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4. intravascular andmultivascular
design freedoms with photopolymerizable hydrogels by using
food dye additives as
biocompatible yet potent photoabsorbers for projection
stereolithography.We demonstrate
monolithic transparent hydrogels, produced in minutes,
comprising efficient intravascular
3D fluid mixers and functional bicuspid valves.We further
elaborate entangled vascular
networks from space-filling mathematical topologies and
explore the oxygenation and
flow of human red blood cells during tidal ventilation and
distension of a proximate airway.
In addition, we deploy structured biodegradable hydrogel
carriers in a rodent model of chronic
liver injury to highlight the potential translational utility of this
materials innovation.
T
he morphologies of the circulatory and
pulmonary systems are physically and evo-
lutionarily entangled (1). In air-breathing
vertebrates, these bounded and conserved
vessel topologies interact to enable the
oxygen-dependent respiration of the entire or-
ganism (2–4). To build and interrogate soft hy-
drogels containing such prescribed biomimetic
and multivascular architectures, we sought to
use stereolithography (fig. S1) (5), commonly em-
ployed to efficiently convert photoactive liquid
resins into structured plastic parts through lo-
calized photopolymerization (6, 7). Comparedwith
extrusion 3D printing, which deposits voxels in
a serial fashion (8–12), photocrosslinking can
be highly parallelized via image projection to
5. simultaneously and independently address mil-
lions of voxels per time step. In stereolithography,
xy resolution isdeterminedby the lightpath,whereas
z resolution is dictated by light-attenuating ad-
ditives that absorb excess light and confine the
polymerization to the desired layer thickness,
thereby improving pattern fidelity. In the ab-
sence of suitable photoabsorber additives, 3D
photopatterning of soft hydrogels has been lim-
ited in the types of patterns that can be generated
(13–16) or has required complex, expensive, and
low-throughput microscopy to enhance z reso-
lution via the multiphoton effect (17–19). How-
ever, common light-blocking chemicals used for
photoresist patterning or plastic part fabrication,
such as Sudan I, are not suitable for biomanu-
facturing owing to their known genotoxic and
carcinogenic characteristics (20). Therefore, we
hypothesized that the identification of nontoxic
light blockers for projection stereolithography
could provide a major advance to the architec-
tural richness available for the design and gen-
eration of widely used biocompatible hydrogels.
Here, we establish that synthetic and natural
food dyes, widely used in the food industry, can
be applied as potent biocompatible photoabsorbers
to enable the stereolithographic production of
hydrogels containing intricate and functional
vascular architectures. We identified candidate
photoabsorbers among food additives whose ab-
sorbance spectra encompass visible light wave-
lengths that can be used for biocompatible
photopolymerization. We initially sought to gen-
eratemonolithic hydrogels, composed primarily
6. of water and poly(ethylene glycol) diacrylate
[PEGDA, 6 kDa, 20weight% (wt%)], with a 1-mm
cylindrical channel oriented perpendicular to the
light-projection axis. The fabrication of even this
trivial design cannot be easily realized because of
the dilute nature of such aqueous formulations,
in which the low mass fraction of crosslinkable
groups and the requisite longer polymerization
times result in inadvertent polymerization and
solidification within the narrow void spaces that
were designed to be hollowperfusable vasculature
(figs. S2 to S4).
We determined that aqueous pre-hydrogel
solutions containing tartrazine (yellow food
coloring FD&C Yellow 5, E102), curcumin (from
turmeric), or anthocyanin (from blueberries) can
each yield hydrogels with a patent vessel (figs. S2
to S5). In addition to these organic molecules,
inorganic gold nanoparticles (50 nm), widely
regarded for their biocompatibility and light-
attenuating properties (21), also function as an
effective photoabsorbing additive to generate
perfusable hydrogels (fig. S4).
To understand how these photoabsorbers
affect the gelation kinetics of photopolymeriz-
able hydrogels, we performed photorheological
characterization with short-duration light ex-
posures, which indicate that these additives cause
a dose-dependent delay in the induction of photo-
crosslinking (figs. S2D and S4E). Saturating light
exposures that extend beyond the reaction termi-
nation point demonstrate that suitable additives
did not ultimately interfere with the reaction be-
7. cause hydrogels eventually reached an equivalent
storage modulus independent of the additive
concentration (figs. S2D and S4E). We selected
tartrazine as a photoabsorber for further studies.
In addition to its low toxicity in humans and
broadutility in the food industry (22), we observed
that this hydrophilic dye is easily washed out of
generated hydrogels (70% elutes within 3 hours
for small gels), resulting in nearly transparent
constructs suitable for imaging (fig. S2E). Some
tartrazine may also be degraded during poly-
merization, as tartrazine is known to be sensitive
to free radicals (23). Submerging gels in water
or saline solution to remove soluble tartrazine
also flushes the vascular topology and removes
unreacted pre-hydrogel solution. In contrast to
tartrazine, curcumin is lipophilic and does not
wash out in aqueous solutions; anthocyanin has
a peak absorbance far fromour intended 405-nm
light source, requiring high concentrations for
suitable potency; and gold nanoparticles are
physically entrapped and make transmission or
fluorescence microscopy impractical (fig. S4E).
We assessed whether this materials insight
could similarly impart new architectural freedoms
tomore-advanced photoactivematerials. Photo-
absorber additives are necessary and sufficient
to enable vessel construction in thiol-ene step-
growth photopolymerization (24) of hydrogels
and in a continuous liquid interface production
(6) workflow for the generation of hydrogels (fig.
S5). We observed strong lamination between ad-
jacent fabricated layers and a rapid response
of the patterned hydrogel tomechanical deforma-
tions (fig. S6). This facile generation of soft
8. hydrogels with patent cylindrical vessels oriented
orthogonal to the light-projection axis sug-
gests an extensive design flexibility toward the
generation of complex vascular topologies, and
RESEARCH
Grigoryan et al., Science 364, 458–464 (2019) 3 May 2019 1 of
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1Department of Bioengineering, Rice University, Houston, TX
77005, USA. 2Department of Bioengineering, University of
Washington, Seattle, WA 98195, USA. 3Institute for Stem Cell
and Regenerative Medicine, University of Washington,
Seattle, WA 98195, USA. 4Department of Pathology,
University of Washington, Seattle, WA 98195, USA.
5Department of Biomedical Engineering, Duke University,
Durham, NC 27708, USA. 6Nervous System, Somerville, MA
02143, USA. 7Department of Biomedical Engineering, Rowan
University, Glassboro, NJ 08028, USA.
*These authors contributed equally to this work. †Present
address:
Computational Science and Engineering Division, Oak Ridge
National Laboratory, Oak Ridge, TN 37830, USA.
‡Corresponding author. Email: [email protected] (K.R.S.);
[email protected] (J.S.M.)
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the optical clarity of resultant hydrogels implies
imaging methodologies suitable for characteri-
zation and validation of fluid flows.
Next, we investigated the ability to form
hydrogels containing functional intravascular
topologies. We first explored chaotic mixers:
intravascular topologies that homogenize fluids
as a result of interactions between fluid flow
streams and the vessel geometry (25, 26). Where-
as macroscale static mixers have found broad
utility in industrial processes (27) because of their
unparalleled efficiency, translation of intravascular
static mixers into microfluidic systems has been
difficult to implement, owing to their complex
3D topology. To this end, we generated mono-
lithic hydrogels with an integrated static mixer
composed of 3D twisted-fin elements (150 mm
thick) of alternating chirality inside a 1-mm
cylindrical channel. We applied laminar fluid
streams to the static mixer at a low Reynolds
number (0.002) and observed rapid mixing per
unit length (Fig. 1A) and as a function of fin
number (fig. S7). The elasticity and compliance of
PEG-based hydrogels (fig. S6) enabled the facile
generation of a 3D functional bicuspid venous
valve (Fig. 1B).We observed that the valve leaflets
10. are dynamic, respond rapidly to pulsatile antero-
grade and retrograde flows, and promote the for-
mation of stable mirror image vortices in the
valve sinuses (Fig. 1B and movie S1) according to
established mappings of native tissue (28, 29).
Solid organs contain distinct fluid networks
that are physically and chemically entangled,
providing the rich extracellular milieu that is
a hallmark of multicellular life. The ability to
fabricate such multivascular topologies within
biocompatible and aqueous environments could
enable a step change in the fields of biomaterials
and tissue engineering. A first objective is the
development of an efficient framework to design
entangled networks that can provide suitable
blueprints for their fabrication within hydrogels.
Separate vascular networks must not make a
direct fluid connection or they would topolog-
ically reduce to a single connected network. We
find that mathematical space-filling and fractal
topology algorithms provide an efficient para-
metric language to design complex vascular blue-
prints and a mathematical means to design a
second vascular architecture that does not inter-
sect the first (Fig. 2). We demonstrate a selection
of hydrogels (20 wt %, 6-kDa PEGDA) contain-
ing entangled vascular networks based on 3D
mathematical algorithms (Fig. 2, A to D): a helix
surrounding an axial vessel, 1° and 2° Hilbert
curves, a bicontinuous cubic lattice (based on a
Schwarz P surface), and a torus entangled with
a torus knot. Perfusion with colored dyes and
micro-computed tomography (mCT) analysis de-
monstrate pattern fidelity, vascular patency, and
11. Grigoryan et al., Science 364, 458–464 (2019) 3 May 2019 2 of
7
Fig. 1. Monolithic hydrogels with functional intravascular
topologies.
(A) Monolithic hydrogels with a perfusable channel containing
integrated fin elements of alternating chirality. These static
elements rapidly promote fluid dividing and mixing (as shown
by
fluorescence imaging), consistent with a computational model
of flow
(scale bars, 1 mm). (B) Hydrogels with a functional 3D bicuspid
valve integrated into the vessel wall under anterograde and
retrograde flows (scale bars, 500 mm). Particle image
velocimetry
demonstrates stable mirror image vortices in the sinus region
behind open valve leaflets.
