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      Recent advances in 3D printing of biomaterials

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      Journal of Biological Engineering
      Springer Science and Business Media LLC

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          Abstract

          3D Printing promises to produce complex biomedical devices according to computer design using patient-specific anatomical data. Since its initial use as pre-surgical visualization models and tooling molds, 3D Printing has slowly evolved to create one-of-a-kind devices, implants, scaffolds for tissue engineering, diagnostic platforms, and drug delivery systems. Fueled by the recent explosion in public interest and access to affordable printers, there is renewed interest to combine stem cells with custom 3D scaffolds for personalized regenerative medicine. Before 3D Printing can be used routinely for the regeneration of complex tissues (e.g. bone, cartilage, muscles, vessels, nerves in the craniomaxillofacial complex), and complex organs with intricate 3D microarchitecture (e.g. liver, lymphoid organs), several technological limitations must be addressed. In this review, the major materials and technology advances within the last five years for each of the common 3D Printing technologies (Three Dimensional Printing, Fused Deposition Modeling, Selective Laser Sintering, Stereolithography, and 3D Plotting/Direct-Write/Bioprinting) are described. Examples are highlighted to illustrate progress of each technology in tissue engineering, and key limitations are identified to motivate future research and advance this fascinating field of advanced manufacturing.

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          Printing three-dimensional tissue analogues with decellularized extracellular matrix bioink

          The ability to print tissue analogue structures through delivering living cells with appropriate material in a defined and organized manner, at the right location, in sufficient numbers, and within the right environment is critical for several emerging technologies. These technologies include, tissue-engineering scaffolds1 2, cell-based sensors3, drug/toxicity screening4 and tissue or tumour models5. The concept of tissue or organ printing, often described as bioprinting6, is essentially an extension of the idea that uses additive manufacturing methods to build complex scaffold structures via a layer-by-layer process7 8 9 10. A crucial aspect of bioprinting is that the printing process must be cytocompatible, as it requires the dispensing of cell-containing media. This restriction reduces the choice of materials because of the necessity to operate in an aqueous or aqueous gel environment11 12. In extrusion-based printing, hydrogels that are solidified through either thermal processes or post-print cross-linking are being used for printing of cells to produce diverse tissues ranging from liver to bone using materials such as gelatin13, gelatin/chitosan14, gelatin/alginate15, gelatin/fibrinogen16, Lutrol F127/alginate17 and alginate18. However, there are some concerns over the outcomes from these studies, such as the use of harsh cross-linking agents, like glutaraldehyde14. Elsewhere, osteogenic differentiation was not prominent on alginate gel and no differentiation was observed on Lutrol F127 (ref. 17). In addition, when alginate gel was used for printing of a cell-printed structure, only a minor fraction of cells in the construct could differentiate towards osteogenic lineage17. Normally, cells remain located specifically in their original deposited position during the whole culture period, as they are unable to adhere or degrade the surrounding alginate gel matrix19. This limited interaction between the cells within the gel can be explained by the noninteractive nature of alginate. Thus, although there were some successful reports about bioprinting of cell-printed structure, minimal cells–material interactions and inferior tissue formation are the foremost concerns. Actually, these materials cannot represent the complexity of natural extracellular matrices (ECMs) and thus are inadequate to recreate a microenvironment with cell–cell connections and three-dimensional (3D) cellular organization that are typical of living tissues. Consequently, the cells in those hydrogels cannot exhibit intrinsic morphologies and functions of living tissues in vivo. It is thus ideal if cells are provided the natural microenvironment similar to their parent tissue. Decellularized extracellular matrix (dECM) is the best choice for doing so, as no natural or man-made material can recapitulate all the features of natural extracellular matrix (ECM)20. Moreover, ECM of each tissue is unique in terms of composition and topology, which is generated through dynamic and reciprocal interactions between the resident cells and microenvironment21. Recent studies of cells and ECM isolated from tissues