Biomaterials 22 (2001) 3045}3051
Smooth muscle cell growth in photopolymerized hydrogels with cell
adhesive and proteolytically degradable domains: synthetic ECM
analogs for tissue engineering
Brenda K. Mann, Andrea S. Gobin, Annabel T. Tsai, Rachael H. Schmedlen,
Jennifer L. West*
Department of Bioengineering, Rice University, 6100 Main Street, MS 142, Houston, TX 77005-1892, USA
Received 28 June 2000; accepted 25 January 2001
Abstract
Photopolymerizable polyethylene glycol (PEG) derivatives have been investigated as hydrogel tissue engineering sca!olds. These
materials have been modi"ed with bioactive peptides in order to create materials that mimic some of the properties of the natural
extracellular matrix (ECM). The PEG derivatives with proteolytically degradable peptides in their backbone have been used to form
hydrogels that are degraded by enzymes involved in cell migration, such as collagenase and elastase. Cell adhesive peptides, such as
the peptide RGD, have been grafted into photopolymerized hydrogels to achieve biospeci"c cell adhesion. Cells seeded homogeneously in the hydrogels during photopolymerization remain viable, proliferate, and produce ECM proteins. Cells can also migrate through
hydrogels that contain both proteolytically degradable and cell adhesive peptides. The biological activities of these materials can be
tailored to meet the requirements of a given tissue engineering application by creating a mixture of various bioactive PEG derivatives
prior to photopolymerization. 2001 Elsevier Science Ltd. All rights reserved.
Keywords: Biomimetic polymers; Hydrogels; Photopolymerization; Proteolysis; Cell migration
1. Introduction
The goal of the current project was the development of
synthetic polymers that can mimic several of the properties of the natural extracellular matrix (ECM), such as
biospeci"c cell adhesion, degradation by proteolytic processes involved in cell migration and tissue remodeling,
and the ability to control cellular functions such as ECM
synthesis. These materials are photopolymerizable derivatives of polyethylene glycol (PEG) that form hydrogel
materials. PEG was chosen for this application due to its
hydrophilicity, biocompatibility, and intrinsic resistance
to protein adsorption and cell adhesion [1,2]. This resistance to protein adsorption and cell adhesion makes the
base material essentially a blank slate, devoid of biological interactions, upon which the desired biofunctionality can be built. Previous work has shown that aqueous
* Corresponding author. Tel.: #1-713-348-5955; fax: #1-713-3485877.
E-mail address: [email protected] (J.L. West).
solutions of acrylated PEG derivatives can be photopolymerized in direct contact with cells and tissues without deleterious e!ects [3,4]. For tissue engineering
purposes, cells can be suspended in the aqueous polymer
solution; after photopolymerization, cells will be homogeneously seeded throughout the hydrogel sca!old.
Photopolymerization can be carried out ex vivo or in situ.
These materials can be rendered bioactive by inclusion
of peptides or polysaccharides in the polymer backbone
or by grafting peptides, proteins, or polysaccharides into
the hydrogel network during the photopolymerization
process. In order to create a material that might be
degraded by tissue formation processes, we have chosen
to target degradation to proteolytic enzymes involved in
cell migration and tissue remodeling [5}8]. Proteases,
such as the matrix metalloproteases, are crucial in the cell
migration process, as they allow cells to clear a pathway
through the dense matrix [9]. We have previously shown
that PEG-based hydrogels with peptides in their backbone that are degradation substrates for particular enzymes, such as collagenase or plasmin, can undergo
proteolytic degradation [10]. This degradation scheme
0142-9612/01/$ - see front matter 2001 Elsevier Science Ltd. All rights reserved.
PII: S 0 1 4 2 - 9 6 1 2 ( 0 1 ) 0 0 0 5 1 - 5
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B.K. Mann et al. / Biomaterials 22 (2001) 3045}3051
should match the rate of material resorption to the rate
of tissue formation. In order to achieve biospeci"c cell
adhesion to these materials, we have grafted cell adhesion
peptides into the hydrogel structure in a manner similar
to that used by Hern and Hubbell [11], who demonstrated peptide concentration-dependent adhesion of
"broblasts to the surface of PEG diacrylate hydrogels
grafted with RGD peptides. The PEG hydrogels o!er an
advantage over most other sca!old materials in that they
are intrinsically resistant to cell adhesion [1,2]. This
allows the cell-material interactions to be limited to the
adhesive ligands provided. If cell selective ligands, such as
the REDV peptide that has been shown to be endothelial
cell speci"c [12], are incorporated into a cell non-adhesive sca!old, the resultant sca!old material should allow
adhesion of only the desired cell type. We have also
previously shown that growth factors, such as transforming growth factor-beta1, can be covalently grafted into
these photopolymerized hydrogel sca!olds and maintain
their activity. Cells grown in sca!olds with immobilized
transforming growth factor-beta exhibited dramatically
increased synthesis of collagen and improved mechanical
properties of the resultant engineered tissues, even compared to cells in similar sca!olds but with the growth
factor soluble in the culture media [13]. The hydrogels
used as tissue engineering sca!olds can be formed from
blends of several of these bioactive PEG derivatives in
order to create a matrix substitute with the desired biological activities for a given application.
