A 3D bioprinting system to produce human-scale tissue constructs with structural integrity

Journal name:
Nature Biotechnology
Volume:
34,
Pages:
312–319
Year published:
DOI:
doi:10.1038/nbt.3413
Received
Accepted
Published online

Abstract

A challenge for tissue engineering is producing three-dimensional (3D), vascularized cellular constructs of clinically relevant size, shape and structural integrity. We present an integrated tissue–organ printer (ITOP) that can fabricate stable, human-scale tissue constructs of any shape. Mechanical stability is achieved by printing cell-laden hydrogels together with biodegradable polymers in integrated patterns and anchored on sacrificial hydrogels. The correct shape of the tissue construct is achieved by representing clinical imaging data as a computer model of the anatomical defect and translating the model into a program that controls the motions of the printer nozzles, which dispense cells to discrete locations. The incorporation of microchannels into the tissue constructs facilitates diffusion of nutrients to printed cells, thereby overcoming the diffusion limit of 100–200 μm for cell survival in engineered tissues. We demonstrate capabilities of the ITOP by fabricating mandible and calvarial bone, cartilage and skeletal muscle. Future development of the ITOP is being directed to the production of tissues for human applications and to the building of more complex tissues and solid organs.

At a glance

Figures

  1. ITOP system.
    Figure 1: ITOP system.

    (a) The ITOP system consists of three major units: (i) 3-axis stage/controller, (ii) dispensing module including multi-cartridge and pneumatic pressure controller and (iii) a closed acrylic chamber with temperature controller and humidifier. (b) Illustration of basic patterning of 3D architecture including multiple cell-laden hydrogels and supporting PCL polymer. (c) CAD/CAM process for automated printing of 3D shape imitating target tissue or organ. A 3D CAD model developed from medical image data generates a visualized motion program, which includes instructions for XYZ stage movements and actuating pneumatic pressure to achieve 3D printing.

  2. 2D/3D patterning using the ITOP system.
    Figure 2: 2D/3D patterning using the ITOP system.

    (a,b) 2D patterning of 'WFIRM' characters written by cell-laden hydrogels through the integrated organ printing. Microscopic (a) and fluorescent images (b) of 'WFIRM' characters, which were produced using cells labeled with Dil and DiO. (ch) Two basic types of 3D patterning: type I pattern (ce) and type II pattern (fh). Two types of 3D patterning, including cell-A (red), cell-B (blue) and PCL (green), were fabricated by the integrated organ printing (c,f); photographs (d,g) and fluorescent image (e,h) of the 3D printed patterns. (i) Cell viability was over 95% on day 0 and then maintained on days 3 and 6 (n = 3). (j) Cell proliferation results showed that the number of cells continuously increased over a 15-d period, and no significant differences between the control and the printed constructs were observed (n = 5). Error bars, mean ± s.d.

  3. Mandible bone reconstruction.
    Figure 3: Mandible bone reconstruction.

    (a) 3D CAD model recognized a mandible bony defect from human CT image data. (b) Visualized motion program was generated to construct a 3D architecture of the mandible bone defect using CAM software developed by our laboratory. Lines of green, blue and red colors indicate the dispensing paths of PCL, Pluronic F-127 and cell-laden hydrogel, respectively. (c) 3D printing process using the integrated organ printing system. The image shows patterning of a layer of the construct. (d) Photograph of the 3D printed mandible bone defect construct, which was cultured in osteogenic medium for 28 d. (e) Osteogenic differentiation of hAFSCs in the printed construct was confirmed by Alizarin Red S staining, indicating calcium deposition.