and organs highlight the necessity of tissue specificity for preserving selected cell functions and phenotype20 22 23 24 25. The dECM materials are harvested and typically processed as two-dimensional (2D) scaffolds from various tissues, including skin26, small intestinal submucosa27, where at the initial stages the infiltrating or seeded cell populations depend on diffusion of oxygen and nutrient for their survival until a supporting vascular network develops. However, printing tissue analogue structures requires a fabrication approach to devise a highly open porous 3D structure to allow the flow of nutrients11 28. Organ transplantation in an orthotopic location is considered as a definitive treatment for end-stage organ failure23 24 29. However, the demand for transplantable organs surpasses the number of available donor organs, compelling the manufacture of tissue or organs to address this unmet demand. Here we develop a bioprinting method for printing of cell-laden structure with novel dECM bioink capable of providing an optimized microenvironment conducive to the growth of 3D structured tissue, where the intrinsic cellular morphologies and functions can be reconstituted. We show the versatility and the functionality of our bioprinting process using various types of dECMs bioinks, including adipose, cartilage and heart tissues. As applications of these functional cell-printed constructs, we observe higher-order assembly of these cellular constructs with organized spatial patterns and tissue-specific gene expression. A key advantage of this methodology is the application of tissue-specific ECM, providing crucial cues for cells engraftment, survival and long-term function. Results Preparation of dECM bioink The method for printing of cell-laden construct using dECM bioink consists of a few steps (Fig. 1). Decellularization of the ECM material is the initial step and practically, the goal is to maximize the removal of cellular material, while minimizing ECM loss and damage30. We successfully decellularized the adipose (adECM), cartilage (cdECM) and heart (hdECM) tissue after harvesting with a combination of physical, chemical and enzymatic process using slightly modified methods described elsewhere31 32 33. Furthermore, 0.1% peracetic acid solution in combination with 4% ethanol was used in the last step of the decellularization method, which is frequently considered as a disinfection step34. Efficiency of decellularization method was evaluated through DNA quantification assay and was found to be optimal for removing the cellular contents with ~98% reduction and only 39±15, 11±1 and 6.7±1.2 ng of DNA per mg of tissue remaining in the dECM of fat, cartilage and heart tissue, respectively (Fig. 2). Hematoxylin and eosin (H&E) staining confirmed the absence of cells and cell debris in the matrix after decellularization (Fig. 2b,c) in all samples of dECMs, including those from the innermost regions of the processed tissues. Although the term decellularization has not been defined by quantitative metrics, 95%) when examined after 24 h (Fig. 5e). Cell-printed structure with only hdECM also revealed similar result of high viability of myoblasts after 24 h (Supplementary Fig. 4). Actually, dECM gels possibly protect the cells through cushioning them during extrusion and allow cells to acquire the desired morphology during their residence in the gel. We also could observe the cell- to -cell connections in 3D within 24 h mimicking the in vivo conditions (Fig. 5e), which is very important for their survival and functions. Moreover, the dECM gels did not produce any deleterious effect on the cells or hindered their migration as the high cell viability (>90%) was maintained when the sample was examined on day 7 and 14 with active cell proliferation (Fig. 5e). Tissue-specific gene expression We investigated cellular morphologies and functions of the cell-laden constructs using stem cells, such as human adipose-derived stem cells (hASCs) and human inferior turbinate-tissue derived mesenchymal stromal cells (hTMSCs), a potential abundant cell source for clinical application from human inferior turbinate tissues generally discarded during turbinate surgery41 42. These cells have been shown to be promising for adipose tissue regeneration25 and cartilage tissue regeneration41, respectively. To assess the differentiation of the printed stem cells, in particular encapsulating in dECM, tissue-specific gene expressions were analysed. Before demonstrating the superiority of each dECM material, cell proliferation test was conducted. This test verified that all the dECMs provide biocompatible microenvironment for cell proliferation and outperformed the other printable materials, such as COL and alginate (Supplementary Figs 5 and 6). Among the various ECM components, COL was selected as a control for comparative analysis of target gene expression, because it is the most abundant component of ECM and has been used for cell