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12. fluidic independence between the two networks
(Fig. 2, A to D, and movie S2).
We sought to evaluate the efficiency of inter-
vascular interstitial transport by measuring the
delivery of oxygen from a source vessel to per-
fused human red blood cells (RBCs) flowing in
an adjacent 3D topology. We tessellated the en-
tangled helical topology shown in Fig. 2A along
a serpentine path while maintaining the inter-
vessel distance at 300 mm (Fig. 2E). Perfusion of
deoxygenated RBCs [oxygen partial pressure
(PO2) ≤ 40 mmHg; oxygen saturation (SO2) ≤
45%] into the helical channel during ventilation of
the serpentine channel with humidified gaseous
oxygen (7 kPa) caused a noticeable color change
of RBCs fromdark red at the inlet to bright red at
the outlet (Fig. 2, E and F). Collection of perfused
RBCs showed significantly higher SO2 and PO2
relative to deoxygenated RBCs loaded at the
inlet and negative control gels ventilated with
humidified nitrogen gas (Fig. 2G and fig. S8).
Although this serpentine-helix design demon-
strates the feasibility of intervascular oxygen trans-
port between 3D entangled networks, we sought
to introduce additional structural features of
native distal lung into a bioinspired model of
alveolar morphology and oxygen transport. In
particular, the realization of 3D hydrogels that
contain branching networks and that can sup-
port mechanical distension during cyclic ventila-
13. tion of a pooled airway could enable investigations
of the performance of lung morphologies derived
from native structure (30) and could provide a
complete workflow for the development and ex-
amination of new functional topologies. Over the
past several decades, alveolar morphology has
been approximated mathematically as 3D space-
filling tessellations of polyhedra (31–34). However,
the translation of these ideas into useful blue-
prints has remained nontrivial because of the
need for efficient space-filling tessellations and
an ensheathing vasculature that closely tracks
the curvature of the 3D airway topography. Our
solution is to calculate a 3D topological offset of
the airway (moving each face in its local normal
Grigoryan et al., Science 364, 458–464 (2019) 3 May 2019 3 of
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Fig. 2. Entangled vascular networks. (A to D) Adaptations of
mathematical
space-filling curves to entangled vessel topologies within
hydrogels
(20 wt % PEGDA, 6 kDa): (A) axial vessel and helix, (B)
interpenetrating
Hilbert curves, (C) bicontinuous cubic lattice, and (D) torus and
(3,10) torus
knot (scale bars, 3 mm). (E) Tessellation of the axial vessel and
its
encompassing helix along a serpentine pathway. The photograph
is a
top-down view of a fabricated hydrogel with oxygen and RBC
delivery to
respective vessels. During perfusion, RBCs change color from
dark red
14. (at the RBC inlet) to bright red (at the RBC outlet) (scale bar, 3
mm). Boxed
regions are magnified in (F) (scale bar, 1 mm). (G) Perfused
RBCs were
collected at the outlet and quantified for SO2 and PO2. Oxygen
flow increased
SO2 and PO2 of perfused RBCs compared with deoxygenated
RBCs perfused
at the inlet (dashed line) and a nitrogen flow negative control
(N ≥ 3
replicates, data are mean ± SD, *P < 2 × 10−7 by Student’s t
test).
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Fig. 3. Tidal ventilation and oxygenation in hydrogels with
vascularized
15. alveolar model topologies. (A) (Top) Architectural design of an
alveolar
model topology based on a Weaire-Phelan 3D tessellation and
topologic offset
to derive an ensheathing vasculature. (Bottom) Cutaway view
illustrates the
model alveoli (alv.) with a shared airway atrium. Convex (blue)
and concave
(green) regions of the airway are highlighted. (B) Photograph of
a printed
hydrogel during RBC perfusion while the air sac was ventilated
with O2 (scale
bar, 1 mm). (C) Upon airway inflation with oxygen, concave
regions of the
airway (dashed black circles) squeeze adjacent blood vessels
and cause RBC
clearance (scale bar, 500 mm). (D) A computational model of
airway inflation
demonstrates increased displacement at concave regions (dashed
yellow
circles). (E) Oxygen saturation of RBCs increased with
decreasing RBC flow
rate (N = 3, data aremean ± SD, *P < 9 × 10−4 by Student’s t
test).The dashed
line indicates SO2 of deoxygenated RBCs perfused at the inlet.
(F) Elaboration
of a lung-mimetic design through generative growth of the
airway, offset
growth of opposing inlet and outlet vascular networks, and
population of branch
tips with a distal lung subunit. (G) The distal lung subunit is
composed of a
concave and convex airway ensheathed in vasculature by 3D
offset and
anisotropicVoronoi tessellation. (H) Photograph of a printed
16. hydrogel containing
the distal lung subunit during RBC perfusion while the air sac
was ventilated
with O2 (scale bar, 1 mm). (I) Threshold view of the area
enclosed by the dashed
box in (H) demonstrates bidirectional RBC flow during
ventilation. (J) Distal lung
subunit can stably withstand ventilation for more than 10,000
cycles (24 kPa,
0.5 Hz) and demonstrates RBC sensitivity to ventilation gas (N2
or O2).
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Fig. 4. Engraftment of functional hepatic hydrogel carriers. (A
to C)
Albumin promoter activity was enhanced in hydrogel carriers
17. containing
hepatic aggregates after implantation in nude mice. Data from
all time points
for each condition are shown in (B) [N = 4, *P < 0.05 by two-
way analysis of
variance (ANOVA) followed by Tukey’s post-hoc test].
Cumulative bio-
luminescence for each condition is shown in (C) (N = 4, *P <
0.05 by one-way
ANOVA followed by Tukey’s post-hoc test). Error bars indicate
SEM. GelMA,
gelatin methacrylate. (D) Gross images of hydrogels upon
resection (scale
bars, 5 mm). (E) (Left) Prevascularized hepatic hydrogel
carriers are created
by seeding endothelial cells (HUVECs) in the vascular network
after printing.
(Right) Confocal microscopy observations show that hydrogel
anchors
physically entrap fibrin gel containing the hepatocyte
aggregates (Hep)
(scale bar, 1 mm). (F) Hepatocytes in prevascularized hepatic
hydrogel
carriers exhibit albumin promoter activity after implantation in
mice with
chronic liver injury.Graft sections
stainedwithH&Eshowpositioning of hepatic
aggregates (black arrows) relative to printed (case, anchor) and
nonprinted
(fibrin) components of the carrier system (scale bar, 50 mm).
(G) Hydrogel
carriers are infiltrated with host blood (gross, H&E). Carriers
contain
aggregates that express the marker cytokeratin-18 (Ck-18) and
are in close
18. proximity to Ter-119–positive RBCs (scale bars, 40 mm).
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direction) and have the new surface serve as the
template on which a vascular skeleton is built.
With this approach, we developed a bioinspired
alveolar model with an ensheathing vasculature
from 3D tessellations of the Weaire-Phelan foam
topology (35) (Fig. 3 and fig. S9). Although the
fundamental units of the Weaire-Phelan foam
are convex polyhedra (fig. S9), 3D tessellations
can produce a surface containing both convex
and concave regions reminiscent of native alveolar
air sacs (30) with a shared airway atrium sup-
porting alveolar buds (Fig. 3A). We extended
the manifold air surface in the normal direc-
tion, removed faces, and ensheathed edges in
a smoothed polygonal mesh to form a highly
branched vascular network (containing 185 vessel
19. segments and 113 fluidic branch points) that
encloses the airway and tracks its curvature
(fig. S9B).
We printed hydrogels (20wt%, 6-kDa PEGDA)
patterned with the alveolar model topology at a
voxel resolution of 5 pl and a print time of 1 hour
(Fig. 3B). Cyclic ventilation of the pooled airway
with humidified oxygen gas (10 kPa, 0.5Hz) led to
noticeable distension and an apparent change
in the curvature of concave airway regions (fig.
S9C). Perfusion of deoxygenated RBCs at the
blood vessel inlet (10 to 100 mm/min) during
cyclic ventilation led to observable compression
and RBC clearance from vessels adjacent to con-
cave airway regions (Fig. 3, B and C). By observing
dilute RBC streams at the early stages of per-
fusion, we also discerned that the cyclic com-
pression of RBC vessels—actuated by the concave
airway regions upon each inflation cycle—acts as
switching valves to redirect fluid streams to
neighboring vessel segments (movie S3). We im-
plemented a simplified 2D computational model
of airway inflation (fig. S9D), which predicts an-
isotropic distension of the airway and compres-
sion of adjacent blood vessels, corresponding to
local curvature (fig. S9E). In addition, analysis
from a 3D computational model supports an-
isotropic distension of the concave regions of the
airway during inflation (Fig. 3D). Despite the
volume of the alveolar model hydrogel (0.8 ml)
being <25% of that of the serpentine-helix model
(3.5 ml), we measured similar oxygenation ef-
ficiencies for the two designs (Fig. 3E). Our data
suggest that branching topology, hydrogel dis-
tension, and redirection of fluid streams during
20. ventilation may boost intravascular mixing and
allow faster volumetric uptake of oxygen by the
well-mixed RBCs. Vascular constriction during
breathing has been previously described as an
important fluid control mechanism in the mam-
malian lung (36), and here we provide a means
to actualize these ideas in completely defined
and biocompatiblematerials andwithin aqueous
environments.