2. Materials and methods
Chemicals were obtained from Sigma Chemical Co.
(St. Louis, MO) unless otherwise stated.
2.1. Cell maintenance
Human dermal "broblasts (HDFs) and human aortic
smooth muscle cells (HASMCs) were obtained from
Clonetics (San Diego, CA). Rat aortic smooth muscle
cells (RASMCs) were isolated as previously described
[14]. HDFs were maintained on Dulbecco's modi"ed
Eagle medium (DMEM) supplemented with 10% fetal
bovine serum (FBS; BioWhittaker, Walkersville, MD),
2 mM L-glutamine, 500 units penicillin and 100 mg/l
streptomycin. HASMCs and RASMCs were maintained
on minimum essential Eagle medium (MEEM) supplemented with 10% FBS, 2 mM L-glutamine, 500 units
penicillin and 100 mg/l streptomycin. Cells were incubated at 373C in a 5% CO environment.
0.4 mmol/ml acryloyl chloride, and 0.2 mmol/ml
triethylamine in anhydrous dichloromethane and stirring
under argon overnight. The resulting PEG-diacrylate
was then precipitated with ether, "ltered, and dried
in vacuo.
2.3. Preparation of PEG-diacrylate derivatives containing
degradable sequences
The ABA block copolymers of PEG (A) and peptides
that are substrates for proteolytic enzymes (B) were
synthesized. The degradable peptide sequences
were GGLGPAGGK (collagenase-sensitive) or AAAAAAAAAK (elastase-sensitive; Research Genetics, Huntsville, AL). The appropriate peptide was reacted with
acryloyl-PEG-N-hydroxysuccinimide
(acryloyl-PEGNHS, 3400 Da; Shearwater Polymers, Huntsville, AL) in
a 1 : 2 (peptide : PEG) molar ratio in 50 mM TRIS bu!er
(pH 8.5) for 2 h. This adds a PEG-monoacrylate chain to
the N-terminus and to the amine group on the lysine at
the C-terminus of the peptide. The product was then
lyophilized and stored frozen. Gel permeation
chromatography equipped with UV/Vis (260 nm) and
evaporative light scattering detectors (Polymer Laboratories, Amherst, MA) was used to analyze the resulting
PEG-peptide copolymers and PEG standards (Polymer
Laboratories).
2.4. Preparation of acryloyl-PEG-peptide copolymers
Peptides utilized in this study were KQAGDV (Research Genetics) and RGDS (Sigma). Peptides were conjugated to PEG monoacrylate by reacting the peptide
with acryloyl-PEG-NHS (3400 Da) in 50 mM TRIS bu!er
(pH 8.5) for 2 h. The mixture was then lyophilized and
stored frozen. Gel permeation chromatography equipped
with UV/Vis (260 nm) and evaporative light scattering
detectors was used to analyze the resulting copolymers
(Polymer Laboratories).
2.5. Photopolymerization of hydrogels
Hydrogels were prepared by combining 0.2 g/ml
PEG-diacrylate and 0.15 mmol/ml triethanolamine in
10 mM HEPES-bu!ered saline (pH 7.4, HBS). A 40 l/ml
of 2,2-dimethyl-2-phenyl-acetophenone in n-vinylpyrrolidone (600 mg/ml) was then added. The resulting solution was then exposed to UV light (365 nm, 10 mW/cm)
for 20 s to convert the liquid polymer solution to a hydrogel.
2.6. Degradation of hydrogels
2.2. Preparation of PEG-diacrylate
PEG-diacrylate was prepared by combining 0.1 mmol/
ml dry PEG (6000 Da; Fluka, Milwaukee, WI),
Hydrogels were prepared as above using 0.1 g/ml
PEG-diacrylate derivative containing either GGLGP
AGGK or the 9-mer alanine in the polymer backbone.
