Biomimetic hydrogels with pro-angiogenic properties

Abstract

To achieve the task of fabricating functional tissues, scaffold materials that can be sufficiently vascularized to mimic functionality and complexity of native tissues are yet to be developed. Here, we report development of synthetic, biomimetic hydrogels that allow the rapid formation of a stable and mature vascular network both in vitro and in vivo. Hydrogels were fabricated with integrin binding sites and protease-sensitive substrates to mimic the natural provisional extracellular matrices, and endothelial cells cultured in these hydrogels organized into stable, intricate networks of capillary-like structures. The resulting structures were further stabilized by recruitment of mesenchymal progenitor cells that differentiated into a smooth muscle cell lineage and deposited collagen IV and laminin in vitro. In addition, hydrogels transplanted into mouse corneas were infiltrated with host vasculature, resulting in extensive vascularization with functional blood vessels. These results indicate that these hydrogels may be useful for applications in basic biological research, tissue engineering, and regenerative medicine.

Full text

Biomimetic hydrogels with pro-angiogenic properties
James J. Moon
a
, Jennifer E. Saik
a
, Ross A. Poche
´
b
, Julia E. Leslie-Barbick
a
, Soo-Hong Lee
a
, April A. Smith
a
,
Mary E. Dickinson
b
, Jennifer L. West
a
,
*
a
Department of Bioengineering, Rice University, P.O. Box 1892, MS 142, Houston, TX 77251-1892, USA
b
Department of Molecular Physiology and Biophysics, Baylor College of Medicine, One Baylor Plaza BCM 335, Houston, TX 77030, USA
article info
Article history:
Received 8 January 2010
Accepted 15 January 2010
Available online 24 February 2010
Keywords:
Hydrogel
Endothelial cell
Smooth muscle cell
Mesenchcymal stem cell
abstract
To achieve the task of fabricating functional tissues, scaffold materials that can be sufficiently vascu-
larized to mimic functionality and complexity of native tissues are yet to be developed. Here, we report
development of synthetic, biomimetic hydrogels that allow the rapid formation of a stable and mature
vascular network both in vitro and in vivo. Hydrogels were fabricated with integrin binding sites and
protease-sensitive substrates to mimic the natural provisional extracellular matrices, and endothelial
cells cultured in these hydrogels organized into stable, intricate networks of capillary-like structures. The
resulting structures were further stabilized by recruitment of mesenchymal progenitor cells that
differentiated into a smooth muscle cell lineage and deposited collagen IV and laminin in vitro. In
addition, hydrogels transplanted into mouse corneas were infiltrated with host vasculature, resulting in
extensive vascularization with functional blood vessels. These results indicate that these hydrogels may
be useful for applications in basic biological research, tissue engineering, and regenerative medicine.
!2010 Elsevier Ltd. All rights reserved.
1. Introduction
Most successes in tissue engineering have been limited to thin
or avascular tissues such as skin, bladder, and cartilage as these
constructs are relatively simple in design without requirement for
intricate blood vessels [1–4]. Although there have been recent
advances in vascularization of tissue constructs in vivo [5], devel-
opment of complex tissues or organs such as heart, kidney, liver and
lung has been elusive due to the lack of proper formation of
vasculature in the engineered constructs. Thus, the most impend-
ing challenge in creating more complex and clinically relevant
tissues is vascularization of engineered tissues.
The process of angiogenesis is achieved by complex interactions
among endothelial cells (ECs), the interstitial extracellular matrix
(ECM), and the neighboring mural cell types via various growth
factors [6,7]. In the initial phase of angiogenesis, vascular endo-
thelial growth factor (VEGF) activates ECs from their normal
quiescent states. When activated, the ECs proliferate and secrete
various proteases, including matrix metalloproteinases (MMPs), to
degrade the basement membrane and ECM. ECs migrate and
extend sprouts to build tubular structures. These nascent vessels
are stabilized by recruitment of mural cells such as mesenchymal
stem cells (MSCs), which differentiate into pericytes and deposit
new ECM proteins to form the basal lamina [8,9].
As these multi-components are all essential parts of neo-
vascularization, the design of pro-angiogenic tissue constructs
needs to address each component in order to truly mimic the
physiological microenvironment in which ECs can form functional
blood vessels. The objectives of the present study were to integrate
cellular, biochemical, and biophysical cues in synthetic biomaterials
to achieve extensive vascularization both in vitro and in vivo.To
achieve this, hydrogels mimicking natural provisional ECM were
synthesized and assessed as a scaffold for angiogenesis.
To fabricate synthetic ECM-mimicking biomaterials, we used
protease-sensitive poly(ethylene glycol) (PEG) hydrogels, first
reported by West and Hubbell [10]. Protease-sensitive peptides
were introduced into the backbone of PEG to render PEG hydrogels
biodegradable in response to cellular proteases. These types of
materials can also be modified with cell-adhesive sequences for use
as scaffolds in regenerative medicine [11–16]. In the present study,
we examined proteolytically-degradable and cell-adhesive PEG
hydrogels as provisional matrices for angiogenesis both in vitro and
in vivo. To form and stabilize new blood vessels in hydrogels, we
exploited cellular interactions between ECs and MSCs. Co-culture of
ECs and MSCs in hydrogels led to formation of extensive tubule-like
structures that were stabilized and not subject to regression during
long term culture in vitro. In addition, we demonstrate vasculari-
zation of these hydrogels with functional blood vessels in vivo.