To extend this work toward a coherent ap-
proximation of scalable lung-mimetic design, we
must consolidate the location of the vascular
inlet, vascular outlet, and air duct, such that dis-
tal lung subunits can be populated on the tips
of multiscale branching architecture. Therefore,
within a given computational bounding volume,we
first derive a branching airway (Fig. 3F). Next,
the centerlines of inlet and outlet blood vessel
networks are grown 180° opposite each other
across and topologically offset from the airway,
and the blood vessels traverse down to the tips of
all daughter branches. The final step is to pop-
ulate the tips of each distal lung with an alveolar
unit cell (Fig. 3G andmovie S4) whose ensheath-
ing vasculature (containing 354 vessel segments
and 233 fluidic branch points) itself is an an-
isotropic Voronoi surface tessellation along a
topological offset of its local airway (fig. S9, F
and G).We found that hydrogels (20 wt %, 6-kDa
PEGDA) could withstand more than 10,000 ven-
tilation cycles (at 24 kPa and a frequency of
0.5 Hz) over 6 hours during RBC perfusion and
while switching the inflow gas between humidi-
fied oxygen and humidified nitrogen (Fig. 3, H
21. to J). Color-filtered views of the early stages of
RBC perfusion (Fig. 3I) indicate that ventilation
promotes RBC mixing and bidirectional flows
within selected vessel segments near the mid-
point of the distal lung subunit (movie S4).
We use our custom stereolithography appa-
ratus for tissue engineering (SLATE) to demon-
strate production of tissue constructs containing
mammalian cells (figs. S1, S10, and S11 andmovie
S5). Lung-mimetic architectures can also be pop-
ulatedwith human lung fibroblasts in the bulk of
the interstitial space and human epithelial-like
cells in the airway (fig. S12), which could facilitate
the development of a hydrogel analog of a lab-
on-a-chip lung design (37). Finally, we subjected
primary humanmesenchymal stem cells (hMSCs)
to SLATE fabrication (with mixtures of PEGDA
and gelatinmethacrylate) and show that the cells
within cylindrical fabricated hydrogels remain
viable and can undergo osteogenic differentia-
tion (fig. S13D). In related multiweek perfusion
tissue culture of hMSCs with osteogenic differ-
entiation media, osteogenic marker–positive
hMSCs were visible throughout the gel (fig. S14).
These studies indicate that SLATE fabrication
supports rapid biomanufacturing, can maintain
the viability of mammalian cell lines, supports
the normal function and differentiation of pri-
mary human stem cells, and provides an ex-
perimentally tractablemeans to explore stem cell
differentiation as a function of soluble factor
delivery via vascular perfusion.
We next sought to establish the utility of this
process for fabricating structurally complex and
22. functional tissues for therapeutic transplanta-
tion. In particular, the liver is the largest solid
organ in the human body, carrying out hundreds
of essential tasks in a manner thought to be
dependent on its structural topology. We created
complex structural features in hydrogel within
the expanded design space imparted by SLATE
to assemblemultimaterial liver tissues. Bioprinted
single-cell tissues andbioprintedhydrogel carriers
containing hepatocyte aggregates were fabricated
(Fig. 4, A to C). The albumin promoter activity
of tissue carriers loaded with aggregates was
enhanced by more than a factor of 60 compared
with that of implanted tissues containing single
cells (Fig. 4, B and C). Furthermore, upon gross
examination of tissues after resection, hydrogel
carrier tissues appeared to havemore integration
with host tissue and blood (Fig. 4D). Despite the
improved utility of hepatic aggregates over single
cells, aggregate size puts substantial architectural
limitations on 3D printing because aggregates
are larger in size than our lowest voxel resolution
(50 mm). To accommodate these design con-
straints, we built a more advanced carrier that
can deliver hepatic aggregates within natural
fibrin gel, has a vascular compartment that can
be seeded with endothelial cells, and incorpo-
rates structural hydrogel anchors to physically,
rather than chemically, retain the fibrin gel and
facilitate remodeling between the graft and host
tissue (Fig. 4E and fig. S15). Microchannel net-
works were seeded with human umbilical vein
endothelial cells (HUVECs) because our previous
studies demonstrated that inclusion of endothelial
cords improved tissue engraftment (38). We then
23. evaluated whether optimized bioengineered liver
tissueswould survive transplantation in a rodent
model of chronic liver injury. After 14 days of
engraftment in mice with chronic liver injury,
hepatic hydrogel carriers exhibited albumin pro-
moter activity indicative of surviving functional
hepatocytes (Fig. 4F). Immunohistological char-
acterization revealed the presence of hepatic
aggregates adhered to printed hydrogel com-
ponents that stained positively for the marker
cytokeratin-18 (Fig. 4, F and G). Further charac-
terization through gross examination and higher-
magnification images of slides stained with
hematoxylin and eosin (H&E) indicated the pres-
ence of host blood in explanted tissues. Immu-
nostaining using a monoclonal antibody against
Ter-119 confirmed the erythroid identity of cells in
microvessels adjacent to hepaticmicroaggregates
in explanted tissues (Fig. 4G, right). This work
provides an approach to address long-standing
design limitations in tissue engineering that have
hindered progress of preclinical studies.
We have identified readily available food dyes
that can serve as potent photoabsorbers for bio-
compatible and cytocompatible production of
hydrogels containing functional vascular to-
pologies for studies of fluidmixers, valves, inter-
vascular transport, nutrient delivery, and host
engraftment.With our stereolithographic process,
there is potential for simultaneous and orthog-
onal control over tissue architecture and bio-
materials for the design of regenerative tissues.
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(2016).
17. M. S. Hahn, J. S. Miller, J. L. West, Adv. Mater. 18, 2679–
2684
(2006).
18. C. A. DeForest, K. S. Anseth, Nat. Chem. 3, 925–931
(2011).
19. K. A. Heintz et al., Adv. Healthc. Mater. 5, 2153–2160
(2016).
20. T. M. Fonovich, Drug Chem. Toxicol. 36, 343–352 (2013).
21. S. Kumar, J. Aaron, K. Sokolov, Nat. Protoc. 3, 314–320
(2008).
22. L. J. Stevens, J. R. Burgess, M. A. Stochelski, T. Kuczek,
Clin. Pediatr. 54, 309–321 (2015).
23. M. Li et al., J. Agric. Food Chem. 62, 12052–12060 (2014).
24. C. A. DeForest, B. D. Polizzotti, K. S. Anseth, Nat. Mater.
8,
659–664 (2009).
25. …
26. 3 1 2 VOLUME 34 NUMBER 3 MARCH 2016 nature
biotechnology
A r t i c l e s
The demand for engineered tissues has risen rapidly owing to
the
limited availability of donor tissues and organs for
transplantation.
Despite some initial successes in engineering relatively simple
tissues,
many challenges remain in developing tissues and organs
suitable for
clinical translation1,2. Three-dimensional (3D) printing
technology
shows promise for creating complex composite tissue
constructs3–8
through precise placement of cell-laden hydrogels in a layer-by-
layer
fashion7,9–17. The most commonly used bioprinting systems
are based
on jetting, extrusion and laser-induced forward transfer
(LIFT)6,18,19.
The jetting method produces picoliter scale drops with a
printing
resolution of 20~100 µm. However, because the hydrogel
concen-
tration is low20–23, the thickness of printed constructs may be
lim-
ited because of inadequate structural support24. Extrusion
methods,
which use a syringe and piston system to dispense material
through
microscale nozzles, can produce more stable 3D cell-laden
structures
using high concentrations of hydrogels such as alginate, fibrin
27. and
Pluronic F-127 (refs. 18,25–27). However, it is difficult to
construct
large free-form tissue structures owing to inadequate structural
integ-
rity, mechanical stability and printability28–30. The LIFT
method can
precisely print cells in relatively small constructs31 but requires
rapid
gelation of hydrogels to achieve high resolution of the printed
pat-
terns, resulting in low flow rates.
Here we describe a system that deposits cell-laden hydrogels
together
with synthetic biodegradable polymers that impart mechanical
strength, thereby overcoming previous limitations on the size,
shape,
structural integrity and vascularization of bioprinted tissue con-
structs. This was accomplished by designing multidispensing
modules
for delivering various cell types and polymers in a single
construct;
by developing an optimized carrier material for delivering cells
to
discrete locations in the 3D structure in a liquid form; by
designing
sophisticated nozzle systems with a resolution down to 2 µm for
bio-
materials and down to 50 µm for cells; by cross-linking cell-
laden
hydrogels after passage though the nozzle system; by
simultaneously
printing an outer sacrificial acellular hydrogel mold that is
dissolved
28. after the tissue construct acquires enough rigidity to retain its
shape;
and by creating a lattice of microchannels permissive to nutrient
and
oxygen diffusion into the printed tissue constructs. These
properties,
all designed to work in a coordinated manner, make up the
ITOP. We
demonstrate the printer by fabricating human-scale mandible
bone,
ear-shaped cartilage and organized skeletal muscle. Evaluation
of
the characteristics and function of these tissues in vitro and in
vivo
showed tissue maturation and organization that may be
sufficient for
translation to patients.
RESULTS
Design of the ITOP system
Multiple cartridges (Fig. 1a and Supplementary Fig. 1) are used
to
deliver and pattern multiple cell-laden composite hydrogels,
support-
ing poly(ε-caprolactone) (PCL) polymer and a sacrificial
Pluronic
F-127 hydrogel (Fig. 1b). The end of each cartridge is
connected
to a microscale nozzle, and the top is connected to an air
pressure
controller for precisely controlling the dispensing volume. A
heating
unit ensures that the PCL remains easily dispensable. A three-
axis
motorized stage system enables 3D patterning of multiple cells
and
29. biomaterials. The system resides in a humidified and
temperature-
controlled (18 °C) enclosure.
The correct shape of a tissue construct is obtained from a human
body by processing computed tomography (CT) or magnetic
resonance
imaging (MRI) data in computer-aided design (CAD) software
(Fig. 1c).