printing43. Although, the major COL of cartilage is type II COL, not type I, we chose it as a control because of its increased application in cartilage tissue engineering. Direct differentiation of hTMSCs into chondrogenic lineage in cdECM was verified through qRT–PCR analysis. We normalized the target genes expressions with glyceraldehyde 3-phosphate dehydrogenase (GAPDH), as it shows least variation when cells were cultured on TCP over time (Supplementary Fig. 7). On day 14, cells printed with cdECM revealed greater messenger RNA (mRNA) expressions of SOX9 that is an early chondrogenic transcription factor and type II COL than that of COL construct (Fig. 6a). Moreover, on day 3, SOX9, adhesion-related genes (focal adhesion kinase; protein tyrosine kinase 2) and integrin β1 were significantly greater than that of COL (Supplementary Fig. 8), indicating the chondrogenic potential of cdECM constructs. Rat myoblast cells (L6, ATCC CRL-1458) were used to verify the effectiveness of hdECM for functional maturation of myoblasts. For assessment of the maturation level of myoblast in the hdECM structure, mRNA expression was analysed through qRT–PCR; in particular, the expression level of cardiac-specific genes like fast myosin heavy chain (Myh6) and alpha-sarcomeric actinin (Actn1). The hdECM constructs disclosed significantly higher levels of both the genes than that of the COL constructs during 14 days study period (Fig. 6b). Moreover, the expression of Myh6 and Actn1 were also higher in hdECM than that of COL at day 4 and 7 (Supplementary Fig. 9), which signify the superiority of hdECM over COL gel. Adipogenic differentiation has been shown to enhance under high-density conditions where the cells have a more rounded morphology and the cellular growth ceases44. As such, the high-density cell printing is an important step that will facilitate adipogenic differentiation for supporting adipogenesis in the culture groups. We have shown the possibility of printing cell-laden construct with high density (5 × 106 cells ml−1) using our bioprinting method without any deleterious effect on cell viability. Moreover, a significant increase in the expression of the master adipogenic regulators peroxisome proliferator-activated receptor gamma (PPARγ) and CCAAT/enhancer-binding protein alpha (C/EBPα) together with an increase in early adipogenic marker lipoprotein lipase (LPL) was observed when hASCs were cultured in adECM in comparison with COL and alginate (Fig. 6c and Supplementary Fig. 10), widely used materials for cell printing2 18. The gene expression trends in the constructs showed increase in the expression of the adipogenic markers over time, consistent with a progression in differentiation, demonstrating the ‘adipoconductive’ nature of adECM. Thus, 3D printed constructs fabricated through our bioprinting method using dECM bioink supported differentiation and maturation of encapsulated cells and functionality of the respective tissues. Tissue formation Tissue formation was evaluated through immunohistological study. Supportive role of cdECM to differentiate hTMSCs into the chondrogenic lineage in cell-printed constructs with cdECM was verified through immunofluorescence imaging, which indicated that the cells synthesized type II COL within the construct (Fig. 6d,e and Supplementary Fig. 11) and was in accordance with the mRNA expression results. We also verified the function of cdECM on enhancing cell adhesion by using F-actin staining (Fig. 6e). Most of the cells displayed spread morphology on the TCP (Supplementary Fig. 12a) and COL (Supplementary Fig. 12b); on the other hand, many cells were migrated into the depth of the cdECM (Supplementary Fig. 12c) and the cells were observed to be elongated and created mesh-like junctions in between them. This phenomenon corresponds to the expression of adhesion-related genes. It is also implied that cdECM have abundant residues to bind cells, which might have triggered the chondrogenesis transcription marker like SOX9 (Supplementary Fig. 8). The maturation of myoblasts in cell-printed structure with hdECM was investigated by cardiac myosin heavy chain (β-MHC) staining and was compared with that of COL (Fig. 6f,g and Supplementary Fig. 13), an abundantly available protein in the myocardium. The result showed that cells expressed higher level of β-MHC in the hdECM construct than that of COL and was also in accordance with the qRT–PCR result. In general, myoblasts were formed into the myofibres during the structural maturation. Subsequent to this, the differentiated myoblasts (myocytes) express the slow myosin heavy chain and synchronize with the native tissue. The slow myosin heavy chain is usually expressed in the newly formed cardiomyocytes. This phenomenon is associated with the graft survival; especially, the enhanced graft survival influences the alignment of the differentiated myoblasts in parallel with the host myocardial fibre45. The above finding advocates the potentiality of