*Corresponding author. Tel.: þ1 713 348 5955; fax: þ1 713 348 5877.
E-mail address: jwest@rice.edu (J.L. West).
Contents lists available at ScienceDirect
Biomaterials
journal homepage: www.elsevier.com/locate/biomaterials
0142-9612/$ – see front matter !2010 Elsevier Ltd. All rights reserved.
doi:10.1016/j.biomaterials.2010.01.104
Biomaterials 31 (2010) 3840–3847

2. Materials and methods
2.1. Cell maintenance
Human umbilical vein endothelial cells (HUVECs) were obtained from Cambrex
(East Rutherford, NJ). The cells were grown in endothelial cell medium EGM-2
(Cambrex) supplemented with 2 m
ML
-glutamine, 1000 U/mL penicillin, and 100 mg/
L streptomycin (Sigma, St. Louis, MO), and they were used throughpassage 8. 10T1/2
cells (American Type Culture Collection, Rockville, MD) were grown and maintained
in DMEM supplemented with 10% fetal bovine serum (FBS), 2 m
ML
-glutamine,
1000 U/mL penicillin, and 100 mg/L streptomycin, and used through passage 18. All
cells were incubated at 37
!
C in a 5% CO
2
environment.
2.2. Synthesis of PEG-polymer hydrogel precursors
An MMP-sensitive peptide sequence, GGGPQGYIWGQGK was synthesized by
solid phase peptide synthesis based on standard Fmoc chemistry using an Apex396
peptide synthesizer (Aapptec, Louisville, KY, USA). Hydrogels with this peptide in the
polymer backbone have been shown to be completely degraded by matrix
metalloproteinases (MMP) [11]. Following purification, synthesis of the peptide was
confirmed with matrix-assisted laser desorption ionization time of flight mass
spectrometry (MALDI-ToF; Bruker Daltonics, Billerica, MA, USA).
Synthesis of the ABA block copolymers with the peptide linker was completed as
described previously [10]. Briefly, the MMP-sensitive peptide sequence was reacted
with acrylate-PEG-N-hydroxysuccinimide (acrylate-PEG-NHS, 3400 Da; Nektar) in
a 1:2 (peptide: PEG) molar ratio in 50 m
M
sodium bicarbonate buffer (pH 8.5) for 2 h.
This step conjugates a PEG-monoacrylate chain to the N-terminus and to the amine
group on the lysine at the C-terminus of the peptide. The resulting product was then
dialyzed (MWCO 5,000; Spectrum Laboratories, Inc., Rancho Dominguez, CA, USA)
to remove unreacted peptide and PEG moieties, and the product was lyophilized,
frozen, and stored under argon at "80
!
C. The cell-adhesive ligand, Arg-Gly-Asp-Ser,
(RGDS, American Peptide, Sunnyvale, CA) was conjugated to acrylate-PEG-NHS in
1:1 molar ratio under similar conditions. Fluorophore-tagged PEG-RGDS was
synthesized by reacting PEG-RGDS with Alexa Fluor-488 carboxylicacid (Invitrogen)
in 10-fold dye molar excess in DMF. VEGF
165
(Sigma) was conjugated to acrylate-
PEG-SMC in 200:1 molar ratio at pH 8.5 at 4
!
C for 4 days, followed by final 4 h at
24
!
C. The resulting PEG-VEGF solution was then lyophilized and reconstituted in
HEPES buffered saline (HBS) with 0.1% BSA at 4
!
C until use. A gel permeation
40
80
120
160
15%12.5%10%7.5%
Pol
y
mer Wei
g
ht %
Compressive
moduli (kPa)
Gels only
With cells
Incubation duration (hrs)
50
100
43210
Degradation (%) Degradation (%)
50
100
43210
Col Plasmin Buffer
7.50%
10.0%
12.5%
15.0%
1
2
3
4
5
Tubule length
(relative)
1
2
No. of branching
points (relative)
Day 1
Day 3
Day 6
**
*
*
*
D
G
D3
7.5% 12.5%10% 15%
AB
C
E
F
D6
D6
Polymer Weight %
Fig. 1. Proteolytically-degradable PEG hydrogels with intermediate rigidity support endothelial tubule formation and maintenance. A) The schematic illustration shows fabrication
of MMP-sensitive PEG hydrogels by UV polymerization of PEG-RGDS and PEG chain with MMP-sensitive peptide in its backbone in the presence of cells. B) Hydrogels were degraded
by collagenase (10
m
g/mL), but not by plasmin (10
m
g/mL) or buffer solutions. C) Hydrogels fabricated with different polymer weight percentages exhibited varying degradation
profiles in collagenase (10
m
g/mL) solution. D) Confocal and bright field images of HUVECs and 10T1/2 cells cultured for 3 and 6 days in hydrogels with varying polymer weight
percentages. HUVECs and 10T1/2 cells were pre-stained with CytoTracker green and red, respectively, prior to encapsulation in hydrogels. E) The total tubule length formed and F)
the number of branching point were measured during a 6 day culture period in hydrogels formulated with varying polymer weight percentages. Only in hydrogels with inter-
mediate polymer weight percentage (10%), HUVECs and 10T1/2 cells formed stable tubule-like structures. G) Compressive moduli ranging from 30 to 110 kPa were measured in
hydrogels with and without encapsulated cells. Data represent mean #SD. (n¼4 for B,C,G; n¼9 for E,F). *P<0.05, analyzed by two-way ANOVA followed by Tukey’s HSD test. Scale
bars ¼50
m
m.
