pervasive network of interconnected channels, which are then lined with HUVECs. The resulting vascularized tissues are perfused via their embedded vasculature on chip over long time periods using an external pump (Movie S3) that generates smooth flow over a wide range of flow rates (24).To demonstrate the formation of stable vasculature, we printed a simple tissue construct composed of two parallel channels embedded within a fibroblast cell-laden matrix (Fig. 2). The channels are lined with HUVECs, perfused with 1:1 ratio of endothelial growth media (EGM-2 Bullet kit) and HNDF growth media [DMEM plus 10% (vol/vol) FBS], and subsequently form a confluent monolayer that lines each blood vessel (Fig. 2A). The medium is preincubated for 5 h in the incubator at 37 °C and 5% CO₂ and replaced every other day. Importantly, after 6 wk of active perfusion, these endothelial cells maintain endothelial phenotype and remain confluent, characterized by expression of CD31, von Willebrand factor (vWF), and VE-Cad (Fig. 2 B and C). The cross-sectional view of a representative vessel reveals lumen formation (Fig. 2D and Movie S4). Confirming the barrier function of the endothelium, we measured a fivefold reduction in the diffusional permeability compared with unlined (bare) channels (Fig. 2E and SI Appendix, Fig. S7). Stromal HNDFs residing within the surrounding matrix exhibit cell spreading and proliferative phenotypes localized to regions within ∼1 mm of the vasculature (Fig. 2F and SI Appendix, Fig. S8); cells further away from these regions become quiescent likely due to an insufficient nutrient supply. As cell density increases, their viability rapidly decreases at distances beyond 1 mm from the embedded blood vessels (e.g., only 5% of the cells remain viable at 7 mm). Clearly, the perfusable vasculature is critical to support living tissues thicker than 1 mm over long time periods.Open in a separate windowFig. 2.Three-dimensional vascularized tissues remain stable during long-term perfusion. (A) Schematic depicting a single HUVEC-lined vascular channel supporting a fibroblast cell-laden matrix and housed within a 3D perfusion chip. (B and C) Confocal microscopy image of the vascular network after 42 d, CD-31 (red), vWF (blue), and VE-Cadherin (magenta). (Scale bars: 100 µm.) (D) Long-term perfusion of HUVEC-lined (red) vascular network supporting HNDF-laden (green) matrix shown by top-down (Left) and cross-sectional confocal microscopy at 45 d (Right). (Scale bar: 100 µm.) (E) Quantification of barrier properties imparted by endothelial lining of channels, demonstrated by reduced diffusional permeability of FITC-dextran. (F) GFP-HNDF distribution within the 3D matrix shown by fluorescent intensity as a function of distance from vasculature.To explore emergent phenomena in complex microenvironments, we created a heterogeneous tissue architecture (>1 cm thick and 10 cm³ in volume) by printing a hMSC-laden ink into a 3D lattice geometry along with intervening in- and out-of-plane (vertical) features composed of fugitive ink, which ultimately transform into a branched vascular network lined with HUVECs. After printing, the remaining interstitial space is infilled with an HNDF-laden ECM (Fig. 3A) to form a connective tissue that both supports and binds to the printed stem cell-laden and vascular features. In this example, fibroblasts serve as model cells that surround the heterogeneously patterned stem cells and vascular network. These model cells could be replaced with either support cells (e.g., immune cells or pericytes) or tissue-specific cells (e.g., hepatocytes, neurons, or islets) in future embodiments. The embedded vascular network is designed with a single inlet and outlet that provides an interface between the printed tissue and the perfusion chip. This network is symmetrically branched to ensure uniform perfusion throughout the tissue, including deep within its core. In addition to providing transport of nutrients, oxygen, and waste materials, the perfused vasculature is used to deliver specific differentiation factors to the tissue in a more uniform manner than bulk delivery methods, in which cells at the core of the tissue are starved of factors (25). This versatile platform (Fig. 3A) is used to precisely control growth and differentiation of the printed hMSCs. Moreover, both the printed cellular architecture and embedded vascular network are visible macroscopically with this thick tissue (Fig. 3B).Open in a separate windowFig. 3.Osteogenic differentiation of thick vascularized tissue. (A) Schematic depicting the geometry of the printed heterogeneous tissue within the customized perfusion chip, wherein the branched vascular architecture pervades hMSCs that are printed into a 3D lattice architecture, and HNDFs are cast within an ECM that fills the interstitial space. (B) Photographs of a printed tissue construct within and removed from the customized perfusion chip. (C) Comparative cross-sections of avascular tissue (Left) and vascularized tissue (Right) after 30 d of osteogenic media perfusion with alizarin red stain showing location of calcium phosphate. (Scale bar: 5 mm.) (D) Confocal microscopy image through a cross-section of 1-cm-thick vascularized osteogenic tissue construct after 30 d of active perfusion and in situ differentiation. (Scale bar: 1.5 mm.) (E) Osteocalcin intensity across the thick tissue sample inside the red lines shown in C. (F) High-resolution image showing osteocalcin (purple) localized within hMSCs, and they appear to take on symmetric osteoblast-like morphologies. (Scale bar: 100 µm.) After 30 d (G and H), thick tissue constructs are stained for collagen-I (yellow), which appears to be localized near hMSCs. (Scale bars: 200 µm.) (I) Alizarin red is used to stain calcium phosphate deposition, and fast blue is used to stain AP, indicating tissue maturation and differentiation over time. (Scale bar: 200 µm.)To develop a dense osteogenic tissue, we transvascularly delivered growth media to the tissue during an initial proliferation phase (6 d) followed by an osteogenic differentiation mixture that is perfused for several weeks. Our optimized mixture is composed of BMP-2, ascorbic acid, and glycerophosphate, to promote mineral deposition and alkaline phosphatase (AP) expression (SI Appendix, Fig. S9). To assess tissue maturation, changes in cell function and matrix composition are observed over time. In good agreement with prior studies (21), we find that AP expression in hMSCs occurs within 3 d, whereas mineral deposition does not become noticeable until 14 d, which coincides with visible collagen-1 deposition by hMSCs (SI Appendix, Fig. S9) (21). Fig. 3C shows an avascular tissue produced with comparable hMSC density, in which positive alizarin stains are only observed within a few hundred microns of the tissue surface. By contrast, the thick vascularized tissue stains positive in hMSC regions deep within its core after 30 d of osteogenic differentiation by perfusion. We characterized the mineral deposits, which consist of particulates ∼20–200 nm in size, using SEM/energy-dispersive X-ray spectroscopy (EDS) analysis. Calcium and phosphorous peaks are only observed for vascularized tissues, not the avascular control (SI Appendix, Fig. S9 E and F). The phenotype of hMSCs varies across the printed filamentary features: cells are close-packed, compacted, and exhibit a high degree of mineralization within the filament core, whereas those in the periphery are more elongated and exhibit less mineralization. We observe that subpopulations of HNDFs and hMSCs migrate from their initial patterned geometry toward the vascular channels and wrap circumferentially around each channel (Fig. 3D). After 30 d, the printed hMSCs express osteocalcin within the tissue, and osteocalcin expression is proportional to distance from the nearest vessel (Fig. 3E). Furthermore, we find that collagen deposition is localized within printed filaments and around the circumference of the vasculature (Fig. 3 F–H and SI Appendix, Fig. S9).In summary, thick, vascularized human tissues with programmable cellular heterogeneity that are capable of long-term (>6-wk) perfusion on chip have been fabricated by multimaterial 3D bioprinting. The ability to recapitulate physiologically relevant, 3D tissue microenvironments enables the exploration of emergent biological phenomena, as demonstrated by observations of in situ development of hMSCs within tissues containing a pervasive, perfusable, endothelialized vascular network. Our 3D tissue manufacturing platform opens new avenues for fabricating and investigating human tissues for both ex vivo and in vivo applications.     