myoblasts printed hdECM structure for myocardial reconstruction. Upon differentiation of hASCs into the adipogenic lineage, they express PPARγ and COL IV. Cell-printed construct with adECM revealed substantial expression of PPARγ and COL IV (Fig. 6h,i, and Supplementary Fig. 14) with higher level of fluorescence intensity to that of COL construct. These observations highlight the tissue-formation capabilities of our cell-printed constructs. Discussion In the present study, for the first time, tissue printing with particular dECM bioink that encompass either living hASCs or hTMSCs is demonstrated. Printed cell dECM constructs revealed higher levels of cell viability, differential lineage commitment and ECM formation. This study addresses recent advancement in the field of bioprinting for engineering biological tissues, consisting of cells encapsulated in dECM with specific organization and matrix composition. The adapted printing technology of depositing cell-laden dECM hydrogel that we used can fulfil several important requirements of tissue-engineered grafts, including tissue-specific matrix materials, engineered interconnected porosity, the introduction of pertinent cells, and the opportunity to explore organizational aspects in tissue regeneration. Encapsulation of cells within dECM hydrogel prior to the deposition process facilitates the formation of cell-laden, viable printed constructs17. In this way, the cell loading can be altered and different types of cells can be laid down in tissue-mimicking architectures. We have also shown that our printing method does not cause stress-induced apoptosis of the encapsulated cells and high level of cell viability was achieved. Moreover, being a robust dispensing method, our bioprinting technique can produce constructs of anatomically relevant (centimetre scale) sizes for tissue replacement. Pepsin digestion cleaves the telopeptide regions of the COL, which allows solubilization of the dECM in dilute acids43. We could achieve highly concentrated dECM solution with adequate viscosity for printing of cell-laden constructs. When pH values of these dECM solutions were adjusted, they behaved similar to temperature-sensitive pre-gel materials, allowing printing of well-defined 3D constructs that were transformed into physically cross-linked gel at 37 °C. The structures were stable in vitro without the need for chemical cross-linking. We used dECMs processed from tissues, such as adipose, cartilage and heart, for printing of particular tissue analogue. Likewise, this method can also be extended to ECM from other tissues with little modification. At present, tissue-engineering strategies utilizing ECM materials are being successfully used clinically for the regeneration of a range of tissues46 47. The main advantage of utilizing ECM material is that it facilitates constructive remodelling at the implantation site and encourages specific tissue formation rather than forming inferior and less functional scar tissue36. We demonstrated that human living progenitor cells survive the generated shear stress during the printing process of cell-laden dECM hydrogel with a high level of cell survival, which is in line with previous reports17. This can be explained through the shear thinning behaviour of ECM hydrogel, which results in a decrease in shear stress. This phenomenon of shear thinning is a common behaviour for polymeric gel or hydrogel and has been described in detail elsewhere48. We could print a precisely defined but flexible designed structure with a homogenous distribution of cells within the construct. In addition, to improve the mechanical integrity of the printed dECM hydrogel constructs, cell-laden hydrogels were combined with structurally stiffer synthetic polymers, such as PCL for cartilage construct. The versatility of of our bioprinting method was demonstrated through printing two different bioinks from various dECMs in a single constructs. The capability to print different dECM hydrogels simultaneously exemplified the ability of our process to reproducibly print heterogeneous 3D structures comprising various kinds of dECM. As a follow-up step, we demonstrated the printing of heterogeneous constructs (that is, printed with different kinds of dECM at different locations). However, further studies should be directed to investigate the heterogeneous tissue formation based on this approach. The ECM microenvironment plays a fundamental role in directing and mediating the differentiation of stem cell while culturing them49. There is strong evidence supporting that this instruction conveyed through both receptor ligand interactions, as well as mechanical signalling events, which in turn stimulate the important signal transduction pathways and control the cellular function49. The cell–ECM interactions are extremely complex in nature and justify the need for a tissue-specific approach while creating the native stem cell niche. Recent studies have validated this tissue-specific