A custom nozzle motion program is generated by incorporating
A 3D bioprinting system to produce human-scale
tissue constructs with structural integrity
Hyun-Wook Kang, Sang Jin Lee, In Kap Ko, Carlos Kengla,
James J Yoo & Anthony Atala
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 mm for
cell survival in engineered tissues. We demonstrate capabilities
of the ITOP by fabricating mandible and calvarial bone,
cartilage
32. the main computer of the 3D printing system
that effects the biofabrication process.
The composite hydrogel for cell delivery
consisted of gelatin, fibrinogen, hyaluronic
acid (HA) and glycerol mixed into DMEM (high glucose). We
tested
various concentrations of each component to achieve proper
print-
ing resolution and dispensing uniformity, mechanical properties
(before and after cross-linking with thrombin) and cell viability
(Supplementary Fig. 2). The optimized concentrations of hydro-
gel ingredients and the numbers of cells needed for fabrication
of
individual tissue constructs are listed in Table 1.
Patterning synthetic polymers confers mechanical strength
Our first series of studies tested the ability of the ITOP to
generate uni-
form two-dimensional (2D) and 3D cell patterns of multiple cell
types.
Using 3T3 fibroblasts labeled with two fluorescent dyes, Dil
(red) and
DiO (green), we demonstrated delivery of the two populations
in a
unique 2D pattern (Fig. 2a,b). To create 3D constructs, we
combined
the fluorescently labeled 3T3 fibroblasts in composite hydrogels
with
supporting PCL and printed them in two patterns—type I (Fig.
2c)
and type II (Fig. 2f). These patterns differ in the placement of
PCL and
thus in the mechanical strength of the printed construct. The
type I
pattern creates multiple PCL frames in each layer throughout
33. the con-
struct, and places cells and gel materials in between the frames.
The
type II pattern consists of cell-laden hydrogel and porous
structures,
surrounded by a PCL framework on the outer layers and corners
of
each layer, thus protecting the contents from external load.
Type I
constructs (Fig. 2d,e) maintained a more stable structure than
type II
constructs (Fig. 2g,h), owing to the abundance of uniformly
distrib-
uted PCL frames. Therefore, we used the type I pattern to
fabricate
mandible bone and ear-shaped cartilage structures and the type
II
pattern to print organized skeletal muscle constructs.
Next, we produced 3D structures by placing either type I or type
II pat-
terns of cell-laden hydrogel and PCL (~130 µm for type I and
~250 µm
wide for type II). The microchannels (type I: 500 × 300 µm2;
type II:
650 × 450 µm2), formed by the PCL patterns, were designed to
maxi-
mize diffusion of nutrients and oxygen. In addition, we used
Pluronic
F-127 hydrogel as a sacrificial outer layer to support the 3D
architecture
of the dispensed cell-laden structures before crosslinking. After
cross-
linking of fibrinogen using thrombin, the uncross-linked
components
34. (gelatin, HA, glycerol and Pluronic F-127) were washed out.
To determine cell viability during printing, we examined
survival of
3T3 fibroblasts at 60 min (day 0), 3 d and 6 d after printing.
Live/dead
cell assays showed ≥95% cell viability on day 0, which was
maintained
through days 3 and 6 (Fig. 2i). Cell proliferation, assessed using
the
AlamarBlue assay system, increased over a 15-d period, similar
to the
proliferation of control cells encapsulated in a fibrin construct
(Fig. 2j).
These data indicate that the optimized composite hydrogel
system
maintained cell viability during the printing process and
provided a
favorable microenvironment for cell proliferation.
Mandible bone reconstruction
To demonstrate construction of a human-sized bone structure,
we fab-
ricated a mandible fragment in a size and shape similar to what
would
be needed for facial reconstruction after traumatic injury (Fig.
3).
The cell type used was human amniotic fluid–derived stem cells
Main
computer
3-axis stage
controller
Pressure
35. controller
Multi-
cartridge
module
3D
printed
construct
PCL (gray)
A: cell A (red)
B: cell B (green)
S: sacrificial material
Closed chamber
Medical imaging
(CT, MRI)
Visualized motion
program
3D printing process
3D bioprinted
tissue product
10 mm
DICOM format STL format Text-based
command list
3D printed
construct
PCL
36. Cell A
Cell B
XYZ
stage
Heating
unit
PCL
A B S
Po
re
(m
icr
oc
ha
nn
el)
3D CAD model
a b
c
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
39. biotechnology
A r t i c l e s
(hAFSCs), which can give rise to osteogenic lineages in
appropriate
media32,33. Mandible bone defects have an arbitrary shape. We
used
data from a CT scan of a human mandible defect in combination
with Mimics software (Materialise, Leuven, Belgium) to
produce a
CAD model of the defect shape, with dimensions of 3.6 cm ×
3.0 cm
× 1.6 cm (Fig. 3a). A text-based command motion program,
gener-
ated from the CAD model with custom CAM software,
determined
the required dispensing paths of cell-laden hydrogel, a mixture
of
PCL and tricalcium phosphate (TCP), and Pluronic F127 (Fig.
3b).
PCL/TCP and hAFSCs mixed with the composite hydrogel
(Table 1)
were printed in a type I pattern with a Pluronic F127 temporary
sup-
port (Fig. 3c). At 1 d of culture, cell viability in the printed
bone
structures was 91 ± 2% (n = 3, Table 1), confirming that the
printing
process did not adversely affect cell viability. After induction
of osteo-
genic differentiation using an established protocol32,33 for 28 d
(n = 5,
Fig. 3d), we stained the structures with Alizarin Red S; staining
at the
40. surface of the 3D bone structures indicated calcium deposition
in the
hAFSC-laden hydrogel (Fig. 3e). 3D constructs before
differentiation
showed no Alizarin Red S staining (data not shown).
Calvarial bone reconstruction
To study maturation of the bioprinted bone in vivo, we
fabricated rat
calvarial bone constructs in a circular shape (8 mm diameter ×
1.2 mm
thickness) with hAFSCs (Fig. 4a,b and Supplementary Fig. 3),
cultured
them in osteogenic media for 10 d, implanted
them in a calvarial bone defect region of
Sprague Dawley rats (n = 4) and analyzed
them 5 months after implantation (Fig. 4c).
Type l
Pore
1 mm
1 mm
1 mm
1 mm
Type l Top Type l - top
Type ll
PCL
43. in
te
n
si
ty
Control
Printed
500 µm
1 3 7 15
a b
c d e
f g h
i
j
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. (c–h) Two basic
types of 3D patterning: type I pattern (c–e) and type II pattern
(f–h). 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
44. 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.
Bony
defect
Red: cells
Green: PCL
Blue: Pluronic F-127
Pluronic
F-127 (sacrificial
material)
Printing nozzle
Pore
PCL
Cell-laden
hydrogel
30 mm
3
6
m
m
P
45. o
re
PCL
Cell-laden
hydrogel
1 mm
a b c
d e
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.
np
g
47. d.
nature biotechnology VOLUME 34 NUMBER 3 MARCH
2016 3 1 5
A r t i c l e s
The bioprinted constructs showed newly
formed vascularized bone tissue through-
out the implants, including the central por-
tion, with no necrosis (Fig. 4j), whereas
the untreated defect and scaffold-only
treated control groups showed fibrotic tis-
sue ingrowth (Fig. 4d) and minimal bone
tissue formation restricted to the periphery
of the implant (Fig. 4g), respectively. The
modified Tetrachrome staining confirmed
mature bone (red) and osteoid (blue) for-
mation (Fig. 4e,h,k). Von Willebrand factor
(vWF) immunostaining showed large blood
vessel formation within newly formed bone
tissue throughout the bioprinted bone con-
structs, including the central portion (Fig. 4l), whereas the
nontreated
(Fig. 4f) and scaffold-only (Fig. 4i) groups had only limited
vascu-
larization restricted to the periphery of the implant.
Ear cartilage reconstruction
Next, we tested the ability of the ITOP to fabricate tissue
constructs
of complex shape by making human-sized external ears, as the
frame-
work of an auricle consists of a single piece of cartilage with a
48. com-
plicated geometry of ridges. A CT image of an ear (Fig. 5a) was
used
to develop a motion program (Fig. 5b) to print a chondrocyte-
laden
hydrogel, PCL and Pluronic F-127. Using rabbit ear
chondrocytes
(passages 3 and 4) mixed with the composite hydrogel (Table
1), we
fabricated human ear–shaped cartilage constructs with
dimensions of
3.2 cm × 1.6 cm × 0.9 cm (Fig. 5c–e) in the type I pattern. Cell
viabil-
ity was 91 ± 8% at 1 d after printing (n = 3, Table 1). After 5
weeks
in the culture medium, the constructs were stained with
Safranin-O
and showed production of a new cartilaginous matrix (Fig. 5f).
The constructs with microchannels showed enhanced tissue
forma-
tion as evidenced by the production of new viable cartilaginous
matrix
throughout the entire ear constructs. In contrast, the constructs
without microchannels showed only limited tissue formation
restricted to the peripheral region, likely owing to the diffusion
limits of nutrients and oxygen. The cells in the newly formed
tissues
demonstrated similar morphological characteristics to those in
native
ear cartilage, with cells located within typical chondrocyte
lacunae,
surrounded by a cartilaginous matrix (Fig. 5g). Native human
ear
tissue served as a positive control.
To determine whether the printed ear constructs would mature
49. in vivo,
we implanted them in the dorsal subcutaneous space of athymic
mice
and retrieved them 1 and 2 months after implantation (n = 4).