J.J. Moon et al. / Biomaterials 31 (2010) 3840–3847 3841

chromatography system equipped with UV–vis and evaporative light scattering
detectors (Polymer Laboratories, Amherst, MA) was used to analyze the products.
2.3. Fabrication of MMP-sensitive PEG hydrogels
The hydrogel precursor solution was prepared in 10 m
M
HBS (pH 7.4) with PEG-
RGDS (3.5
m
M
/mL), a photoinitiator, Irgacure-2969 (0.3 mg/mL; Ciba Corporations,
Basel, Switzerland), and varying amount of MMP-sensitive PEG-GGGPQGIWGQGK-
PEG. In this work, MMP-sensitivePEG precursor was added to the final concentrations
of 7.5 mg/mL, 10 mg/mL, 12.5 mg/mL, and 15 mg/mL to fabricate hydrogels with
polymer weight percentages of 7.5%, 10%, 12.5%, and 15%, respectively. For HUVECs
only culture, 6 !10
6
cells were added to the hydrogel precursor solution. For HUVEC
and 10T1/2 co-culture, 4.8!10
6
HUVECs and 1.2 !10
6
10T1/2 cells were added. On
coverglasses that had been cleaned with 70% ethanol and sterilized under UV lamp
overnight, the PEG precursor-cell suspension was dispensed as 5
m
L droplets and
photopolymerized by exposure to long-wavelength UV light (365 nm, 10 mW/cm
2
)
for 7 min. The hydrogels were then immersed in EGM-2 media and incubated at 37
"
C
in a humidified atmosphere containing 5% CO
2
. For PEG hydrogels used in corneal
angiogenesis assays in vivo, VEGF was mixed into the polymer solution to create
concentrations of 512 ng VEGF per geland 1.9 ng PEG-VEGF per gel.
2.4. Characterization of hydrogels
The swelling ratio and water content of the hydrogels were determined as
described previously [17]. The compressive moduli of the different hydrogels with
and without cells were determined using a 10 N load cell at a crosshead speed of
0.5 mm/min in an Instron-3342 (Canton, MA) mechanical tester as previously
described [18]. Degradation rates of MMP-sensitive PEG hydrogels formulated with
varying polymer weight percentages were measured by monitoring the release of
tryptophan (W) in the MMP-sensitive peptide sequence with a UV–Vis spectro-
photometer (Carey 5000 Varian, Walnut Creek, CA) at 280 nm.
2.5. Measurement of tubule length and the number of branching points in 3D
network of hydrogels
Endothelial tubule formation was evaluated by measuring total tubule length
formed and the number of branching points over 6 days. On day 1, 3, and 6,
cells in the hydrogels were stained with 2
m
M
calcein AM and visualized with
a confocal microscope (Zeiss LIVE 5, Carl Zeiss, Thornwood, NY) with zstack
depth ranging 100–200
m
m. Scion Image (Scion Corporation, Frederick, MD) was
used to trace and measure the total tubule length and the number of branching
points in each area. The data were normalized to the zstack depth in each area
to account for the different optical slice volumes of the images.
2.6. Time-lapse video confocal microscopy
To record time-lapse images during endothelial tubule formation, HUVECs
and 10T1/2 were pre-labeled with 50
m
g CellTracker Green CMFDA (5-chlor-
omethylfluorescein diacetate; Invitrogen) and 50
m
g CellTracker Red CMTPX
(Invitrogen), respectively, per manufacturer’s instructions prior to encapsulation
200
400
600
800
1000
1200
605040302010
Hrs
Total tubule length ( m)
CF
E
D
A
B
5 hr 21 hr 37 hr 53 hr 69 hr
HUVEC
HUVEC + 10T1/2
Fig. 2. Time-lapse confocal videomicroscopy shows HUVECs and 10T1/2 undergoing tubule formation in hydrogels. Cellular interactions in hydrogels encapsulated with either A)
HUVECs only or B) HUVECs and 10T1/2 cells were visualized over w70 h with time-lapse confocal videomicroscopy. A) In HUVEC mono-culture conditions, tubule-like structures
initially formed by cells failed to maintain their networks and quickly regressed after w50 h of encapsulation in hydrogels. B) Tubule-like structures formed in co-culture conditions
maintained their morphologies throughout the time-lapse experiments. C) The movies were analyzed quantitatively to plot tubule length formed over time. Whereas the total
tubule length formed by HUVECs alone initially increased and sharply declined, there was a gradual increase and maintenance of the tubule length in co-culture conditions. D)
HUVECs with large vacuoles in their cell bodies and E) in the process of intercellular fusion of multiple vesicles to form large lumens were observed.10T1/2 cells were recruited to
the newly formed lumen or tubular structures. A,B,D,E) HUVECs and 10T1/2 cells were pre-stained with CellTracker Green and Red, respectively, prior to encapsulation into the
hydrogels. F) HUVECs and 10T1/2 cells cultured for 28 days in hydrogels maintained the highly interconnected networks of tubules. F-actin and nuclei were stained by phalloidin-
TRITC and DAPI, respectively. Data represent mean #SD (n¼10–12). *P<0.05, analyzed by two-way ANOVA followed by Tukey’s HSD test compared to the HUVEC mono-culture
groups. Scale bars ¼50
m
m.