outlook and shown the manifestation of enhanced functions20 and intricate tissue formation22 23 when site-specific ECM scaffolds were being employed. The ECM scaffolds were shown to have the capacity to direct tissue-specific stem cell lineage commitment and maintaining the phenotype of mature cell populations50. Here, we obtain a synergistic effect through combining the beneficial effects of dECM with tissue printing process to get a cell-laden construct with engineered porosity in a tissue-specific manner. With this approach, we were able to generate a tissue analogue in vitro with either adipogenic or chondrogenic potential. In particular, we demonstrated that hASCs or hTMSCs could be printed via encapsulating in dECM pre-gel and that the in vitro culture did not modify cell survival and proliferation, but increased the commitment of stem cells towards either adipogenic lineage within dECM from fat tissue or chondrogenic lineage within cartilage dECM. Enhanced structural maturation of myoblasts was observed in constructs prepared with dECM from heart tissue. ECM has been shown to guide tissue morphogenesis through modulating cell–cell interactions51 52. Thus, we demonstrated that tissue printing with dECM bioink is an attractive option that opens up new avenues for both in vitro and in vivo tissue reconstruction. Nevertheless, there are other potential application areas where a tissue analogue structure developed by our bioprinting method is worthwhile (Fig. 1). In vitro models of human physiology and pathology can be developed using our bioprinting method and can benefit to convincingly and accurately predict the outcomes of in vivo drug administrations and potential toxic exposures, if any4. Furthermore, cancer model can be developed by recreating the microenvironmental characteristics representative of tumours in vivo with our bioprinting method with dECM that provide cancer tissue-mimicking 3D environment5. Methods Materials As synthetic biomaterial, FDA-approved PCL (molecular weight=70,000–90,000 g mol−1; Polyscience, Inc., PA, USA) was used. SDS solution 10% and 1M Tris-HCL solution were purchased from Bioneer, South Korea. Triton X-100 was purchased from Affymetrix-USB, Santa Clara, CA. Pepsin, DNAse and RNAse A were purchased from Sigma-Aldrich, St Louis, MO. Sodium chloride (NaCl) was purchased from Junsei, Japan. Unless otherwise stated, cell culture reagents were obtained from Sigma-Aldrich. Decellularization of tissues Porcine cartilage and heart from a 6-month-old pig was collected from a nearby slaughter house and used with approval from the supplier. The left ventricle was isolated from the whole porcine heart. Heart tissue decellularization was conducted by following the protocol published elsewhere with slight modification32. Tissues were cut into pieces of about 1 mm in thickness. The chopped heart tissue was stirred in 1% SDS in phosphate-buffered saline (PBS) solution for 48 h followed by treatment with 1% Triton X-100 solution for 30 min. The decellularized heart tissue was washed using PBS at least for 3 days. The hyaline cartilage was collected from porcine articular cartilage and decellularized by following the protocol published elsewhere with modification33. Briefly, the minced cartilage was placed into a hypotonic Tris-HCL buffer solution (10 mM Tris–HCL, pH 8.0) and 6 cycles of freezing (at −80 °C) and thawing (at 37 °C) were conducted. The cartilage slurry was homogenized and treated with 0.25% trypsin in PBS for 24 h at 37 °C with vigorous agitation. The trypsin solution was replaced with the fresh one at every 4 h. Trypsinized cartilage slurry was washed with a hypertonic buffer solution (1.5 M NaCl in 50 mM Tris-HCL, pH 7.6) and treated with nuclease solution (50 U ml−1 DNAse and 1 U ml−1 RNAse A in 10 mM Tris–HCL, pH 7.5) with gentle agitation at 37 °C for 4 h. To remove all the enzymes, the enzyme-treated cartilage slurry was washed with the hypotonic Tris–HCL solution for 20 h following treatment with 1% Triton X-100 solution for 24 h. The decellularized cartilage tissue was washed at least for 3 days to remove all the detergent. The adipose tissue was collected from hospital (St Mary’s Hospital, Seoul, South Korea) after liposuction of seven different female donors at the ages between 35 and 54 with informed consent and under approval from the Catholic University of Korea Institutional Review Board. The collected tissue was centrifuged to separate the oil and blood from the tissue. The adipose tissue was washed with PBS solution and decellularized with 0.5% SDS solution for 48 h with changing the solution every 12 h. Decellularized adipose tissue was then treated with isopropanol to remove the lipid for 48 h with changing the isopropanol every 12 h followed by washing several times with PBS solution. Finally, all the decellularized and delipidated tissues were treated with a solution of 0.1% peracetic acid in 4% ethanol for 4 h followed by washing several times