The
shape was well maintained, with substantial cartilage formation
upon
gross examination (Fig. 5h). Histological analysis showed the
for-
mation of cartilage tissue (Fig. 5i). The glycosaminoglycan
(GAG)
content (2.7 ± 0.2 µg/mg at 1 month and 4.2 ± 0.3 µg/mg at 2
months)
increased over time, reaching 20% of that of native ear GAG
content
(Fig. 5j). Vascularization of the printed constructs in the outer
region
was suggested by endothelial cell marker expression at 1 and 2
months
after implantation (Supplementary Fig. 4). The inner regions
were
avascular (Supplementary Fig. 4), as in native cartilage, but the
car-
tilage cells were viable, suggesting adequate nutrient diffusion
during
development. Biomechanical analyses (n = 4, Fig. 5k) showed
that
maturation in vivo strengthened the tissue constructs, resulting
in a
higher normalized load during bending compared with pre-
implant
constructs. In addition, resilience, measured by the ∆Load%,
was tested
by repeated bending and relaxation cycles. Resilience between
the
repeated bending cycles was much higher in the constructs
50. implanted
for 1 month (Fig. 5m and Supplementary Table 1) than in the
con-
structs before implantation (Fig. 5l) . These results demonstrate
the
generation of ear-shaped cartilage with resilience properties
similar to
those of native cartilage (rabbit ear) (Supplementary Table 1).
Skeletal muscle reconstruction
Finally, we applied the ITOP to fabricate an organized soft
tissue—a
3D muscle construct 15 mm × 5 mm × 1 mm in dimension
containing
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. (d–l) Histological and
immunohistological images of nontreated (d–f),
scaffold only without cells (g–i) and hAFSCs-printed
construct at 5 months after implantation (j–l).
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.
51. Red: cells
Green: PCL/TCP
200 µm2 mm
Top view
530 µm
Side view
Day 0
5 months
5 mm
NB
NB
PCL/
TCP
PCL/
TCP
PCL/
TCP
PCL/
TCP
NB
NB
54. A r t i c l e s
mouse myoblasts (Table 1) printed in the type II pattern (Fig.
6a,b).
Immediately after printing, the printed structures contained
mus-
cle fiber–like bundles (~400 µm width), supporting PCL pillars
and Pluronic F-127 hydrogel as a temporary structure (Fig. 6c
and
Supplementary Fig. 5a). Notably, the printed cells began
stretch-
ing along the longitudinal axis of the constructs at day 3 in
growth
media (Fig. 6e and Supplementary Fig. 5b) with high cell
viability
(Fig. 6f), and the constructs underwent compaction34, keeping
the
fibers taut during cell growth and differentiation, whereas the
printed
cells without PCL support did not show cellular alignment (Fig.
6d).
After 7 d in differentiation media, muscle-like structures with
aligned
myotubes were observed (Fig. 6g and Supplementary Fig. 5c).
To study whether these structures could mature into functional
muscle in vivo, we implanted 7-d differentiated structures
subcutane-
ously (ectopically) in 14- to 16-week-old nude rats (n = 6). The
dis-
sected distal end of the proximal stump of the common peroneal
nerve
(CPN) was embedded within the constructs to promote
integration
(Fig. 6h,i). Adequate innervation of implanted muscle is
55. essential to
achieve and maintain muscle function. Our model allowed us to
eval-
uate nerve integration of the implanted muscle construct
independ-
ent of the surrounding muscle tissue. After 2 weeks of
implantation,
the retrieved muscle constructs showed well-organized muscle
fiber
structures (Fig. 6j), the presence of acetylcholine receptor
(AChR)
clusters on the muscle fibers (MHC+ and α-BTX+) (Fig. 6k), as
well
as nerve (neurofilament) contacts with α-BTX+ structures
within the
implants (Fig. 6l), indicating that the printed muscle constructs
were
robust enough to maintain their structural characteristics and
induce
nerve integration in vivo. In addition, vascularization
throughout the
muscle constructs was indicated by endothelial cell marker
expression
(Fig. 6m). To examine muscle function, we performed
electromyogra-
phy to evaluate electrical and neurological activation of the
constructs
2 weeks after implantation. Compound muscle action potential,
which
is evoked by motor nerves and measures muscle function, was
3.6 mV,
compared to 10.7 mV for the control gastrocnemius muscle, and
Red: cells
Green: PCL
Blue: Pluronic F-127
57. 500 µm 500 µm
100 µm 100 µm
500 µm
100 µm
100 µm100 µm100 µm
PCL
Microchannel
10 mm 16 mm
3
2
m
m
Pluronic
F-127
Printed ear
construct
PCL
PCL
a b c d e f
g
Pre-implantation
62. 0
0 0.5 1.0 1.5 2.0
Extension (mm)
0 0.5 1.0 1.5 2.0
Extension (mm)
Cycle 1
Cycle 2
Cycle 3
Cycle 4
h i
j
k l m
500 µm 500 µm
Alcian Blue
Figure 5 Ear cartilage reconstruction. (a–f) 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
63. 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. (h–m) 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
65. gh
ts
r
es
er
ve
d.
nature biotechnology VOLUME 34 NUMBER 3 MARCH
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A r t i c l e s
0 mV for the negative controls (subcutaneous tissue), indicating
that
the implanted muscle constructs responded to electrical
stimulation
to an extent consistent with immature, developing muscle (Fig.
6n).
DIScUSSIOn
Bioprinters based on jetting, extrusion and LIFT methods can
deliver
viable cells, biomaterials and macromolecules to generate 3D
tissue
structures. However, in general they are limited in their ability
to gen-
erate large biological constructs with sufficient structural
integrity for
surgical implantation28–30, and the few in vivo studies of
66. bioprinted
tissue structures tested less complex constructs with low
mechanical
stability35,36. The ITOP can address the limitations of size and
sta-
bility by sequentially printing cell-laden hydrogels with a
synthetic
polymer and a temporary scaffolding, creating tissue constructs
with
the structural integrity needed for surgical implantation. A
computer-
generated 3D tissue model can be converted to a motion
program that
operates and guides the dispensing nozzles to take defined paths
for
delivery of cells and materials. The cell-laden hydrogel protects
cell
viability and promotes growth and expansion, whereas the
adjacent
sacrificial scaffolding provides the initial structural and
architectural
integrity. As the cells anchored three dimensionally within the
hydro-
gel initiate the transition to tissue formation, they start to
secrete their
own matrix, replacing the hydrogel as it slowly degrades over
time.
The system’s modular design enables printing of a wide array of
tissue
constructs. Here we used up to four material repositories, but
many
additional repositories could be installed to print constructs
contain-
ing multiple cell types and biomaterials.
67. Cell carriers for bioprinting must provide adequate mechanical
support, cell-specific cues and negligible cytotoxicity. As few
such
materials are available37,38, we fulfilled these requirements
with a
mixture of gelatin, fibrinogen, HA and glycerol. …
R E V I E W
T I S S UE E N G I N EE R I N G
Engineering Complex Tissues
Anthony Atala,1 F. Kurtis Kasper,2 Antonios G. Mikos2*
D
Tissue engineering has emerged at the intersection of numerous
disciplines to meet a global clinical need for
technologies to promote the regeneration of functional living
tissues and organs. The complexity of many tissues
and organs, coupled with confounding factors that may be
associated with the injury or disease underlying the
need for repair, is a challenge to traditional engineering
approaches. Biomaterials, cells, and other factors are
needed to design these constructs, but not all tissues are created
equal. Flat tissues (skin); tubular structures
(urethra); hollow, nontubular, viscus organs (vagina); and
complex solid organs (liver) all present unique chal-
lenges in tissue engineering. This review highlights advances in
tissue engineering technologies to enable
regeneration of complex tissues and organs and to discuss how
such innovative, engineered tissues can affect
the clinic.
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lo
69. ://stm
.scie
n
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a
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.o
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fro
m
INTRODUCTION
A tremendous clinical need exists for the development of
technologies
to facilitate the regeneration of injured or diseased tissues and
organs.
The unrelenting prevalence of trauma, congenital defects, and
diseases
such as cancer drives the demand, which becomes increasingly
urgent
as the global population expands and ages. A wide variety of
tissues
and organs would benefit from engineering-based repair or
70. regenera-
tion, from musculoskeletal tissues, such as bone and cartilage,
to entire
organs, including the bladder and liver. The field of tissue
engineering
is at the interface of bioengineering, materials science,
chemistry, biol-
ogy, and medicine, poised to meet these unmet clinical needs
through
the development of new technologies and refinement of existing
ones.
Increasing levels of complexity in the tissues or organs targeted
for
repair generally necessitate a concomitant increase in the
complexity
of the associated tissue engineering approach. For example,
solid
organs, such as the kidney, would require several essential
structures
to restore function, whereas tubular hollow organs, such as the
urethra,
are more easily recreated from basic cells and materials (1).
Similarly,
complexity can be found at the interfaces between tissues, such
as the
transition from cartilage to bone in the osteochondral interface
in ar-
ticulating joints. Such interfaces are receiving increasing
attention as
targets for repair, given the prevalence of injuries affecting
them (2, 3).
Indeed, a complex tissue injury or defect may involve multiple
tissue
types, may be associated with compromised vascularity, or may
be at
71. risk for infection.
Regardless of the complexity of the target for repair, tissue
engi-
neering strategies generally involve the application of
combinations
of biomaterials, cells, and biologically active factors to effect
tissue for-
mation. This process can involve de novo growth in tissue
culture (in
vitro, ex vivo) or induction of tissue regeneration in vivo at
sites or
under conditions where it otherwise would not occur. Increasing
em-
phasis is being placed on the development of tissue engineering
ap-
proaches within the context of the injury or disease underlying
the
defect. For example, traumatic injuries to the extremities may
involve
multiple tissue types (bone, muscle, vasculature, lymphatics,
nerve),
and biomaterial-based approaches for regeneration are being
devel-
1Wake Forest Institute for Regenerative Medicine, Wake Forest
School of Medicine,
Winston Salem, NC 27157, USA. 2Department of
Bioengineering, Rice University, Houston,
TX 77251, USA.