B.K. Mann et al. / Biomaterials 22 (2001) 3045}3051
100 l of the polymer solution was placed in a diskshaped mold (6.5 mm diameter, 3.1 mm thickness) and
photopolymerized. Hydrogels were placed in HBS with
0.2 mg/ml sodium azide for 24 h at 373C to swell to their
equilibrium state. Hydrogels containing GGLGPAGGK
were then placed in HBS with 0.2 mg/ml sodium azide
and 0, 0.2, or 2 mg/ml collagenase or 2 mg/ml elastase.
Hydrogels containing the 9-mer of alanine were placed in
HBS with 0.2 mg/ml sodium azide and 0, 0.2, or 2 mg/ml
elastase or 2 mg/ml collagenase. The samples were incubated at 373C for the duration of the experiment. Every
2 days, the protease solution was removed, the hydrogel
was weighed, and fresh protease solution was added.
2.7. Migration through scawolds
Hydrogels containing 1.4 mol/ml acryloyl-PEGRGDS with 0.1 g/ml PEG-diacrylate derivative containing GGLGPAGGK were prepared as described above.
A 50 l of the polymer solution was pipetted into a transwell cell culture insert (6.4 mm diameter, 8 m pore,
Becton Dickinson, Franklin Lakes, NJ) and photopolymerized. The resulting hydrogel was 1 mm thick. As
a reference material, 1 mm thick collagen gels with
physical characteristics similar to the PEG hydrogels
were also formed in transwell cell culture inserts.
A 2 mg/ml solution of collagen (Vitrogen 100, Cohesion
Technologies, Palo Alto, CA) was prepared with the "nal
pH 7.4. A 100 l of this solution was pipetted into each
transwell insert. The inserts were incubated at 373C for
60 min to form 1 mm thick gels after contraction. The
inserts were placed in 24-well plates (Becton Dickinson),
and HDFs were seeded on top of the hydrogel layer
(3;10 cells/well). DMEM containing 10% FBS was
added to the insert and well, and the plates were incubated at 373C with 5% CO . After 7 days of culture, cells
were removed from the insert and well with trypsin and
counted using a Coulter counter (Multisizer C 0646,
Coulter Electronics, Hialeah, FL). The migration index
was calculated as the number of cells that had migrated
through the hydrogel and insert membrane divided by
the sum of the cells that had migrated into the well and
those that remained in the insert.
2.8. Cell viability in hydrogels
Hydrogels were formed as described above, except that
the aqueous polymer solution contained 0.4 g/ml PEGdiacrylate and 0.3 mmol/ml triethanolamine. An equal
volume of this was added to a suspension of HASMCs to
create a cell-polymer solution with a cell density of
1;10 cells/ml and a PEG-diacrylate concentration of
0.2 g/ml. Acryloyl-PEG-KQAGDV of 1.4 mol/ml and
the photoinitiator solution were then added. The
KQAGDV peptide is a potent adhesion ligand for SMCs
[15]. A 250 l of the resulting cell-polymer mixture was
3047
placed in a disk-shaped mold (20 mm diameter;2 mm
thick) and photopolymerized as described above. The
hydrogels were then transferred to a 12-well tissue culture plate, and 2 ml of MEEM with 10% FBS was added
to each well. The hydrogels were incubated at 373C in
a 5% CO environment, and media were changed every
3 days. After 1 week of culture, the hydrogels were incubated with serum-free MEEM containing 0.1 M chloromethyl#uorescein diacetate (CMFDA; Molecular
Probes, Eugene, OR) at 373C for 4 h. CMFDA is a non#uorescent #uorescein derivative that is taken up by
viable cells, where the diacetate group is cleaved to form
a #uorescent #uorescein derivative. The 50 m sections of
these hydrogels were taken on a cryostat (HM505E, Carl
Zeiss, Thornwood, NY), and the sections were evaluated
under phase-contrast microscopy and #uorescence
microscopy at 100 X (Axiovert 135, Carl Zeiss) to examine cell viability.
2.9. Proliferation in hydrogels
Hydrogels containing 7 mol/ml acryloyl-PEGKQAGDV were formed as described above for the cell
viability study, except that RASMCs were used. After
7 days of culture, hydrogels were sectioned to a thickness
of 20 m using a cryostat (Carl Zeiss). The sections were
"xed with 10% bu!ered formalin and stained for proliferating cell nuclear antigen (PCNA). Cells were permeabilized with methanol, incubated with mouse antiPCNA IgG (Dako, Carpinteria, CA), incubated with
HRP-conjugated anti-mouse IgG, incubated with 3amino-9-ethylcarbazole (AEC) substrate-chromagen,
and "nally counterstained with Mayer's hematoxylin.