with PBS solution and distilled water. The obtained dECMs from heart, cartilage and adipose tissues were lyophilized and stored in −20 °C freezer. Biochemical characterization of dECMs To verify the extent of decellularization, the residual DNA was measured and histological sections were analysed with H&E staining. ECM components like COL and GAGs in the decellularized tissue were also assessed. For DNA quantification, all the dECMs were digested in 1 ml of papain solution (125 μg ml−1 papain in 0.1 M sodium phosphate with 5 mM Na2-EDTA and 5 mM cysteine-HCl at pH 6.5) for 16 h at 60 °C. Papain solution without sample for using as a diluent in biochemical assays was also incubated. Native tissues of similar weight were also digested in a same manner as controls. The DNA content was determined using Hoechst 33258 assay53. The fluorescence intensity was measured to assess the amount of remaining DNA within the dECMs and the native tissue using a fluorescence spectrophotometer (TECAN, Switzerland; excitation wavelength: 360 nm, emission wavelength: 450 nm). The standard curve for DNA was generated in advance using calf thymus DNA and used for quantifying the DNA in samples. For histological evaluation, both native and decellularized tissues were fixed in 4% formalin, washed several times with distilled water, embedded in OCT compound, sectioned using a cryotome (Leica CM1850 Cryostat, Germany), stained with H&E, and observed under microscope. The GAGs content was estimated via quantifying the amount of sulphated glycosaminoglycans using 1,9-dimethylmethylene blue solution54. The absorbance was measured with microplate reader at wavelength of 492 nm. The standard curve was made using chondroitin sulphate A in advance and used for estimating the sulphated glycosaminoglycans in samples. The total COL content was determined via a conventional hydroxyproline assay55. The absorbance of the samples was measured at 550 nm and quantified by referring to a standard curve made in advance with hydroxyproline. Preparation of dECM solution and pre-gel Lyophilized dECM was crushed into powder using a mortar and pestle with the help of liquid N2. Required amount of dECM powder was weighed and digested in a solution of 0.5 M acetic acid with 10 mg of pepsin for 100 mg dECM (P7125, Sigma-Aldrich) for 48 h. After complete solubilization of dECM, the solution was centrifuged (only for adipose dECM) to remove the particulate things if any and used for further examinations. The pH of dECM solution was in the range of 2.8–3. The pH was adjusted with dropwise addition of cold 10 M NaOH solution while maintaining the temperature below 10 °C to avoid gelation of the dECM. The pH-adjusted dECM pre-gel was stored in refrigerator at 4 °C. Rheological characterization Rheological investigations were conducted on an Advanced Rheometric Expansion System (TA Instrument, USA) using 25 mm diameter plate geometry. To assess the viscosity, steady shear sweep analysis of various dECM pre-gels were performed at a constant temperature of 15 °C. Dynamic frequency sweep analysis was conducted to measure the frequency-dependent storage (G′) and loss (G″) moduli of various dECM gels in the range of 1–1,000 rad s–1 at 2% strain after incubation for 30 min at 37 °C. A temperature sweep was used to study the gelation kinetics of dECM pre-gels. The complex modulus was measured at each temperature while the dECM pre-gels were subjected to a temperature ramp in the range of 4–37 °C with a increment rate of 5 °C min−1. In addition, at 15 °C, the dECM pre-gels were held for 5 min. All measurements were conducted in triplicate. Cell encapsulation in the dECM pre-gel hASCs and hTMSCs were used for assessment of the effectiveness of adECM and cdECM on adipogenic and chondrogenic differentiation of adult stem cells, respectively. The cells were isolated and cultured by a previously described method41 42 56. Rat myoblast cells (L6, ATCC CRL-1458) were used for verifying functional enhancement of myoblasts in the hdECM. hASCs and L6 cells were cultured in the DMEM high glucose supplemented with 10% fetal bovine serum and 1% antibiotics (penicillin/streptomycin), and hTMSCs were cultured in the αMEM supplemented with 10% fetal bovine serum and 1% antibiotics (penicillin/streptomycin). All the cells used were from passage 2 to 5. Cultured cells were detached from the tissue culture plate with 0.25% trypsin solution, centrifuged at 1,500 r.p.m. for 3 min, suspended in fresh media and counted. To maintain the same osmotic pressure, 10 × concentrated culture media was added to the pH-adjusted dECM pre-gel (1/10th volume) and the suspended cells were mixed thoroughly with the pH-adjusted ECM pre-gel. The concentration of cells used was from 1 to 5 × 106 cells ml−1. All processes were conducted on ice. The prepared bioink was loaded in a sterilized syringe for cell printing. Printing of cell-laden construct A MtoBS, a bioprinting system developed in our lab, with six different