*To whom correspondence should be addressed. E-mail:
[email protected]
www.ScienceT
oped and evaluated using preclinical composite tissue defect
models
(4). The present review will focus on advances in tissue
engineering
72. and regenerative medicine that may enable the repair of tissues
with
high complexity, while highlighting bottlenecks to clinical
translation
of these technologies.
TISSUE ENGINEERING SCAFFOLDS
Biomaterials can provide a three-dimensional (3D) structure to
sup-
port tissue growth. These scaffolds define and maintain the
space in
which the target tissue will form and can be tailored to support
the
attachment and proliferation of cells to effect the desired tissue
forma-
tion (5). Ideally, a scaffold should serve as a transient structure
that
will degrade or resorb with time, such that it is replaced with
the tissue
of interest. Advances in biomaterials science combined with
increas-
ing knowledge of extracellular matrix (ECM) biology and the
role of
environmental factors in tissue formation have led to the
development
of scaffolds tailored to provide appropriate structural support
and, in
some cases, biological and mechanical cues to promote tissue
regen-
eration in vivo (6–9). Moreover, scaffold biomaterials can be
modified
to present biologically active signals, including cell-adhesion
peptides
and growth factors, to facilitate cell attachment and to direct
tissue
formation (10–12). In some instances, the scaffolds depend
73. entirely
on the migration of cells from the body into the defect for tissue
for-
mation to occur, whereas other approaches leverage the
scaffolds for
the transplantation of cell populations to supplement the body.
In either
case, tissue engineering scaffolds seek to mimic key elements of
the
ECM and local microenvironment to support and perhaps induce
tis-
sue formation.
Naturally derived polymeric materials, including polypeptides
(for
example, collagen) and polysaccharides (for example,
hyaluronic acid),
have been explored extensively in the development of tissue
engineering
scaffolds for applications ranging from cartilage repair to
functional
pancreatic replacements (13). Indeed, a key advantage
associated with
naturally derived polymers is the general capacity of these
materials
to support the attachment, proliferation, and differentiation of
cells
(14, 15). Although naturally derived polymers are typically
enzymati-
cally degradable, the kinetics of degradation may not be easily
con-
trolled or predicted. The generally weak mechanical strength
associated
with naturally derived polymers is also a limitation, but it may
be
ranslationalMedicine.org 14 November 2012 Vol 4 Issue 160
76. improved through the introduction of intermolecular cross-links
(16).
However, cross-linking may prolong the degradation of the
materials
(17). A concern with naturally derived polymeric materials is
the var-
iability inherent in the production of the materials as well as the
po-
tential, albeit small, of the materials to evoke an immune
response.
Synthetic polymers present several key advantages relative to
nat-
urally derived polymers. Synthetic polymers can be
reproducibly manu-
factured with a wide range of mechanical properties and
degradation
kinetics to enable the production of scaffolds with properties
tailored
for a particular application (18). For example, scaffolds
comprising
poly(lactic-co-glycolic acid) (PLGA) have been investigated for
the re-
generation of tissues ranging from blood vessels to bone. Many
syn-
thetic polymers undergo hydrolytic degradation, which may be
more
readily predicted and controlled than enzymatic degradation in
vivo,
given the lack of dependence on local enzyme concentrations.
Certain
classes of synthetic polymers, such as poly(a-hydroxy esters),
produce
acidic products upon degradation (19), which may elicit a
prolonged
77. inflammatory response (20). Nevertheless, synthetic polymers
them-
selves typically do not carry a risk for inducing an immune
response
owing to a lack of biologically functional domains. This is also
a lim-
itation because synthetic polymers cannot facilitate cell
attachment or
direct phenotypic expression as a natural polymer would.
However, a
variety of synthesis techniques have been developed to
incorporate bi-
ologically active domains into synthetic polymer scaffolds,
thereby
enabling the production of biomimetic scaffolds with a defined
and
tunable composition (21).
Materials derived from the native ECM have also been explored
as
scaffolds. Tissues like the urinary bladder submucosa or the
small in-
testinal submucosa can be processed through mechanical and
chemical
manipulation to remove the cellular components, yielding a
collagen-
rich matrix, in a process called “decellularization.” The
structures and
arrangement of the various ECM proteins in the resulting
acellular
matrices are largely conserved, which results in a general
retention
of the mechanical properties of the original tissue (22).
Moreover, acel-
lular tissue matrices have been shown to support the ingrowth
of cells
78. and tissues in several applications, without inducing a gross
immune
response (23, 24). Indeed, given the natural origin of the
matrices, the
materials degrade slowly after implantation and are replaced or
re-
modeled with matrix produced by cells (25). Decellularized
matrices
may also be processed to form particulates that can be used
either
alone or in combination with other materials to promote tissue
re-
pair (26).
In other cases, synthetic polymeric scaffolds have been
fabricated
and modified through covalent immobilization of ECM-derived
moi-
eties to control presentation of growth factors, promote cell
attachment,
and enhance directed differentiation of progenitor cell
populations (27).
Additional methods to introduce an ECM-mimetic coating on
scaf-
folds have been explored, including coating of synthetic
polymeric scaf-
folds with naturally derived polymers (collagen or gelatin) and
ceramics
(calcium phosphate) for bone tissue engineering applications
(28, 29).
Loai et al. (30) combined particles of acellular tissue matrices
(urinary
bladder submucosa) with polymeric materials in the fabrication
of
scaffolds with biological activity and tunable properties for
generation
79. of a vascularized bladder in murine and porcine preclinical
models.
Alternative approaches have seeded cell populations onto
scaffolds
and leveraged culture conditions to drive the differentiation of
cells
and the concomitant production of ECM. Recently, this was
demon-
strated in the production of bone-like ECM (31, 32).
www.ScienceT
To support tissue formation, a tissue engineering scaffold must
present an interconnected porosity or be capable of resorbing as
a func-
tion of time to create space for new tissues. Many fabrication
techniques
have been developed to enable the fabrication of 3D scaffolds
with an
interconnected porosity, ranging from particulate leaching
techniques
to electrospinning methods (33, 34). Although traditional
methods for
scaffold fabrication can enable introduction of interconnected
pores
with a tunable pore size, control of pore architecture has been a
chal-
lenge (35). Three-dimensional printing methods have emerged
to en-
able the fabrication of scaffolds with precise control of the
architecture
throughout the structure (36, 37). Printing techniques have even
been
used to produce scaffolds with controlled gradients in
mechanical prop-
erties and gradients of biologically active factors (38). With this
technol-
ogy, scaffolds with spatially controlled properties have been
80. created for
the regeneration of complex tissue structures, such as bone and
cartilage
(39–41). However, in some cases, printing of complex scaffolds
of di-
mensions of clinical relevance, such as whole kidneys or livers,
may be
too time-consuming for widespread application.
Tissue engineering scaffolds should support the attachment and
proliferation of cells and the subsequent formation of the tissue
of in-
terest. However, scaffold materials alone often lack the
biological cues
to induce tissue formation. Accordingly, scaffolds are
commonly used
for the presentation or controlled delivery of biologically active
factors
to induce tissue regeneration. Growth factors, ranging from
angiogenic
factors, such as vascular endothelial growth factor, to
osteogenic fac-
tors, such as bone morphogenetic protein-2, have been
incorporated
into scaffolds to promote tissue formation (42, 43). Key
challenges as-
sociated with growth factor delivery in tissue engineering
include not
only selection of the appropriate factor or combination of
factors neces-
sary to induce the desired response but also the dose and
spatiotemporal
delivery needed for proper tissue development (44–46). Another
chal-
lenge has been the maintenance of the biological activity of the
factor,
81. especially once released from the scaffold.
CELLS IN ENGINEERED COMPLEX TISSUES
Scaffolds used in tissue engineering approaches are commonly
divided
into two general categories, namely, acellular scaffolds, which
depend
on cells in the recipient to effect tissue formation, and cellular
scaf-
folds, which serve as cell transplantation vehicles. In both
cases, the
success of a scaffold technology toward achieving tissue growth
de-
pends largely on the action of the cells. Accordingly, many
current ef-
forts in tissue engineering seek to identify and optimize cell
populations
that can be leveraged for delivery with a scaffold to promote
tissue
repair where it otherwise might not occur.
Autologous cell populations have been of great interest for
appli-
cation in tissue engineering approaches because they have
minimal
risk of rejection. Some early efforts in the field focused on
isolating
primary cells from a biopsy of the tissue or organ of interest
and grow-
ing the cells ex vivo for subsequent introduction back into the
patient
in a tissue engineering therapy. However, a major limitation
encountered
in this area has been the difficulty in expanding cells to
sufficient num-
bers for clinical application. As an alternative, precursor cells
82. and their
necessary culture conditions for tissue engineering have been
identified
for several tissues and organs. For instance, urothelial cells
have been
grown and expanded in vivo, but traditionally, the expansion
has
been limited (47, 48). Methods have been developed in recent
years to
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85. phase
to obtain sufficient numbers of cells for seeding of scaffolds
(47, 49).
These methods have enabled the isolation of urothelial cells
from a
single specimen with dimensions of 1 cm2 and the expansion of
these
cells over a period of 8 weeks to sufficient numbers to cover the
equivalent of a football field (4202 m2) (47).
Although advances in cell culture protocols have allowed for
ex-
pansion of autologous cells to sufficient numbers for clinical
applica-
tion, expansion of primary cells from some tissues and organs,
such as
the pancreas, remains a challenge. Additionally, in some cases,
tissue
engineering strategies rely on autologous cells derived from
diseased
tissues or organs, which may not yield a sufficient number of
normal
cells for clinical application. As a result, tissue engineers seek
to leverage
autologous stem and progenitor cell populations, such as bone
marrow–
derived mesenchymal stem cells (MSCs) and adipose-derived
stem
cells (50). Although MSCs have received a great deal of
attention in
the tissue engineering literature, advances with other adult-
derived stem
cell populations have generally progressed slowly, owing in part
to diffi-
culties associated with maintaining the stem cells in culture or
achiev-
86. ing attachment of the cells to scaffolds (51). Nevertheless, some
clinical
strategies have involved seeding of patient-derived stem and
progen-
itor cells on biomaterial scaffolds and then leveraging the body
as a
bioreactor for tissue growth. For example, a ceramic scaffold
within
a titanium mesh was seeded with bone marrow as a source of
stem
cells and implanted in the latissimus dorsi of a patient to grow a
man-
dibular replacement ectopically (52).