The numbers of proliferating and non-proliferating cells
in each section were counted under light microscopy
(Carl Zeiss).
2.10. DNA and hydroxyproline measurement
in hydrogels
Hydrogels containing 7 mol/ml acryloyl-PEGKQAGDV and homogeneously seeded with RASMCs
were formed as described above for the proliferation
study. After 3 and 7 days of culture, hydrogels were
removed from the culture media, weighed, and digested
with 1 ml 0.1 N NaOH overnight at 373C. Digested hydrogels were then neutralized with 1 ml 0.1 N HCl. DNA
content of the digested, neutralized hydrogels was determined using a #uorescent DNA binding dye, Hoechst
33258 (Molecular Probes). Fluorescence of the samples
was determined on a #uorometer (VersaFluor, Bio-Rad
Laboratories, Hercules, CA) with excitation "lter at
360 nm and emission "lter at 460 nm, and compared to
#uorescence of calf thymus DNA standards. Hydroxyproline concentration was determined by oxidation
with chloramine T (ICN Biomedicals, Aurora, OH) and
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B.K. Mann et al. / Biomaterials 22 (2001) 3045}3051
development with p-dimethylaminobenzaldehyde (ICN
Biomedicals) [16]. Hydroxyproline is a marker for
collagen production [16].
2.11. Matrix synthesis in degradable hydrogels
Hydrogels were made as described above containing
2.8 mol/ml acryloyl-PEG-KQAGDV and homogeneously seeded with HASMCs at a "nal concentration of
1.4;10 cells/ml. The hydrogels were made using either
PEG-diacrylate or a PEG-diacrylate derivative containing the 9-mer of alanine. After 7 and 26 days of culture,
hydrogels were digested and assayed for DNA and
hydroxyproline as described above. Additionally, a piece
of each gel was removed at 7 days and 20 m sections
were taken on a cryostat. These sections were then
stained with Biebrich's Scarlet Acid Fuschin to stain the
cytoplasm and were evaluated under light microscopy at
400;.
Fig. 1. Degradation of GGLGPAGGK-derivatized PEG hydrogels in
solutions containing collagenase. (䢇) 2 mg/ml collagenase; (*)
0.2 mg/ml collagenase; (䉱) no collagenase.
2.12. Statistical analysis
Data sets were compared using two-tailed, unpaired
t-tests. p-Values less than 0.05 were considered to be
signi"cant.
3. Results
3.1. Proteolytic degradation of hydrogel scawolds
In order to link sca!old degradation to tissue formation, sca!olds were synthesized that could be degraded
by proteolytic enzymes secreted by cells during
migration. These sca!olds were made by incorporating
proteolytically degradable peptide sequences into the
backbone of a PEG diacrylate derivative as an ABA
block copolymer (A"PEG monoacrylate, B"peptide).
The peptide sequences examined were LGPA, targeted
for degradation by collagenase [5,6], and a 9-mer of
alanine, targeted for degradation by elastase [7,8].
Degradation of the hydrogels by their respective enzymes
were examined by placing the hydrogels in a solution
containing the enzyme and tracking their mass over time.
Degradation of LGPA-derivatized hydrogels in solutions
containing collagenase is shown in Fig. 1. The hydrogels
initially swelled as some of the LGPA sequences were
cleaved, loosening the hydrogel network, and more water
was able to enter the hydrogels. The hydrogels then lost
mass as more sequences were degraded. As the amount of
collagenase in the solution increased, so did the rate of
hydrogel degradation. Additionally, hydrogels placed in
solutions without collagenase did not degrade during the
timecourse of this experiment, indicating that the degradation of the hydrogels was due to the presence of the
action of the collagenase. Further, LGPA-containing
Fig. 2. Degradation of AAAAAAAAAK-derivatized PEG hydrogels in
solutions containing elastase. (䢇) 2 mg/ml elastase; (*) 0.2 mg/ml elastase; (䉱) no elastase.
hydrogels placed in solutions containing elastase did not
degrade, indicating that degradation of the LGPA is
speci"c to collagenase. Hydrogels containing the 9-mer
of alanine responded to solutions containing elastase in
a similar manner (Fig. 2). That is, the hydrogels degraded
in a dose-dependent fashion to the amount of elastase
present, and did not degrade when placed in solutions
without proteolytic enzymes or in solutions containing
collagenase.