J.J. Moon et al. / Biomaterials 31 (2010) 3840–38473842

in the hydrogels. After photoencapsulation of the cells, the hydrogels were
incubated in EGM-2 media for 5 h before time-lapse imaging. A confocal
microscope (Zeiss LIVE 5) equipped with a motorized XYZ stage and a temper-
ature hood was used to record time-lapse images. The temperature hood was
perfused with 5% CO
2
during the experiments. Confocal images were obtained at
1 h intervals for 66 h, and the motorized stage allowed automatic collection of
images from pre-recorded, multiple locations.
2.7. Isolation of cellular lysates and Western blotting
The hydrogels were prepared as 5
m
L droplets each with either HUVEC mono-
culture (24 !10
6
cells/mL), 10T1/2 mono-culture (6 !10
6
cells/mL), or HUVECs and
10T1/2 co-culture (24!10
6
and 6 !10
6
cells/mL, respectively). After 6 days, the
cells were isolated from the bulk of hydrogels by degrading away the hydrogels
with 2 mg/mL collagenase-1 solution (Sigma) for 10 min at 37
"
C. Cell suspensions
from four 5
m
L hydrogel droplets were pooled for each cell culture condition to
obtain enough cell lysate for Western blotting. In parallel experiments, cells were
plated on 6 well tissue culture plates with either HUVEC mono-culture (24 !10
4
cells/well), 10T1/2 mono-culture (6 !10
4
cells/well), or HUVECs and 10T1/2 co-
culture (24 !10
4
and 6 !10
4
cells/well, respectively). SDS-PAGE and Western
blotting were performed following a standard protocol. The primary antibodies
used for Western blotting include antibodies against smooth muscle
a
-actin (SM-
a
actin, 1:3000; ABCam, Cambridge, MA), calponin (1:500; ABCam), and caldesmon
(1:500; Sigma). The secondary antibody was used at 1:2000, 1:1000, and 1:1000
dilutions, respectively.
2.8. Immunofluorescence staining
Immunofluorescence staining was performed to confirm differentiation of
10T1/2 cells into SMC lineages and to visualize deposition of ECM proteins
adjacent to the tubular structures. On days 1 and 6, the hydrogels containing
HUVEC and 10T1/2 co-culture were fixed, permeabilized, and stained with
primary and secondary antibodies. The primary antibodies used were mouse
anti-SM
a
-actin (Sigma), rabbit anti-CD31 (Bethyl), rabbit anti-collagen type IV
(ABCam), and rabbit anti-laminin (Sigma) IgGs. The secondary antibodies were
anti-rabbit and anti-mouse antibodies conjugated with either Alexa flour 488,
Alexa fluor 594, or rhodamine (Invitrogen). For some samples, actin cytoskele-
tons and nuclei were stained with TRITC-conjugated phalloidin (5 U/mL, Sigma)
for 1 h and DAPI (300 n
M
, Invitrogen) for 5 min.
2.9. Hydrogel implantation into the mouse cornea
Using a modified version of the previously described corneal micropocket
angiogenesis assay [19], hydrogels were implanted into mouse cornea in Flk1-
myr::mCherry transgenic mouse, which exhibits EC-specific expression of a myr-
istoylated mCherry fluorescence protein in the EC membrane [20]. Briefly, mice
were anesthetized, and a partial thickness incision was made into the mouse
cornea. The micropocket was created using a von Graef knife, and hydrogels
were implanted into the micropocket immediately after UV photopolymerization.
Seven days post-implantation, some mice were injected intravenously with
dextran-Texas red (MW 70 KDa), and the mice were sacrificed with CO
2
asphyxiation. Eyes from the mice were enucleated and fixed in 4% para-
formaldehyde for 1 h at 4
"
C. Corneal flat-mount preparations were made, and
imaging was performed using a Zeiss LSM 510 META inverted microscope system
(Carl Zeiss Inc) equipped with a Zeiss Plan-Apochromat 20!/0.75 NA objective.
543-nm and 488-nm lasers were used to excite the Flk1-myr::mCherry and flu-
orophore-tagged PEG-RGDS, respectively.
2.10. Statistical analysis
Statistical analysis was performed with Jmp 5.1 (SAS Institute Inc, Cary, NC).
Datasets were analyzed using two-way analysis of variance (ANOVA), followed by
Fig. 3. The tubule structures are stabilized by smooth muscle-like cells differentiated from 10T1/2 cells and by new deposition of collagen IV and laminin in hydrogels. A) HUVECs
and 10T1/2 cells each cultured in mono-culture conditions had minimal expression of SM-
a
actin, calponin, and caldesmon on tissue culture wells and in 3D network of hydrogels as
shown by Western blotting. In contrast, in co-culture conditions, the expression levels of these SMC protein markers were dramatically up-regulated both on tissue culture wells
and in the hydrogels by day 6. B) In hydrogels with HUVECs and 10T1/2 cells, there was minimal expression of SM-
a
actin on day 1, but C) by day 6, expression of SM-
a
actin was
localized adjacent to CD31 staining specific for ECs, indicating that 10T1/2 cells up-regulated expression of the SMC marker protein. In the PEG hydrogels cultured with HUVECs and
10T1/2 cells for 1 day, there was minimal expression of D) collagen type IV or F) laminin; however, by day 6, the tubule-like structures were highly decorated with E) collagen type
IV and G) laminin, indicating that the encapsulated cells are actively producing their own set of ECM proteins, thereby remodeling the synthetic matrices. B,C) Anti-CD31
immunostainings and anti-SM-
a
actin are shown in green and red, respectively. D–G) Anti-CD31 is shown in red, while anti-collagen type IV (D,E) and anti-laminin (F,G) are shown
in green. Scale bars ¼50
m
m.