  4. Calvarial bone reconstruction.
    Figure 4: Calvarial bone reconstruction.

    (a) Visualized motion program (top) used to print a 3D architecture of calvarial bone construct. Green and red color lines indicate the dispensing paths of the PCL/TCP mixture and cell-laden hydrogel, respectively. Photograph of the printed calvarial bone construct (bottom). (b) Scanning electron microscope images of the printed bone constructs. (c) Photographs of the printed bone constructs at day 0 (top) and 5 months (bottom) after implantation. (dl) Histological and immunohistological images of nontreated (df), scaffold only without cells (gi) and hAFSCs-printed construct at 5 months after implantation (jl). H&E staining (d,g,j), modified tetrachrome staining (e,h,k) and vWF immunostaining (f,i,l). Tetrachrome staining: red, mature bone; blue, osteoid and lining of lacunae. vWF immunofluorescent image: red, blood vessel. NB: new bone; PCL/TCP: remaining scaffold.

  5. Ear cartilage reconstruction.
    Figure 5: Ear cartilage reconstruction.

    (af) In vitro bioprinted ear construct. (a) 3D CAD of a human ear. (b) Visualized motion program used to print 3D architecture of human ear. The motion program was generated by using 3D CAD model. Lines of green, blue and red indicate dispensing paths of PCL, Pluronic F-127 and cell-laden hydrogel, respectively. (c) 3D printing process using the integrated organ printing system (Supplementary Movie 1). The image shows patterning of a layer of the construct. (d,e) Photographs of the 3D printed ear cartilage construct with sacrificial Pluronic F-127 (d) and after removing sacrificial material by dissolving with cold medium (e). (f) Safranin-O staining of the 3D printed cartilage constructs with microchannels (porous; left) and without microchannels (nonporous; right) after culture in chondrogenic medium for 5 weeks in vitro. The constructs with microchannels showed the production of new cartilaginous matrix throughout the entire constructs, whereas the constructs without microchannels showed limited tissue formation due to limited diffusion of nutrients and oxygen. The staining indicates the production of GAGs. (g) Safranin-O staining, Alcian Blue staining and immunohistochemistry for type II collagen of the 3D printed ear cartilage constructs after culture in chondrogenic medium for 5 weeks in vitro. Histological images of the samples showed the production of a new cartilaginous matrix within the 3D printed constructs. The chondrocytes in the newly formed tissue demonstrated similar morphological characteristics to those in native cartilage, with cells located within typical chondrocyte lacunae, surrounded by cartilaginous matrix. The newly formed matrix generated in the constructs stained intensely with Safranin-O and Alcian Blue, showing the presence of sulfated proteoglycans. Immunohistochemical staining indicated the presence of type II collagen in the constructs. Human ear was used a positive control. (hm) In vivo bioprinted ear construct. (h,i) Gross appearance at 1 month after implantation (h), Safranin-O staining and collagen type II immunostaining (i) of the retrieved ear construct at 1 month and 2 months after implantation. (j) GAG contents of the bioprinted ear cartilage tissues after 1 and 2 months of implantation. Error bars, mean ± s.d. (k) Gross examination of bending testing of the bioprinted ear constructs: pre-implantation vs. 1-month implantation. (l,m) Stress-strain curve of pre-implanted construct (l) and1-month implanted construct under four-cycle three-point bending test (m).