High‐throughput fabrication of vascularized adipose microtissues for 3D bioprinting

For patients with soft tissue defects, repair with autologous in vitro engineered adipose tissue could be a promising alternative to current surgical therapies. A volume‐persistent engineered adipose tissue construct under in vivo conditions can only be achieved by early vascularization after transplantation. The combination of 3D bioprinting technology with self‐assembling microvascularized units as building blocks can potentially answer the need for a microvascular network. In the present study, co‐culture spheroids combining adipose‐derived stem cells (ASC) and human umbilical vein endothelial cells (HUVEC) were created with an ideal geometry for bioprinting. When applying the favourable seeding technique and condition, compact viable spheroids were obtained, demonstrating high adipogenic differentiation and capillary‐like network formation after 7 and 14 days of culture, as shown by live/dead analysis, immunohistochemistry and RT‐qPCR. Moreover, we were able to successfully 3D bioprint the encapsulated spheroids, resulting in compact viable spheroids presenting capillary‐like structures, lipid droplets and spheroid outgrowth after 14 days of culture. This is the first study that generates viable high‐throughput (pre‐)vascularized adipose microtissues as building blocks for bioprinting applications using a novel ASC/HUVEC co‐culture spheroid model, which enables both adipogenic differentiation while simultaneously supporting the formation of prevascular‐like structures within engineered tissues in vitro.     

Meniscus ECM‐functionalised hydrogels containing infrapatellar fat pad‐derived stem cells for bioprinting of regionally defined meniscal tissue

Injuries to the meniscus of the knee commonly lead to osteoarthritis. Current therapies for meniscus regeneration, including meniscectomies and scaffold implantation, fail to achieve complete functional regeneration of the tissue. This has led to increased interest in cell and gene therapies and tissue engineering approaches to meniscus regeneration. The implantation of a biomimetic implant, incorporating cells, growth factors, and extracellular matrix (ECM)‐derived proteins, represents a promising approach to functional meniscus regeneration. The objective of this study was to develop a range of ECM‐functionalised bioinks suitable for 3D bioprinting of meniscal tissue. To this end, alginate hydrogels were functionalised with ECM derived from the inner and outer regions of the meniscus and loaded with infrapatellar fat pad‐derived stem cells. In the absence of exogenously supplied growth factors, inner meniscus ECM promoted chondrogenesis of fat pad‐derived stem cells, whereas outer meniscus ECM promoted a more elongated cell morphology and the development of a more fibroblastic phenotype. With exogenous growth factors supplementation, a more fibrogenic phenotype was observed in outer ECM‐functionalised hydrogels supplemented with connective tissue growth factor, whereas inner ECM‐functionalised hydrogels supplemented with TGFβ3 supported the highest levels of Sox‐9 and type II collagen gene expression and sulfated glycosaminoglycans (sGAG) deposition. The final phase of the study demonstrated the printability of these ECM‐functionalised hydrogels, demonstrating that their codeposition with polycaprolactone microfibres dramatically improved the mechanical properties of the 3D bioprinted constructs with no noticeable loss in cell viability. These bioprinted constructs represent an exciting new approach to tissue engineering of functional meniscal grafts.     