J.J. Moon et al. / Biomaterials 31 (2010) 3840–3847 3843

Tukey’s HSD test for multiple comparisons. P-values less than 0.05 were considered
statistically significant. All values are reported as mean !standard deviation.
3. Results and discussion
3.1. Proteolytically-degradable hydrogel and its mechanical
characterization
To fabricate synthetic ECM-mimicking hydrogels used in the
current studies, we incorporated an MMP-sensitive sequence
(GGGPQGYIWGQGK) into the backbone of PEG block polymers with
acrylate terminal groups, which allow crosslinking of precursors
into networks [10,11,21] (Fig. 1A). GGGPQGYIWGQGK is a mutated
version of
a
1(I) collagen chain for increased degradation kinetics
with various MMPs [11,22]. In addition, a ligand for integrin, RGDS,
was grafted onto PEG chains to support integrin-mediated cellular
adhesion and migration while VEGF was incorporated into hydro-
gels to promote robust angiogenesis [12,23].
We first characterized protease-mediated degradation of
hydrogels. Degradation of the hydrogels was monitored by
measuring release of tryptophan from the MMP-sensitive
sequences in hydrogels incubated with collagenase, a subset of
MMP family of proteases (Fig. 1B). Hydrogels with 10% polymer
weight percentage incubated with collagenase solution underwent
complete degradation within 1.5 h, but not with either plasmin or
the buffer solution even after 21 h, demonstrating specificity of
degradation mediated by the MMP family of proteases. Suscepti-
bility of hydrogels to proteolysis was correlated with their polymer
weight percentage as denser networks took progressively longer
time to degrade (Fig. 1C). As the polymer weight percentage was
increased from 7.5% to 15%, the compressive modulus of the
hydrogels was increased from 30.2 !6.05 kPa to 110 !23.8 kPa
(Fig. 1G). Encapsulation of cells did not alter the compressive
moduli of hydrogels. The swelling ratios for the 7.5,10,12.5, and 15%
hydrogels were 27.9 !8.37, 17.0 !2.59, 14.7 !3.11, and 13.2 !2.33,
and their water contents were 96.2 !1.11, 94.0 !0.903, 92.9 !1. 47,
and 92.3 !1.28%, respectively.
3.2. Co-culture of HUVECs and 10T1/2 cells in hydrogels for
angiogenesis
To form and stabilize new blood vessels in hydrogels, we
exploited heterotypic cell interactions between ECs and mesen-
chymal progenitor cells. ECs are able to form primitive tubule
Fig. 4. Proteolytically-degradable PEG hydrogels promote neovascularization in murine cornea. Hydrogels incorporated with soluble and immobilized forms of VEGF via PEG
linkage were implanted into cornea in Flk1-myr::mCherry transgenic mice. Newly formed blood vessels were visualized by confocal microscopy, and depth profiles of the vessels
were generated to reveal their Z position with respect to the hydrogels. A–D) Non-degradable hydrogels releasing soluble VEGF supported angiogenesis adjacent to the hydrogels,
but the depth decoding graph indicates lack of vessel penetration into the hydrogels. E–H) MMP-sensitive PEG hydrogels with 10% polymer weight promoted robust neo-
vascularization adjacent to the hydrogels and extensive infiltration of host vasculature into the hydrogels. The arrow in E) indicates a portion hydrogel undergoing active
degradation, and the arrow in H) points to regions within hydrogels with vessel infiltration. I–L) MMP-sensitive PEG hydrogels with 15% polymer weight had blood vessel growth on
surface of hydrogels as pointed by the arrow in L), but the hydrogels remained mostly intact (I) without any significant vessel infiltration into the core. Scale bars ¼100
m
m in A–L).
J.J. Moon et al. / Biomaterials 31 (2010) 3840–38473844

structures when cultured alone; however, for the rudimentary blood
vessels to become matured and stabilized, they require interaction
with mural cell types. 10T1/2 cells, a subset of multipotent mesen-
chymal progenitor cells, are known to support endothelial tubule
formation in naturally occurring ECM-derived materials [24–26].
MMP-sensitive PEG hydrogel precursors were formulated with
human umbilical vein endothelial cells (HUVECs) and 10T1/2 cells
(30 million cell/mL with HUVEC:10T1/2 ratio of 4:1) and exposed to
long-wavelength UV to photopolymerize the precursors into
hydrogels. The cells encapsulated in hydrogels were cultured, and
examined for their angiogenic responses at various time points
(Fig. 1D–F). In order to find the optimal mechanical composition of
the hydrogels for the angiogenic responses, tube formation by
HUVECs and 10T1/2 cells was quantified in hydrogels with 7.5–15%
polymer weight percentage. In hydrogels with an intermediate
polymer weight percentage of 10%, HUVECs and 10T1/2 cells
Fig. 5. Newly formed blood vessels in hydrogels are functional. A,B) Small incision was made in cornea and photopolymerized hydrogels were implanted into the micropocket.
C). Blood vessels formed in the hydrogels were perfused with Dextran-Texas red (70 KDa MW) injected intravenously into mice.