  6. Skeletal muscle reconstruction.
    Figure 6: Skeletal muscle reconstruction.

    (a–g) In vitro bioprinted muscle. (a) Designed fiber bundle structure for muscle organization. PCL pillars (green) were used to maintain the structure and to induce the compaction phenomenon for cell alignment. (b) Visualized motion program for 3D printing muscle construct. Lines of green, white and blue indicate the dispensing paths of PCL, cell-laden hydrogel and sacrificial material, respectively. (c) 3D patterning outcome of designed muscle organization (left) before and (after) removing the sacrificial material (Pluronic F127). The printed construct was cross-linked with thrombin solution to induce gelation of fibrinogen and the uncross-linked sacrificial material was removed by dissolving with cold medium. (d,e) The PCL pillar structure is essential to stabilize the 3D printed muscle organization and to induce a compaction phenomenon of the patterns of the cell-laden hydrogel that causes cell alignment in a longitudinal direction of the printed constructs; without PCL pillar (d) and with PCL pillar (e). The cells with PCL pillar showed unidirectionally organized cellular morphologies that are consistently aligned along the longitudinal axis of the printed construct, which is in contrast to the randomly oriented cellular morphologies without PCL pillar. (f) The live/dead staining of the encapsulated cells in the fiber structure indicates high cell viability after the printing process (green: live cells; red: dead cells). (g) Immunofluorescent staining for myosin heavy chain of the 3D printed muscle organization after 7-d differentiation. The encapsulated myoblasts aligned along the longitudinal direction of the fiber structure. (hm) Structural maintenance and host nerve integration of the bioprinted muscle construct in in vivo study. (h) Schematic diagram of ectopic implantation of bioprinted muscle construct in vivo. (ik) The bioprinted muscle construct was subcutaneously implanted with the dissected CPN inserted into the printed muscle construct, and the harvested implants after 2 weeks of implantation showed the presence of organized muscle fibers and innervating capability (α-BTX-positive structures) within the implanted construct, as confirmed by immunostaining using skeletal muscle markers (desmin (j) and MHC+ and α-BTX+ structure (arrows) (k)). (l) The evidence of nerve integration was demonstrated with double staining of neurofilament (NF+) and α-BTX+ structure (arrows). (m) The vascularization of implanted muscle construct was confirmed by vWF immunostaining. (n) Functional assessment of bioprinted muscle constructs after 4 weeks of implantation (*P < 0.05). Positive control: the normal gastrocnemius muscle; negative control: the gluteus muscle after dissected CPN.

  7. Integrated tissue and organ printing (ITOP) system
    Supplementary Fig. 1: Integrated tissue and organ printing (ITOP) system

    The ITOP system consists of three major units; 1) 3-axis stage/controller, 2) dispensing module including multi-cartridge and pneumatic pressure controller, and 3) a closed acrylic chamber with temperature controller and humidifier.

  8. Optimization of composite hydrogel system for 3D bioprinting
    Supplementary Fig. 2: Optimization of composite hydrogel system for 3D bioprinting

    Dispensing rates (dispensed volume per unit time) with different concentrations of (a) gelatin (n=30) and (b) HA (n=30). The dispensing rate could by increased or decreased by decreasing or increasing the concentration of gelatin, respectively. Irregularities in dispensing rates to varying gelatin concentrations were directly related to the coefficient of variation (COV); where a COV of greater than 30% was associated with uneven dispensing of the hydrogel. However, introduction of HA to gelatin significantly improved uniformity of dispensing rate with low COV values. *Coefficient of variation (COV) was calculated by dividing average with standard deviation.

  9. Optimization of PCL and TCP ratio
    Supplementary Fig. 3: Optimization of PCL and TCP ratio

    (a) Compression modulus and (b) water uptake ability of the printed PCL/TCP constructs with different ratios (*P<0.05, n = 3). (c) Quantification of calcium content showing the osteogenic differentiation of hAFSCs seeded on the PCL/TCP constructs with different ratios (no significance, n = 3).

  10. Vascularization of 3D printed ear
    Supplementary Fig. 4: Vascularization of 3D printed ear

    The engineered cartilage tissues showed vascularization of the implanted constructs in the periphery region at 1 and 2 months after implantation, as confirmed by vWF immunostaining, but similar to normal cartilage tissue, no vascularization was noted in the central region.

  11. Immunofluorescent images of 3D printed muscle organization
    Supplementary Fig. 5: Immunofluorescent images of 3D printed muscle organization

    3D printed muscle structures were cultured in the growth medium up to 3 days, and then induced myotube formation in the differentiation medium for 7 days. Live/dead staining of the printed muscle organization at (a) 1 day and (b) 3 days. (c) Immunofluorescent staining for MHC of the printed muscle organization at 7 days after cell differentiation. The myotubes formed in the printed constructs showed unidirectionally organized myotubes that are consistently aligned along the longitudinal axis of the printed organization.