Other types of stem cells have been included in biomaterial
scaf-
folds for tissue engineering applications. These cell-based
therapies are
beyond the scope of this review, but the reader is referred to
(53–55).
It should be noted that a vascular network is generally needed to
sup-
port the viability of cells throughout a larger, more complex
tissue-
engineered construct. Accordingly, a variety of methods have
been
developed to promote vascularization of tissue-engineered
constructs,
ranging from functionalization of scaffolds with bioactive
factors to
www.ScienceT
development of bioreactor systems to promote vessel formation
ex vivo
(56–58). A detailed discussion of vascularization strategies is
provided
in (59, 60).
87. CREATING COMPLEX ORGANS
An expansive toolbox of biomaterial- and cell-based
technologies stands
ready to contribute to the production of tissue engineering
solutions
to meet clinical needs. However, immense complexity can be
found
in the various tissues and organs targeted for replacement.
More-
over, the injury or disease driving the need for tissue repair or
replace-
ment can add levels of complexity. A common challenge
encountered
in the development of tissue engineering technologies is the
need to
repair tissue defects or to regenerate organs that have intricate
3D
structures. Furthermore, it is challenging to integrate the
regenerating
tissue with surrounding tissues and to maintain cell viability in
large
constructs.
To better understand the structural design of human tissues and
organs that regenerative medicine attempts to replicate, it may
be
helpful to categorize them into four levels according to their
increasing
complexity: flat tissue structures; tubular structures; hollow,
nontubu-
lar, viscus structures; and complex solid organs (Fig. 1). Within
these
levels of complexity, there are several strategies used to achieve
resto-
ration of function. We also consider the unmet clinical needs in
88. these
areas and the barriers to translation in existing demonstrations.
Flat structures
Sheets of cells consisting of multiple layers of predominantly
one cell
type represent the simplest architectural subtype in the body.
This
level of tissue complexity is exemplified by the integument
system,
which represents one of the earliest attempts at culturing
autologous
cells in vitro for repair purposes (61). The effects of substantial
loss
of skin surface area are detrimental, as can be seen in burn
patients.
Traditional treatments, such as skin grafts harvested from
unburned
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Kidney
Bladder
89. Solid organsHollow, viscus structuresTubular structuresFlat
tissue structures
Cornea
Trachea
Fig. 1. Four structural levels of complex tissues and organs. Hu-
man tissues and organs can be categorized generally into four
tures, such as the bladder; and solid organs, such as the kidney.
The
complexity of a tissue engineering approach generally increases
with
levels of structural complexity: flat tissue structures, such as
the
cornea; tubular structures, such as the trachea; hollow, viscus
struc-
the structure and metabolic functions of the tissue or organ
targeted
for repair.
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portions of the body or allogeneic grafts that provide temporary
pro-
tection, are the current clinical “gold standard.” However, skin
auto-
grafts require harvesting healthy tissue, which may not be
available in
adequate supply in some clinical cases, such as severe burns
affecting
large surface areas. Likewise, skin allografts present a risk of
immuno-
logic rejection and disease transmission.
Accordingly, several technologies are currently being used to
92. engi-
neer adequate skin for human replacement. Normal skin cells
are being
harvested from the patient in the operating room and are then
sprayed
over the burn area (62, 63). Patient-derived skin cells are also
being ex-
panded and layered ex vivo, with subsequent implantation over
the
burn area, thereby reducing the donor site morbidity required
for burn
coverage (64, 65). Although coverage of a burn with a tissue-
engineered
skin construct can facilitate repair, the size and severity of the
burn play
an important role in determining the ultimate outcome. Large,
full-
thickness burns present a greater clinical challenge for repair
than
small, superficial, partial-thickness burns because blood vessels
and
regenerative epithelial elements of the dermis are destroyed in
full-
thickness wounds. Nevertheless, clinical and commercial
success has
been realized with tissue engineering approaches for functional
repair
of skin in several applications (66, 67). The functional and
cosmetic
outcomes, however, may be improved through ongoing efforts
to re-
capitulate more fully with the tissue-engineered constructs the
com-
plex strata; vascular, lymphatic, and neural elements; pigment;
hair
follicles; and secretory glands of natural skin.
93. The cornea represents another flat tissue structure that has been
the target of biomaterial-based tissue engineering approaches
for re-
pair in the clinic. It performs a fundamental function in the
refraction
of light for vision and depends on maintenance of its
characteristic
transparency for efficacy. The cornea retains its transparency in
vivo
through maintenance of the shape and organization of a highly
aligned
collagen matrix and active pumping for continuous removal of
aqueous
humor from the tissue. A variety of disorders can disrupt proper
cor-
neal function, and surgical transplantation of donor corneal
tissue has
long served as a clinical standard of treatment for such
conditions.
However, transplantation requires procurement of donor tissue
matched
to the specific requirements of the recipient. To address this
challenge,
biomaterial-based tissue engineering approaches have been
developed
and translated clinically to enable corneal repair without the
need for
human donor tissue (68).
Tubular structures
Regenerative medicine has been able to successfully replicate
many
types of tubular structures, including the urethra, trachea, and
esoph-
agus, in both animals and humans. In general, these structures
94. consist
of two different cell types arranged as sheets of cells. These
sheets form
into circular, bilayered tissues, which usually serve as means of
trans-
porting fluid or air throughout the body. The tubular tissue
structures
are composed of an inner layer of epithelial or endothelial cells
that
provide a functional barrier and an outer layer of smooth muscle
and
connective tissue to provide support. Whereas the single cell–
layered
skin constructs do not require a complex foundation, tubular
structures
need to incorporate a matrix of synthetic or naturally derived
scaffold-
ing for support.
Tubular, tissue-engineered urethral constructs comprising
synthetic,
biodegradable poly(glycolic acid) (PGA)/PLGA scaffolds
seeded with
autologous urethral muscle and epithelial cells were implanted
into five
patients needing complex urethral reconstruction, and the
engineered
www.ScienceT
urethras remained functional over the clinical follow-up period
of up
to 6 years (69). For blood vessels, autologously derived cells
cultured
from peripheral vein biopsies have been grown in both
biodegradable
collagen and synthetic scaffolds and successfully used as
pulmonary
95. artery transplants (70). With a different method, vascular access
grafts
for patients with end-stage renal disease requiring hemodialysis
have
been engineered and implanted in humans (71). To accomplish
this,
fibroblasts and endothelial cells were harvested from patients,
ex-
panded ex vivo as sheets of cells, and then wrapped around a
stainless
steel cylinder to allow for fusion. In both situations (70, 71),
clinical
trials have yielded functional implants. Synthetic materials have
also
been seeded with cells to create new blood vessels for
implantation.
For example, human and canine smooth muscle cells were
cultured
on PGA tubular scaffolds, then treated with detergents to
produce
acellular vascular grafts capable of long-term storage. These
vessels
demonstrated patency in both baboon and canine models (72).
Decellularized scaffolds have been used to create tracheas. In
animal models, autologous chondrocytes cultured from cartilage
bi-
opsies were seeded in biodegradable collagen scaffolds and
success-
fully implanted in the pulmonary tree (73). Autologously
derived
chondrocytes have been differentiated from bone marrow MSCs,
and epithelial cells were isolated from a bronchial mucosa
biopsy.
The cells were seeded in the decellularized donor trachea and
cultured
96. in a bioreactor (74).
Hollow, viscus structures
Like tubular structures, hollow, viscus organs, such as the
bladder and
vagina, generally consist of an inner layer of epithelial-type
cells sur-
rounded by an outer layer of smooth muscle and/or connective
tissue
to provide functional capacity and to anchor the structure in
place.
Whereas tubular structures generally serve as conduits for air or
fluid,
viscus, nontubular organs have wider functional parameters,
higher
metabolic requirements, and more complex intracellular and
inter-
organ interactions. Similar to tubular organs, the biofabrication
pro-
cess depends on a scaffold seeded with at least two different
cell types.
However, the scaffold design is more complex in terms of both
its ar-
chitecture and its predetermined anatomical space limitations,
which
are often patient-specific. In addition, once the engineered
construct is
completed, there are special considerations for implanting the
engi-
neered construct and for connecting it with other tissues and
organs.
Regeneration of bladder tissue has been accomplished in
patients by
using autologously derived urothelial and smooth muscle cells
(75). A
computed tomography scan was performed on patients before
97. tissue
biopsy to determine the size of the organ to be constructed.
Thus, the
scaffold architecture and size were individualized for each
patient. The
cells were harvested, cultured in vitro, and seeded onto
biodegradable
collagen-PGA composite scaffolds, and the cell-scaffold
constructs
were placed in bioreactors to develop the tissues.
Similarly, a construct for vaginal replacement was synthesized
by
combining PGA fibers with a coating of PLGA, seeding this
scaffold
with rabbit epithelial cells, and culturing within a perfusion
bioreactor.
After implantation as a total vaginal replacement in a rabbit
model,
the biomaterial constructs yielded organs that were successfully
inte-
grated by the host animal and subsequently demonstrated
histological
characteristics similar to natural tissue after several months of
growth
(76). The engineered constructs had to be designed with
determined
specifications that would allow a patent connection with both
the uter-
us superiorly and the introitus opening inferiorly. As a result of
these
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experiments, human clinical trials for vaginal regeneration are
under
99. way (COFEPRIS HIM87120BSO).