J.J. Moon et al. / Biomaterials 31 (2010) 3840–3847 3845

formed primitive tubule-like structures by day 3 and continued to
mature into extensive interconnections throughout the 3D network
of hydrogels over 6 days. In soft, compliant hydrogels with 7.5%
polymer weight, the cells underwent self-assembly to form thick
interconnections, but the resulting structures rapidly regressed as
the gels were completely degraded within 6 days. In stiffer
hydrogels with 12.5 or 15% polymer weight, the degree of tubule
formation was reduced as cells failed to migrate as much in softer
matrices, and the cell clusters remained short and rudimentary. In
contrast, HUVECs and 10T1/2 cells co-cultured on tissue culture
plates for 14 days did not form any tubule-like structures but
remained as monolayers. The varying degrees of angiogenic
responses in these hydrogels did not seem to result from different
cellular viability as more than 80% of cells remained viable after 6
days in all hydrogel formulations tested, which is within a typical
range reported in other work using hydrogel systems
(Supplementary Figure 1)[27,28]. These results show that angio-
genesis organized and promoted by HUVECs and 10T1/2 cells is
well recapitulated in proteolytically-degradable hydrogels and that
there is a particular formulation of hydrogels with an intermediate
rigidity that maximizes these angiogenic responses.
These results do not demonstrate mechanical properties of
hydrogels as a sole determinant of angiogenic potential since other
parameters such as density of cell-adhesive ligand or degradable
sites were not held constant. However, since 10% polymer weight
PEG hydrogels produced robust angiogenic responses, all the
subsequent studies were performed with this particular formula-
tion of hydrogels unless noted otherwise.
3.3. Dynamics of tubule formation by HUVECs and 10T1/2 cells
To better understand the dynamics of angiogenesis in the
hydrogels, the process of tubule formation was captured by time-
lapse confocal microscopy. In hydrogels with HUVECs in mono-
culture condition (30 million cells/mL), the cells started migrating
toward each other to form large clusters (t¼5 h) and organized into
primitive tubule-like structures (t¼21 h) (Fig. 2A, C, and Supporting
Video 1). However, at later time points (t¼53 and 69 h), the struc-
tures lost their tubular morphology and rapidly regressed into
individual cell clusters. The total tubule length decreased over the 6
day culture period, while the number of branching points stayed at
minimal levels (Supplementary Figure 2).
In contrast, HUVECs and 10T1/2 cells co-cultured in the hydrogels
(30 million cell/mL with HUVEC:10T1/2 ratio of 4:1) formed large
clusters that rapidly organized into tubule-like structures throughout
the hydrogels (t¼21 and 37 h) (Fig. 2B, and Supporting Video 2). The
tubule-like structures were maintained stably even at later time
points (t¼53 and 69 h) with extensive branches and typical length
reaching 700
m
m(Fig. 2C, and Supplementary Figure 2).
The endothelial tubular structures formed were reminiscent of
capillary structures in their cellular organizations and morphology.
10T1/2 cells were frequently observed to align adjacent to and
migrate rapidly along the tubular structures that were assembled
by HUVECs. Fig. 2D shows HUVECs undergoing lumen formation
and 10T1/2 cells adjacent to the ECs supporting the process. Subsets
of the HUVECs had multiple small vesicles, while others had their
vesicles fused together intercellularly to form lumens enclosed by
multiple ECs (Fig. 2D, E), and these processes of lumen formation
and perivascular association of mural cell types parallel a series of
physiological events that accompany neovascularization as
described recently in vivo [29]. In addition, HUVECs and 10T1/2
cells kept in culture for 28 days maintained their extensivenetwork
of tubular structures throughout the hydrogels (Fig. 2F), demon-
strating their durability. The hydrogels remained intact and stable
when examined up to 1 month later. Taken together, these results
suggest that 10T1/2 cells enhance endothelial tubule formation and
maintenance in hydrogels.
3.4. Differentiation of 10T1/2 cells toward smooth muscle cell
lineage
We next addressed the question whether 10T1/2 MSCs differ-
entiate toward SMC lineages in co-cultureconditions with HUVECs.
10T1/2 cells are known to up-regulate expressions of smooth
muscle marker proteins, such as SM-myosin, SM22
a
, and calponin
when co-cultured together with ECs [24]. We indeed observed
dramatic up-regulation of SMC protein markers such as SM
a
-actin,
calponin, and caldesmon in the co-culture conditions in hydrogels,
but not in mono-culture conditions of either cell type as examined
by Western blotting (Fig. 3A). Similar results were obtained from
cells cultured in tissue culture plates. Over-proliferation of either
cell type contributing to this result can be ruled out since time-
lapse confocal images revealed a minimal number of cell division
during 72 h of culture as reported in the literature [30].
Differentiation of 10T1/2 cells into SMC lineages was further
confirmed by immunofluorescence staining. There was a minimal
level of SM
a
-actin staining on day 1 in the hydrogels with co-
culture conditions, while anti-CD31 staining showed HUVECs
dispersed in the matrix (Fig. 3B). After 6 days, anti-CD31 staining
showed HUVECs in tubule-like structures, whereas the cells stained
with anti-SM
a
-actin antibody aligned adjacent to the HUVECs
(Fig. 3C). Taken together, these results suggest that 10T1/2
precursor cells differentiate toward SMC lineages in co-culture
conditions and associate closely with the tubule-like structures
composed of HUVECs.
3.5. Cellular remodeling of hydrogels
Collagen type IV and laminin, both the major components of
basal lamina of blood vessels [31,32], were found to be secreted and
deposited along the tubule-like structures (Fig. 3D–G). This indi-
cates that these cells actively remodel the matrices by secreting
their own set of ECM proteins, which are known to serve as
a reservoir for various growth factors and provides structural
integrity to newly formed capillaries [33].
3.6. Formation of functional blood vessels in vivo
Proteolytically-degradable PEG hydrogels were implanted into
mouse cornea using a micropocket angiogenesis assay [19] and
examined 7 days post-implantation (Fig. 5A,B). We utilized Flk1-
myr::mCherry transgenic mice, which exhibit EC-specific expres-
sion of a reporter mCherry in EC membrane [20] and allow moni-
toring of host vasculature invading into the hydrogels by confocal
microscopy. In order to recruit the host blood vessels into hydro-
gels, pro-angiogenic growth factor, VEGF, was incorporated into the
hydrogels both in a soluble and an immobilized form via PEG
linkage as described previously [23]. In the absence of VEGF, there
was not any significant angiogenic response (Supplementary
Figure 3). VEGF released from non-degradable hydrogels lacking
the MMP-sensitive peptide sequence promoted neovascularization
underneath, but not in the hydrogels as shownby the vessels in red
in the depth profile graph (Fig. 4A–D). In contrast, MMP-sensitive
PEG hydrogels with 10% polymer weight had significant portions of
the scaffold degraded after 7 days, and robust neovascularization
was observed (Fig. 4E–H). The newly formed vessels infiltrated into
the core of hydrogels as shown in green in the depth profile graph
(Fig. 4H). Dextran-Texas red (MW 70 KDa) was injected intrave-
nously in the host mice to delineate functionality of the blood
vessels formed in the hydrogels. Fig. 5C shows that indeed the
J.J. Moon et al. / Biomaterials 31 (2010) 3840–38473846

blood vessels in the hydrogels were functional and perfused with
the host’s circulatory system. More rigid hydrogels with 15% poly-
mer weight showed the initial stages of vessel penetration into the
hydrogels (Fig. 4L arrow); however, cells failed to invade into the
core of hydrogels (Fig. 4I–L). These data in corroboration with the in
vitro results indicate that degradation of hydrogels and the ensuing
formation of functional blood vessels can be controlled by provision
of appropriate biophysical and biochemical cues in the hydrogels.
4. Conclusions
This work aimed to combine cellular, biochemical, and biome-
chanical cues to promote neovascularization in synthetic biomate-
rials. By providing appropriate cellular and molecular components
in a microenvironment that mimics the physiological landscape of
angiogenesis, we have been able to recapitulate and promote blood
vessel formation both in vitro and in vivo in completely synthetic
biomaterials. These results suggest applications of these systems for
angiogenesis and anti-angiogenesis studies including its develop-
ment as a diagnostic platform for screening angiogenic and anti-
angiogenic compounds as well as clinical applications in tissue
engineering and regenerative medicine.
Acknowledgements
This work was supported by grants from NIH and NSF, and an
NSF Graduate Student Fellowship (JES). The authors would like to
thank Iris Kim, Melissa Scott, and Tegy Vadakkan for technical
support.
Appendix. Supplementary data
Supplementary data associated with this article can be found in
the online version at doi:10.1016/j.biomaterials.2010.01.104.
References
[1] Langer R, Vacanti JP. Tissue engineering. Science 1993;260:920–6.
[2] Gentzkow GD, Iwasaki SD, Hershon KS, Mengel M, Prendergast JJ, Ricotta JJ,
et al. Use of dermagraft, a cultured human dermis, to treat diabetic foot ulcers.
Diabetes Care 1996;19:350–4.
[3] Atala A, Bauer SB, Soker S, Yoo JJ, Retik AB. Tissue-engineered autologous
bladders for patients needing cystoplasty. Lancet 2006;367:1241–6.
[4] Temenoff JS, Mikos AG. Review: tissue engineering for regeneration of artic-
ular cartilage. Biomaterials 2000;21:431–40.
[5] Levenberg S, Rouwkema J, Macdonald M, Garfein ES, Kohane DS,
Darland DC, et al. Engineering vascularized skeletal muscle tissue. Nat
Biotechnol 2005;23:879–84.
[6] Folkman J, DAmore PA. Blood vessel formation: what is its molecular basis?
Cell 1996;87:1153–5.
[7] Jain RK. Molecular regulation of vessel maturation. Nat Med 2003;9:685–93.
[8] Bell SE, Mavila A, Salazar R, Bayless KJ, Kanagala S, Maxwell SA, et al. Differ-
ential gene expression during capillary morphogenesis in 3D collagen
matrices: regulated expression of genes involved in basement membrane
matrix assembly, cell cycle progression, cellular differentiation and G-protein
signaling. J Cell Sci 2001;114:2755–73.
[9] Davis GE, Senger DR. Endothelial extracellular matrix: biosynthesis, remod-
eling, and functions during vascular morphogenesis and neovessel stabiliza-
tion. Circ Res 2005;97:1093–107.
[10] West JL, Hubbell JA. Polymeric biomaterials with degradation sites for prote-
ases involved in cell migration. Macromolecules 1999;32:241–4.