Videos

  1. Supplementary Movie 1
    Video 1: Supplementary Movie 1
    Supplementary Movie 1

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Author information

Affiliations

  1. Wake Forest Institute for Regenerative Medicine, Wake Forest School of Medicine, Medical Center Boulevard, Winston-Salem, North Carolina, USA.

    • Hyun-Wook Kang,
    • Sang Jin Lee,
    • In Kap Ko,
    • Carlos Kengla,
    • James J Yoo &
    • Anthony Atala

Contributions

H.-W.K., S.J.L., J.J.Y. and A.A. developed the concept of the integration tissue and organ printing (ITOP) system and designed all experiments. H.-W.K. performed in vitro experiments and composite hydrogel development, analyzed data and wrote the manuscript. C.K. performed in vivo experiments of the printed cartilage and bone constructs and analyzed data. I.K.K. performed in vivo experiments of the printed skeletal muscle construct and analyzed data. S.J.L., J.J.Y. and A.A. analyzed data and wrote the manuscript. A.A. provided direction and supervised the project. S.J.L., J.J.Y. and A.A. edited the manuscript.

Competing financial interests

The authors declare no competing financial interests.

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Supplementary information

Supplementary Figures

  1. Supplementary Figure 1: Integrated tissue and organ printing (ITOP) system (237 KB)

    The ITOP system consists of three major units; 1) 3-axis stage/controller, 2) dispensing module including multi-cartridge and pneumatic pressure controller, and 3) a closed acrylic chamber with temperature controller and humidifier.

  2. Supplementary Figure 2: Optimization of composite hydrogel system for 3D bioprinting (56 KB)

    Dispensing rates (dispensed volume per unit time) with different concentrations of (a) gelatin (n=30) and (b) HA (n=30). The dispensing rate could by increased or decreased by decreasing or increasing the concentration of gelatin, respectively. Irregularities in dispensing rates to varying gelatin concentrations were directly related to the coefficient of variation (COV); where a COV of greater than 30% was associated with uneven dispensing of the hydrogel. However, introduction of HA to gelatin significantly improved uniformity of dispensing rate with low COV values. *Coefficient of variation (COV) was calculated by dividing average with standard deviation.

  3. Supplementary Figure 3: Optimization of PCL and TCP ratio (54 KB)

    (a) Compression modulus and (b) water uptake ability of the printed PCL/TCP constructs with different ratios (*P<0.05, n = 3). (c) Quantification of calcium content showing the osteogenic differentiation of hAFSCs seeded on the PCL/TCP constructs with different ratios (no significance, n = 3).

  4. Supplementary Figure 4: Vascularization of 3D printed ear (350 KB)

    The engineered cartilage tissues showed vascularization of the implanted constructs in the periphery region at 1 and 2 months after implantation, as confirmed by vWF immunostaining, but similar to normal cartilage tissue, no vascularization was noted in the central region.

  5. Supplementary Figure 5: Immunofluorescent images of 3D printed muscle organization (482 KB)

    3D printed muscle structures were cultured in the growth medium up to 3 days, and then induced myotube formation in the differentiation medium for 7 days. Live/dead staining of the printed muscle organization at (a) 1 day and (b) 3 days. (c) Immunofluorescent staining for MHC of the printed muscle organization at 7 days after cell differentiation. The myotubes formed in the printed constructs showed unidirectionally organized myotubes that are consistently aligned along the longitudinal axis of the printed organization.

Video

  1. Video 1: Supplementary Movie 1 (16 MB, Download)
    Supplementary Movie 1

PDF files

  1. Supplementary Text and Figures (546 KB)

    Supplementary Figures 1–5

  2. Supplementary Table 1 (128 KB)

    Supplementary Table 1

  3. Supplementary Source Code (9.23 MB)

Additional data