基于同轴细胞打印双网络生物墨水优化及类血管支架的打印

文题释义：双网络生物墨水：生物墨水是指可以用于生物3D打印机的材料,具有类似细胞外基质的理化性质,可用于制造与人体器官相似的组织。双网络生物墨水内部具有两种交联网络,能使体外构建的组织具有良好的机械性能,适用于不同的应用场景。 同轴细胞打印：生物3D打印也叫细胞打印,是指操控细胞生物墨水体外构建活性组织的过程。同轴细胞打印是生物3D打印的延伸和发展,通过结合多层同轴针头可以直接快速制备含有内部连通网络的组织工程支架。 背景：细胞体外培养情况下无法在远离营养物质200 μm以上的区域存活,血管网络构建对组织工程领域厚组织和器官再生至关重要,同轴细胞打印为体外构建类血管通道提供了一种新的方式。 目的：优化生物墨水的同轴细胞打印性能,制备具有类血管结构的组织工程支架。 方法：通过间歇式巴氏灭菌制备无菌海藻酸钠溶液,冷冻保存;以脱胶蚕丝为原料制备无菌丝素蛋白冻干粉,密封保存;将丝素蛋白冻干粉加入解冻的海藻酸钠溶液中,再加入人脐静脉内皮细胞,作为生物墨水;将生物3D打印机的外轴连接生物墨水,内轴连接交联剂,同轴打印类血管支架材料,进行光学相干层析成像扫描、扫描电镜观察;拉伸测试海藻酸钠与丝素蛋白/海藻酸钠同轴打印环形试件(不含细胞)的弹性模量。采用冷冻保存7 d的海藻酸钠溶液与人脐静脉内皮细胞制作同轴打印支架,冷冻保存7 d的海藻酸钠溶液、人脐静脉内皮细胞与密封保存6个月的丝素蛋白冻干粉制作同轴打印支架,培养24 h后死活染色观察细胞存活率。设计打印串联与并联结构的类血管支架,培养1,3,7,10,14 d后检测细胞增殖情况。 结果与结论：①光学相干层析成像扫描显示,该混合生物墨水最高打印高度为9层,整体厚度约为4.4 mm;扫描电镜显示,类血管支架的中空纤维丝外壁呈无规则条状卷曲,存在微米级内部连通孔隙结构,中空纤维丝内壁具有更致密的孔隙结构;②丝素蛋白/海藻酸钠同轴打印环形试件的弹性模量大于单纯海藻酸钠同轴打印环形试件(P < 0.05);③采用保存7 d海藻酸钠溶液制作的支架细胞存活率为(86.7±3.4)%,加入丝素蛋白冻干粉支架的细胞存活率为(98.1±1.2)%,说明冷冻保存7 d的海藻酸钠溶液未染菌,丝素蛋白的保质期可达6个月;④并联结构类血管支架培养7,10,14 d的细胞增殖活性高于串联结构的类血管支架(P < 0.05);⑤结果表明,实验制备的类血管支架材料具有良好的生物相容性与机械性能。 ORCID: 0000-0002-5556-6672(张一帆) 中国组织工程研究杂志出版内容重点：生物材料;骨生物材料; 口腔生物材料; 纳米材料; 缓释材料; 材料相容性;组织工程     

3D bioprinting in orthopedics translational research

Abstract

The repair of critical-size bone defect remains a challenge for orthopedic surgeons. With the advent of an aging society and their accompanying chronic diseases, it is becoming more difficult to treat bone defects, especially large segmental bone defects that are caused by trauma, tumors, infections, and congenital malformations. New materials and technologies need to be developed to address these conditions. 3D bioprinting is a novel technology that bridges the biomaterial and living cells and is an important method in tissue engineering projects. 3D bioprinting has the advantages of replacing or repairing damaged tissue and organs. The progress in material science and 3D printing devices make 3D bioprinting a technology which can be used to create various scaffolds with a large range of advanced material and cell types. However, in regard to the widespread use of bioprinting, biosafety, immunogenicity and rising costs are rising to be concerned. This article reviews the developments and applications of 3D bioprinting and highlights newly applied techniques and materials and the recent achievements in the orthopedic field. This paper also briefly reviews the difference between the methods of 3D bioprinting. The challenges are also elaborated with the aim to research materials, manufacture scaffolds, promote vascularization and maintain cell viability.