[11] Lutolf MP, Lauer-Fields JL, Schmoekel HG, Metters AT, Weber FE, Fields GB,
et al. Synthetic matrix metalloproteinase-sensitive hydrogels for the conduc-
tion of tissue regeneration: engineering cell-invasion characteristics. Proc Natl
Acad Sci U S A 2003;100:5413–8.
[12] Zisch AH, Lutolf MP, Ehrbar M, Raeber GP, Rizzi SC, Davies N, et al. Cell-
demanded release of VEGF from synthetic, biointeractive cell ingrowth
matrices for vascularized tissue growth. FASEB J 2003;17:2260–2.
[13] Mann BK, Gobin AS, Tsai AT, Schmedlen RH, West JL. Smooth muscle cell
growth in photopolymerized hydrogels with cell adhesive and proteolytically
degradable domains: synthetic ECM analogs for tissue engineering. Bioma-
terials 2001;22:3045–51.
[14] Lee SH, Moon JJ, West JL. Three-dimensional micropatterning of bioactive
hydrogels via two-photon laser scanning photolithography for guided 3D cell
migration. Biomaterials 2008;29:2962–8.
[15] Lutolf MP, Raeber GP, Zisch AH, Tirelli N, Hubbell JA. Cell-responsive synthetic
hydrogels. Adv Mater 2003;15:888–92.
[16] DeForest CA, Polizzotti BD, Anseth KS. Sequential click reactions for synthe-
sizing and patterning three-dimensional cell microenvironments. Nat Mater
2009;8:659–64.
[17] Canal T, Peppas NA. Correlation between mesh size and equilibrium degree of
swelling of polymeric networks. J Biomed Mater Res 1989;23:1183–93.
[18] Bikram M, Fouletier-Dilling C, Hipp JA, Gannon F, Davis AR, Olmsted-Davis EA,
et al. Endochondral bone formation from hydrogel carriers loaded with BMP2-
transduced cells. Ann Biomed Eng 2007;35:796–807.
[19] Rogers MS, Birsner AE, D’Amato RJ. The mouse cornea micropocket angio-
genesis assay. Nat Protocols 2007;2:2545–50.
[20] Poche RA, Larina IV, Scott ML, Saik JE, West JL, Dickinson ME. The
Flk1-myr::mCherry mouse as a useful reporter to characterize multiple
aspects of ocular blood vessel development and disease. Dev Dyn 2009;
238:2318–26.
[21] Lee SH, Miller JS, Moon JJ, West JL. Proteolytically degradable hydrogels with
a fluorogenic substrate for studies of cellular proteolytic activity and migra-
tion. Biotechnol Prog 2005;21:1736–41.
[22] Barrett AJ, Rawlings ND, Woessner JJF. Handbook of proteolytic enzymes. San
Diego: Academic Press; 1998.
[23] Leslie-Barbick JE, Moon JJ, West JL. Covalently-immobilized vascular endo-
thelial growth factor promotes endothelial cell tubulogenesis in poly(ethylene
glycol) diacrylate hydrogels. J Biomat Sci 2009;20:1763–79.
[24] Hirschi KK, Rohovsky SA, D’Amore PA. PDGF, TGF-beta, and heterotypic cell-
cell interactions mediate endothelial cell-induced recruitment of 10T1/2
cells and their differentiation to a smooth muscle fate. J Cell Biol
1998;141:805–14.
[25] Koike N, Fukumura D, Gralla O, Au P, Schechner JS, Jain RK. Creation of long-
lasting blood vessels. Nature 2004;428:138–9.
[26] Wang ZZ, Au P, Chen T, Shao Y, Daheron LM, Bai H, et al. Endothelial cells
derived from human embryonic stem cells form durable blood vessels in vivo.
Nat Biotechnol 2007;25:317–8.
[27] Mahoney MJ, Anseth KS. Three-dimensional growth and function of neural
tissue in degradable polyethylene glycol hydrogels. Biomaterials 2006;27:
2265–74.
[28] Williams CG, Malik AN, Kim TK, Manson PN, Elisseeff JH. Variable cyto-
compatibility of six cell lines with photoinitiators used for polymerizing
hydrogels and cell encapsulation. Biomaterials 2005;26:1211–8.
[29] Kamei M, Saunders WB, Bayless KJ, Dye L, Davis GE, Weinstein BM.
Endothelial tubes assemble from intracellular vacuoles in vivo. Nature
2006;442:453–6.
[30] Hirschi KK, Rohovsky SA, Beck LH, Smith SR, D’Amore PA. Endothelial cells
modulate the proliferation of mural cell precursors via platelet-d erived
growth factor-BB and heterotypic cell contact. Circ Res 1999;84:2 98–305.
[31] Schlingemann RO, Rietveld FJ, Kwaspen F, van de Kerkhof PC, de Waal RM,
Ruiter DJ. Differential expression of markers for endothelial cells, pericytes,
and basal lamina in the microvasculature of tumors and granulation tissue.
Am J Pathol 1991;138:1335–47.
[32] Beltramo E, Buttiglieri S, Pomero F, Allione A, D’Alu F, Ponte E, et al. A study of
capillary pericyte viability on extracellular matrix produced by endothelial
cells in high glucose. Diabetologia 2003;46:409–15.
[33] Mancuso MR, Davis R, Norberg SM, O’Brien S, Sennino B, Nakahara T, et al.
Rapid vascular regrowth in tumors after reversal of VEGF inhibition. J Clin
Invest 2006;116:2610–21.
J.J. Moon et al. / Biomaterials 31 (2010) 3840–3847 3847