3.2. Cell migration through biomimetic hydrogels
A modi"ed Boyden chamber assay was used to investigate cell migration through the synthetic ECM analogs.
Hydrogels containing the proteolytically degradable
sequence GGLGPAGGK and the adhesive sequence
B.K. Mann et al. / Biomaterials 22 (2001) 3045}3051
RGDS were formed in transwell cell culture inserts, and
migration of "broblasts through the hydrogel was examined and compared to migration through collagen gels of
the same thickness. Similar numbers of cells migrated
through the synthetic collagen-mimetic hydrogel and the
collagen gel (migration index of 6.96$0.41 and
9.78$2.60, respectively, p'0.05).
3.3. Cell viability in hydrogels
HASMCs were homogeneously seeded into hydrogels
containing the adhesion ligand KQAGDV and were
examined after 24 h and 1 week of culture to ensure that
the cells survive photopolymerization and remain viable
in the hydrogels after extended culture periods. Viable
cells, indicated by #uorescence upon CMFDA staining,
were evident 24 h after photopolymerization and after
1 week of culture. Approximately 95% viability was
observed in the hydrogels at both time points. Viable
cells were found uniformly distributed throughout the
hydrogel with no di!erence in viability at the periphery
versus the center of the hydrogel.
3.4. Cell proliferation in hydrogels
RASMCs were homogeneously seeded into hydrogels
containing KQAGDV to examine cell proliferation within the hydrogels. After 7 days of culture, 20 m sections of
the hydrogels were taken, and the cells were stained for
PCNA and counterstained with hematoxylin. Proliferating and non-proliferating cells were observed in all of the
hydrogel sca!olds (Fig. 3), with 47.3$3.1% of the cells
within the hydrogel proliferating.
3049
ly seeded within hydrogels containing KQAGDV. After
in vitro culture, the hydrogels were digested, and the
amounts of DNA and hydroxyproline were determined.
The DNA content was used as a marker for total cell
density. Since hydroxyproline is a marker for collagen
[16], it is an indication of how much extracellular matrix
has been produced by the cells. DNA increased 26% in
the hydrogels from day 3 to day 7 (2717.7$84.6 and
3415.2$64.8 ng DNA/g gel, respectively) and collagen
was produced by cells in the hydrogels by day 7
(2807.3$751.4 ng hydroxyproline/g gel).
3.6. Matrix synthesis in degradable hydrogels
We examined DNA content and matrix protein production in proteolytically degradable versus non-degradable PEG hydrogels homogeneously seeded with
HASMCs. The degradable hydrogels contained a polyalanine sequence which is elastase-sensitive. SMCs have
previously been shown to secrete elastase during migration [17]. As shown in Fig. 4, more hydroxyproline was
produced at both 7 and 26 days in the degradable hydrogels than in the non-degradable hydrogels. Additionally,
HASMCs seeded in the degradable hydrogels can be seen
extending processes at 7 days while those in the nondegradable hydrogels do not (Fig. 5).
4. Discussion
Materials used as sca!olds to support and guide tissue
formation must meet certain basic criteria; they must be
3.5. DNA and hydroxyproline production in hydrogels
To examine DNA content and matrix protein production within the hydrogels, RASMCs were homogeneous-
Fig. 3. RASMCs growing in KQAGDV-containing hydrogels stained
for PCNA and counterstained with hematoxylin after 1 week of culture.
Proliferating cells are red; non-proliferating cells are blue.
Fig. 4. Hydroxyproline production by HASMCs growing in nondegradable PEG-diacrylate hydrogels (PEG) and in AAAAAAAAAderivatized PEG hydrogels (9AK). Solid bars: 7 days; hashed bars: 26
days.
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B.K. Mann et al. / Biomaterials 22 (2001) 3045}3051
Fig. 5. HASMCs growing in PEG hydrogels and stained with Biebrich's Scarlet Acid Fuschin. A: in AAAAAAAAA-derivatized PEG
hydrogels; B: in PEG-diacrylate hydrogels.
biocompatible, must support cell growth, must be bioresorbable, must provide the mechanical support necessary to maintain the tissue form, and should be easily
processed. A number of synthetic biodegradable polymers, such as poly(lactic-co-glycolic acid), largely meet
these requirements [18}20]. Ideally, however, a sca!old
material should also have a number of biological functions so that one might elicit speci"c cellular functions,
dictate cellular interactions, or actively encourage ingrowth of cells from the surrounding tissue. The natural
tissue sca!old, the extracellular matrix (ECM), exerts
precise control over cellular functions, interactions, and
growth. The ECM components, such as collagen materials [21] and decellularized tissues [22,23], have been
investigated for tissue engineering sca!olds in order to
take advantage of their inherent biological functionality.