syringe holders and capable of simultaneously dispensing six different materials, was used for printing of cell-laden construct. PCL was loaded in a syringe and heated to 80 °C to melt the polymer. The dECM pre-gel encapsulated cells were loaded to the other syringe and maintained at temperatures below 15 °C. Cell-printed constructs were fabricated according to the method described elsewhere39. We used two syringes for constructing each structure, one containing PCL and maintained at 80 °C, the other one loaded with cell-laden pre-gel and maintained at temperatures below 15 °C. The applied pneumatic pressure was in the range of 400–650 kPa for fabrication of the PCL framework. The cell-laden pre-gel was dispensed using a plunger-based low-dosage dispensing system. In the process of printing synthetic polymer, temperature of the molten PCL comes down very fast in the air gap before reaching the stage. Cell viability and apoptosis test Cells were under a relatively high shear-stressed environment when the mechanical perturbations were applied to the cells by dispensing bioinks. Therefore, we verified the viability and apoptosis of cells after the printing. For cell apoptosis assay, we selected hdECM as a representative bioink due to its highest viscosity among the developed bioinks. The hTMSCs were encapsulated at a concentration of 5 × 105 cells ml−1 into 3% hdECM pre-gel and the prepared bioink was loaded into the piston syringe. The grid type structure of 100 μm in thickness was fabricated with the hdECM bioink at 2 s−1 shear rate, which was selected to simulate the shear rate generated during printing. Finally, the fabricated structure was cross-linked by incubating at 37 °C temperature for 30 min. After 24 h of incubation in the complete medium, the structure was fixed by 4% paraformaldehyde and treated with the DeadEnd Fluorometric TUNEL System kit (Promega, WI) following manufacturer’s protocol with little modification. We treated all reagents for a longer time than that of the time mentioned in the protocol due to the larger thickness of the structure. The TUNEL system can measure the fragmented DNA of apoptotic cells by catalytically binding fluorescein-12-dUTP; thus, the fluorescein-12-dUTP-labelled DNA in the fabricated structure was visualized by confocal microscope (Olympus FluoView FV1000, Tokyo, Japan). Cell nucleus was counterstained by DAPI (GBI Labs, WA). To conduct viability test, cell-printed structures with dECM was evaluated by staining with live and dead assay kit (LIVE/DEAD Cell Viability Assays, Invitrogen Life Technologies, USA) following manufacturer’s protocol. Gene expression analysis through RT–PCR After culturing the constructs for the specified time period, total RNA of cells was isolated using TRIzol reagent (Invitrogen Life Technologies, USA) following the manufacturer’s protocol. RNA concentration was measured using a Nanodrop (Thermo Scientific, USA). Reverse transcription was performed with the cDNA synthesis kit (Thermo Scientific, USA) following manufacturer’s instructions. Gene expression was analysed quantitatively with SYBR-green using 7500 Real-Time PCR system (Applied Biosystems, Life Technologies, USA). The primers and probes for collagen type II (COL2A1), SOX9, protein tyrosine kinase 2, Integrin β1, PPARγ, C/EBPα, LPL, Myh6, Actn1, beta-actin, 18S ribosomal RNA and GAPDH were designed on the basis of published gene sequences (NCBI and PubMed). We checked the expression patterns of various endogenous control genes (GAPDH, beta-actin and 18S ribosomal RNA) of cells cultured on TCP over time. GAPDH has shown low level of variance among samples over time than the other genes (Supplementary Fig. 7). Thus, we have chosen GAPDH as an endogenous control for our study. Expression levels for each gene were normalized with GAPDH and analysed using the 2−ΔΔCT method. Each sample was assessed in triplicate. Immunohistological study The cell-printed structure was harvested and fixed with a solution of 4% paraformaldehyde after 14 days culture. The structure was embedded in OCT compound and sectioned using cryotome in 10-μm thickness. The sliced samples were washed repeatedly with PBS solution to remove OCT compound and then permeabilized with a solution of 0.1% Triton X-100 in PBS for 5 min. To reduce non-specific background, sections were treated with 0.2% bovine serum albumin solution in PBS for 20 min. For visualization of differentiation and maturation of stromal cells to chondrogenic cells, sections were immunostained using primary antibody against anti-collagen II antibody (1:100; Abcam, UK) followed by incubation with Alexa Fluor 594 goat anti-mouse antibody (1:100; Invitrogen, CA). Sections were also stained with phalloidin-FITC solution (1:100; Sigma-Aldrich, MA) for 40 min and DAPI for 15 min. The adipogenic cell-printed structure was harvested and fixed with a solution of 4% paraformaldehyde after 14 days culture and stained following similar protocol using PPARγ (1:100; Santa Cruz Biotechnology Inc, Catalogue No. sc-81152) and Collagen IV (1:100; Santa Cruz Biotechnology Inc, COL4A5 (H-234): sc-11360). Secondary antibody of Alexa Fluor 594 goat anti-mouse antibody (1:100; Invitrogen, CA) and Alexa Fluor 488 goat anti-rabbit antibody (1:100; Invitrogen, CA) were used and counterstained with DAPI. For detection of the myotube formation in the hdECM construct, sections were stained with anti-heavy chain cardiac myosin antibody (1:100; Abcam), followed by incubation with secondary goat anti-mouse-Alexa 594 antibody (1:100) and counterstained with DAPI. For F-actin expression analysis, the cells were seeded on the each material (tissue culture plate, TCP; COL and cdECM gel), and were cultured for 3 days and stained with phalloidin-FITC solution for 40 min and DAPI for 15 min. Stained samples were visualized and images were captured using a confocal microscope. Statistical analysis All data are expressed as mean±s.d. All statistical analysis was performed by using SigmaPlot software (ver. 10.0, Systat Software Inc., San Jose, CA). Statistical significance was determined by a one-way analysis of variance. Differences were considered to be statistically significant at P<0.05. Author contributions Decellularization, solubilization and gelation of dECM, printing of cell-laden construct, and gene expression were performed by F.P., J.J. and D.-H.H. The paper was written by F.P., J.J. and D.-W.C. The study was conceived and designed F.P., J.J., J.-H.S., and D.W.C. The hTMSCs and hASCs were provided by S.W.K. and J.-W.R., respectively. The study was directed by D.W.C., J.-W.R. and D.-H.K. Additional information How to cite this article: Pati, F. et al. Printing three-dimensional tissue analogues with decellularized extracellular matrix bioink. Nat. Commun. 5:3935 doi: 10.1038/ncomms4935 (2014). Supplementary Material Supplementary Figures. Supplementary Figures 1-14 Supplementary Movie 1 Printing of cell-laden structure using dECM bioink Supplementary Movie 2 Printing of heterogeneous structure using two kinds of dECM bioinks
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            Porous scaffold design for tissue engineering.

            A paradigm shift is taking place in medicine from using synthetic implants and tissue grafts to a tissue engineering approach that uses degradable porous material scaffolds integrated with biological cells or molecules to regenerate tissues. This new paradigm requires scaffolds that balance temporary mechanical function with mass transport to aid biological delivery and tissue regeneration. Little is known quantitatively about this balance as early scaffolds were not fabricated with precise porous architecture. Recent advances in both computational topology design (CTD) and solid free-form fabrication (SFF) have made it possible to create scaffolds with controlled architecture. This paper reviews the integration of CTD with SFF to build designer tissue-engineering scaffolds. It also details the mechanical properties and tissue regeneration achieved using designer scaffolds. Finally, future directions are suggested for using designer scaffolds with in vivo experimentation to optimize tissue-engineering treatments, and coupling designer scaffolds with cell printing to create designer material/biofactor hybrids.
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              A review on stereolithography and its applications in biomedical engineering.

              Stereolithography is a solid freeform technique (SFF) that was introduced in the late 1980s. Although many other techniques have been developed since then, stereolithography remains one of the most powerful and versatile of all SFF techniques. It has the highest fabrication accuracy and an increasing number of materials that can be processed is becoming available. In this paper we discuss the characteristic features of the stereolithography technique and compare it to other SFF techniques. The biomedical applications of stereolithography are reviewed, as well as the biodegradable resin materials that have been developed for use with stereolithography. Finally, an overview of the application of stereolithography in preparing porous structures for tissue engineering is given. 2010 Elsevier Ltd. All rights reserved.
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                Author and article information

                Journal
                Journal of Biological Engineering
                J Biol Eng
                Springer Science and Business Media LLC
                1754-1611
                December 2015
                March 1 2015
                December 2015
                : 9
                : 1
                Article
                10.1186/s13036-015-0001-4
                032b3695-a869-46b4-9101-10cde410d5f8
                © 2015

                http://creativecommons.org/licenses/by/4.0

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