Solid organs
Solid organs, such as the kidney, heart, pancreas, and liver,
have the
highest level of tissue complexity. The traditional treatment for
end-
stage solid organ disease is either temporary supportive
treatment with
drugs or devices or whole-organ transplantation. Conventional
trans-
plantation allows select patients to regain a functional organ,
yet it is
exceptionally complicated to obtain a histocompatible match
that does
not require the use of immunosuppressive agents. The ultimate
goal of
regenerative medicine is to bioengineer and transplant complex,
solid
organs composed of cells derived from the patient in need. This
ob-
jective, however, presents an exceedingly difficult and
challenging task
given the tissue complexity and developmental process of these
organs.
Complete regeneration of these whole organs requires
incorporation of
extensive vascular networks to support the viability of cells
throughout
the organ as well as precise organization of multiple cell
types—two
challenges traditionally not faced in the biofabrication process
of sim-
pler tissues like the skin. Whereas creation of flat, tubular, and
hollow
viscus organs primarily uses cell-seeded scaffolds, replication
100. of solid
organ function must incorporate other methods to be successful
because
these organs have complex architecture that extends beyond
simple lay-
ers. To this end, efforts are focused on the development of
biomaterial-
based approaches that incorporate gradients of growth factors,
hybrid
composite materials (77), and use 3D printing methods (37).
Patients with end-stage renal disease suffer major medical
sequelae
secondary to loss of the many physiological duties carried out
by
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the kidneys. With complete renal fail-
ure, these patients must undergo me-
chanical dialysis to replace the waste
disposal function of the kidneys and
must be closely monitored for electro-
lyte and …
BIOMEDICINE
3D bioprinting of collagen to rebuild
components of the human heart
A. Lee1*, A. R. Hudson1*, D. J. Shiwarski1, J. W. Tashman1,
T. J. Hinton1, S. Yerneni1,
J. M. Bliley1, P. G. Campbell1,2, A. W. Feinberg1,3†
Collagen is the primary component of the extracellular matrix in
the human body. It has proved
102. challenging to fabricate collagen scaffolds capable of
replicating the structure and function
of tissues and organs.We present a method to 3D-bioprint
collagen using freeform reversible
embedding of suspended hydrogels (FRESH) to engineer
components of the human heart
at various scales, from capillaries to the full organ. Control of
pH-driven gelation provides
20-micrometer filament resolution, a porous microstructure that
enables rapid cellular
infiltration and microvascularization, and mechanical strength
for fabrication and perfusion
of multiscale vasculature and tri-leaflet valves. We found that
FRESH 3D-bioprinted
hearts accurately reproduce patient-specific anatomical
structure as determined by
micro–computed tomography.Cardiac ventricles printed with
human cardiomyocytes showed
synchronized contractions, directional action potential
propagation, and wall thickening
up to 14% during peak systole.
F
or biofabrication, the goal is to engineer
tissue scaffolds to treat diseases for which
there are limited options, such as end-stage
organ failure. Three-dimensional (3D) bio-
printing has achieved important milestones
including microphysiological devices (1), pat-
terned tissues (2), perfusable vascular-like net-
works (3–5), and implantable scaffolds (6).
However, direct printing of living cells and soft
biomaterials such as extracellular matrix (ECM)
proteins has proved difficult (7). A key obstacle
103. is the problem of supporting these soft and dy-
namic biological materials during the printing
process to achieve the resolution and fidelity
required to recreate complex 3D structure and
function. Recently, Dvir and colleagues 3D-
printed a decellularized ECM hydrogel into a
heart-like model and showed that human car-
diomyocytes and endothelial cells could be in-
tegrated into the print and were present as
spherical nonaligned cells after 1 day in culture
(8). However, no further structural or functional
analysis was performed.
We report the ability to directly 3D-bioprint
collagen with precise control of composition and
microstructure to engineer tissue components
of the human heart at multiple length scales.
Collagen is an ideal material for biofabrication
because of its critical role in the ECM, where it
provides mechanical strength, enables struc-
tural organization of cell and tissue compart-
ments, and serves as a depot for cell adhesion
and signaling molecules (9). However, it is dif-
ficult to 3D-bioprint complex scaffolds using
collagen in its native unmodified form because
gelation is typically achieved using thermally
driven self-assembly, which is difficult to control.
Researchers have used approaches including
RESEARCH
Lee et al., Science 365, 482–487 (2019) 2 August 2019 1 of 5
1Department of Biomedical Engineering, Carnegie
Mellon University, Pittsburgh, PA 15213, USA. 2Engineering
104. Research Accelerator, Carnegie Mellon University, Pittsburgh,
PA 15213, USA. 3Department of Materials Science and
Engineering, Carnegie Mellon University, Pittsburgh, PA
15213, USA.
*These authors contributed equally to this work.
†Corresponding author. Email: [email protected]
Fig. 1. High-resolution
3D bioprinting of
collagen using FRESH
v2.0. (A) Time-lapse
sequence of 3D bioprint-
ing of the letters “CMU”
using FRESH v2.0.
(B) Schematic of acidified
collagen solution extruded
into the FRESH support
bath buffered to pH 7.4,
where rapid neutralization
causes gelation and
formation of a collagen
filament. (C and D) Rep-
resentative images of
the gelatin microparticles
in the support bath
for (C) FRESH v1.0 and
(D) v2.0, showing the
decrease in size and
polydispersity. (E) Histo-
gram of Feret diameter
distribution for gelatin
microparticles in FRESH
v1.0 (blue) and v2.0
(red). (F) Mean Feret
diameter of gelatin micro-
particles for FRESH v1.0
and v2.0 [N > 1200, data are means ± SD, ****P < 0.0001
105. (Student t test)]. (G) Storage (Gʹ) and loss (Gʺ) moduli for
FRESH v1.0 and v2.0 support
baths showing yield stress fluid behavior. (H) A “window-
frame” print construct with single filaments across the middle,
comparing G-code (left), FRESH v1.0
(center), and FRESH v2.0 (right). (I) Single filaments of
collagen showing the variability of the smallest diameter (~250
mm) that can be printed using
FRESH v1.0 (top) compared to relatively smooth filaments 20
to 200 mm in diameter using FRESH v2.0 (bottom). (J)
Collagen filament Feret diameter as a
function of extrusion needle internal diameter for FRESH v2.0,
showing a linear relationship.
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107. moreversible. However, these hydrogels are typ-
ically soft and tend to sag, and they are difficult
to print with high fidelity beyond a few layers
in height. Instead, we developed an approach
that uses rapid pH change to drive collagen self-
assembly within a buffered support material,
enabling us to (i) use chemically unmodified
collagen as a bio-ink, (ii) enhance mechanical
properties by using high collagen concentra-
tions of 12 to 24 mg/ml, and (iii) create complex
structural and functional tissue architectures.
To accomplish this, we developed a substantially
improved second generation of the freeform
reversible embedding of suspended hydrogels
(FRESH v2.0) 3D-bioprinting technique used
in combination with our custom-designed open-
source hardware platforms (fig. S1) (14, 15).
FRESH works by extruding bio-inks within a
thermoreversible support bath composed of
a gelatin microparticle slurry that provides
support during printing and is subsequently
melted away at 37°C (Fig. 1, A and B, and
movie S1) (16).
The original version of the FRESH support
bath, termed FRESH v1.0, consisted of irregularly
shaped microparticles with a mean diameter of
~65 mm created by mechanical blending of a
large gelatin block (Fig. 1C) (16). In FRESH v2.0,
we developed a coacervation approach to gen-
erate gelatin microparticles with (i) uniform
spherical morphology (Fig. 1D), (ii) reduced poly-
dispersity (Fig. 1E), (iii) decreased particle diam-
eter of ~25 mm (Fig. 1F), and (iv) tunable storage
modulus and yield stress (Fig. 1G and fig. S2).
108. FRESH v2.0 improves resolution with the ability
to precisely generate collagen filaments and ac-
curately reproduce complex G-code, as shown
with a window-frame calibration print (Fig. 1H).
Using FRESH v1.0, the smallest collagen filament
reliably printed was ~250 mm in mean diameter,
with highly variable morphology due to the rela-
tively large and polydisperse gelatin micropar-
ticles (Fig. 1I). In contrast, FRESH v2.0 improves
the resolution by an order of magnitude, with
collagen filaments reliably printed from 200 mm
down to 20 mm in diameter (Fig. 1, I and J).
Filament morphology from solid-like to highly
porous was controlled by tuning the collagen
gelation rate using salt concentration and buffer-
ing capacity of the gelatin support bath (fig. S3).
A pH 7.4 support bath with 50 mM HEPES was
the optimal balance between individual strand
resolution and strand-to-strand adhesion and
was versatile, enabling FRESH printing of mul-
tiple bio-inks with orthogonal gelation mech-
anisms including collagen-based inks, alginate,
fibrinogen, and methacrylated hyaluronic acid in
the same print by adding CaCl2, thrombin, and
UV light exposure (fig. S4) (15).
We first focused on FRESH-printing a sim-
plified model of a small coronary artery–scale
linear tube from collagen type I for perfusion with
a custom-designed pulsatile perfusion system
(Fig. 2A and fig. S5). The linear tube had an inner
diameter of 1.4 mm (fig. S6A) and a wall thick-
ness of ~300 mm (fig. S6B), and was patent and
manifold as determined by dextran perfusion
(fig. S6, C to E, and movie S2) (15). C2C12 cells
![Title
ABC/123 Version X
1
Leadership Newsletter Article
LDR/300 Version 7
1
University of Phoenix Material
Headline
[The title or overall description of the newsletter article]
Newsletter Tips
1. Interview your individual and research any information
necessary.
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an article that is engaging and worthwhile.
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audience’s attention.
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leadership profile.
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Opening
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