While there are numerous advantages to these types of
materials, their use has been limited by their range of
mechanical properties, processing conditions, reproducibility, and cost. An alternative approach has been the
modi"cation of synthetic sca!olds with bioactive moieties [24}28]. For example, poly(lactide-co-lysine) polymers have been functionalized with RGD peptides to
enhance cell adhesion [29,30]. This imparts at least a limited degree of biofunctionality to a synthetic sca!old.
In the current study, we have described a photopolymerizable hydrogel system that allows one to tailor
the biological activity of the hydrogel sca!old, including
cell adhesive characteristics, susceptibility to proteolytic
remodeling, and presentation of growth factors, to meet
the requirements of a given tissue engineering or regeneration application. These materials interact with cells in
a manner similar to the natural ECM and perform many
functions of the natural ECM. Bioactive signals can also
be incorporated to control aspects of cell behavior such
as the rate of proliferation, the amount of extracellular
matrix synthesized, and the speed and directionality of
cell migration. For example, we have previously demonstrated that the covalent incorporation of TGF-beta1
into photopolymerized hydrogel sca!olds signi"cantly
increases collagen production by SMCs within the scaffold, resulting in an improvement in the mechanical
properties of the engineered tissue [13].
We have demonstrated that cells can be homogeneously incorporated into these hydrogel sca!olds during
photopolymerization without a loss in cell viability. The
cell-polymer suspensions can be photopolymerized
ex vivo or in situ and can be made into complex shapes
using transparent molds. For example, multi-layered
tubes can be fabricated for development of tissue engineered vascular grafts. The incorporated cells remain viable (as shown with CMFDA staining), undergo
proliferation (as demonstrated by both increases in DNA
content in the sca!olds and PCNA immunohistochemistry), and secrete matrix proteins such as collagen (as
indicated by increases in hydroxyproline) to begin tissue
formation. Similar materials based on PEG dimethacrylate but lacking bioactive moieties have been investigated for use as sca!olds for chondrocytes [31]. With the
materials described here, we have further demonstrated
that cells can recognize and interact with the bioactive
signals incorporated into the hydrogels. Cell adhesion to
the surface of these materials requires inclusion of cell
adhesion peptides, and the degree of adhesion is dependent on the amount of peptide grafted to the hydrogel
structure [11]. Cells were able to adhere to and migrate
through hydrogels that contained both the appropriate
adhesive and degradable ligands. If either the adhesive or
degradable sequence was omitted from the hydrogel
structure, no cell migration through the material was
observed. This is expected as cells require interaction
with cell adhesive receptors in order to generate forces
required for cell migration and secrete proteolytic enzymes to create pathways for migration through three
dimensional environments [9,32].
Additionally, we have compared aspects of tissue
formation within non-degradable PEG diacrylate hydrogels and proteolytically degradable PEG diacrylate hydrogels that contained an elastase-sensitive polyalanine
sequence in the center of the polymer backbone. It appears that tissue formation is fostered in the degradable
materials, presumably since cells can degrade surrounding hydrogel material to provide space for additional
cells and deposition of a new extracellular matrix to
replace the sca!old. For example, collagen synthesis was
signi"cantly increased in the proteolytically degradable
hydrogels as compared to the non-degradable PEG diacrylate hydrogels. This degradation mechanism is desirable over hydrolytic degradation since resorption of the
synthetic material is caused by cellular activity. As a result, the rate of material degradation should be closely
linked to the rate of tissue formation. To achieve similar
B.K. Mann et al. / Biomaterials 22 (2001) 3045}3051
results with hydrolytically degradable materials would
require a precise understanding of the kinetics of tissue
formation and degradation of the sca!old materials.
In conclusion, the sca!old materials we have described
are able to provide many of the normal signals and
interactions provided to cells by the ECM in tissues. This
may allow greater control over tissue formation and cell
phenotype. Furthermore, these synthetic ECM analogs
may be useful not only in tissue engineering and regeneration applications, but also as a tool to examine the
e!ects of one or a combination of bioactive molecules on
various aspects of cell behavior, such as mechanisms
involved in cell migration.
Acknowledgements
Funding for this work was provided by the NIH Heart,
Lung, and Blood Institute (R01 HL60485) and the Theodore N. Law Foundation.
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