Communication
A Flexible Approach to the Preparation ofPolymer Scaffolds for TissueEngineering
Poonam Borgaonkar, Sachin Sharma, Ming Chen, Sankha Bhowmick,Daniel F. Schmidt*
Porous polyester thermoset xerogels have been produced via sol–gel chemistry as a first step inthe development of sol–gel derived tissue engineering scaffolds templated by replica moldingand/or salt leaching. The pore structure of these untemplated thermosets is tunable and can bealtered independent of or in tandem withalterations in composition. Cytocompatibilitystudies on these xerogels imply the effects ofboth pore size and materials chemistry, withfully aliphatic polyesters with large pore struc-tures allowing the growth of mammalian cells.To the best of our knowledge, this representsthe first report examining the preparation andpotential of sol–gel derived porous polymerxerogels as tissue engineering scaffolds.
Introduction
Tissue engineering combines cell biology, materials
chemistry, and processing to recreate viable tissue that
restore and maintain the function of the body. Three
dimensional biodegradable porous bodies referred to as
‘‘scaffolds’’ are often created for this purpose. Typically, an
P. Borgaonkar, S. Sharma, D. F. SchmidtDepartment of Plastics Engineering, University of MassachusettsLowell, 1 University Avenue, Lowell, MA 01854 USAFax: þ1 (978) 458 4141;E-mail: [email protected]. Chen, S. BhowmickDepartment of Mechanical Engineering/Bioengineering andBiotechnology Program, University of Massachusetts Dartmouth,285 Old Westport Road, North Dartmouth, MA 02747 USA
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appropriate material is chosen and attempts are made to
produce a porous, cell-friendly scaffold optimized for a
specific application. Much has been done, but the state of
the art is far from perfect, and there is much more to do in
order to realize the potential of tissue engineering.[1,2]
Specifically, there is a critical need for appropriate scaffolds
to serve as the base for engineered or regenerated
tissues.[3] At the least, such scaffolds must possess a
significant amount of porosity (>30%), a large pore size (on
the order of tens or hundreds of microns), and a continuous
pore structure in order for cell infiltration. The ideal
scaffold is also mechanically robust, biodegradable,
non-toxic, immunocompatible, and with cell-directing
surface chemistry,[4] and topology/topography/surface
mechanics.[5] It will also act as drug-delivery system for
agents whose activity must be preserved during materials
preparation.[6]
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P. Borgaonkar, S. Sharma, M. Chen, S. Bhowmick, D. F. Schmidt
202
One criterion not mentioned above is the need for a
flexible preparation technique. The body consists of hund-
reds of different types of tissues; no one material will fit the
bill in every tissue engineering application. It is, therefore,
critical that we have a means of rapidly developing new
materials as needs indicate. Likewise, the ability to prepare a
wide range of porous materials whose composition and pore
structures may be independently varied would be an
invaluable tool for scientists attempting to understand
the effects of such variations. It is with these aims that we
have approached the subject, and have selected organic
sol–gel chemistry as a flexible means of scaffold synthesis.
While sol–gel chemistry already sees application in the
field of tissue engineering, most of the efforts have focused
on the development of hydrogel-based systems[7,8] or the
use of inorganic sol–gel chemistry to prepare porous
bioceramics.[9,10] We report on the development of organic
sol–gel chemistry as a means of preparing tissue
engineering scaffolds with structures and chemistries
appropriate for colonization by mammalian cells. While a
single publication has been found describing the prepara-
tion of polyurethane hydrogels via organic sol–gel chemis-
try,[11] we believe this to be the first report describing the
Figure 1. Structure of hyperbranched polyester polyol Boltorn H40. Thyperbranched/dendritic polyester polyol based on a pentaerthyritol (PTheoretically, Mn ¼ 7316 g�mol�1, with 64 primary OH groups pe2800 g�mol�1 (Mw=Mn ¼ 1.8) with �24–25 OH groups per molecule.
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preparation of polymer xerogels for tissue engineering
applications.
The developed systems exhibit rapid and efficient
sol–gel transitions to form open, fully interconnected
pore structures that survive ambient drying to form robust
xerogels. Step polymerization and network formation can
be made to favor gelation over precipitation, while resi-
dual reactive groups may be used to add functionality to
the material. We address the issue of materials selection by
taking a library approach, allowing materials with arbi-
trary properties (hard vs. soft, rapidly vs. slowly degrading,
etc.) to be rapidly prepared by selecting appropriate
components and reaction systems. Variations in reaction
media and conditions allow tuning of the phase separation
conditions and pore structure and accommodation of
active agents (metal precursors, dyes, biomolecules, living
cells, etc.). Additional templating agents (colloids, surfac-
tant phases) may be introduced, while porous inorganic
materials may be formed via pyrolysis or calcination. The
underlying principles at work are simple and versatile,
giving a single synthetic methodology that can address the
needs of a broad range of applications for specific, highly
functional porous materials.
he idealized structure of Boltorn H40, a pseudo-fourth generationE) core reacted with 2,2-bis(hydroxymethyl)propanoic acid (bis-MPA).
r molecule. From supplier data, typical characteristics are Mn ¼
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A Flexible Approach to the Preparation of . . .
Boltorn H40
OH
OH
OHOH
HO
HO
HO
HO OH
Boltorn H40
OH
OH
OHOH
HO
HO
HO
HO OH
++R
O O
Cl Cl
Boltorn H40
OH
O R
O O
O
OHOH
HO
HO
HO
HO OH
Boltorn H40
OH
OH
OHOH
HO
HO
HO OH
-HCl
-HCl
Figure 2. Sol–gel reaction scheme. Schematic of the reaction that forms the basis for the organic sol–gel process described in thiscommunication (triethylamine catalyst not shown). In the case of Samples 1 and 3, terephthaloyl chloride was used and R¼C6H4 (p-phenyl).In the case of Sample 2, adipoyl chloride was used and R¼C4H8 (n-butyl).
In addition to the well-known and readily applicable
approach of salt-leaching as a means of controlling
macroscale porosity,[12] we are developing the means to
directly replicate natural porous bodies using the sol–gel
techniques reported in this communication. Past work has
shown that natural cytocompatible porous bodies (bone,
coral, sponge, etc.) can be infiltrated with wax, then
dissolved to leave the wax ‘‘negative’’, after which a
polymerization reaction may be carried out within the
porous wax to form a porous polymer replica.[13,14] Future
reports will cover these efforts in detail.
In this communication, we focus on the development of
sol–gel systems for use in the aforementioned replica
molding scheme. A variety of systems have been studied
based primarily on the formation of polyester and
polyurethane networks in solution. Of these, three have
been selected on the basis of rapid reaction, robust
materials properties and reasonable homogeneity, and
their structure and cytocompatibility reported.
Experimental Part
Materials
Commercial hyperbranched aliphatic polyester Boltorn H40
(Perstorp Corporation; Mw ¼ 5 100 g �mol�1, Mw=Mn ¼1.8, hydro-
xyl number¼ 470–500 mg KOH � g�1; see Figure 1), terephthaloyl
chloride (99þ%, Acros Organics), adipoyl chloride (98%, Aldrich),
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triethylamine (99.5%, Sigma–Aldrich), and tetrahydrofuran (ACS
reagent, �99.0%, contains 250 ppm BHT as inhibitor, Sigma–
Aldrich) were used as received.
Scaffold Preparation
Sample 1: 20% w/v (total solids weight per total solution volume,
20 g per 100 ml in this case) solutions of terephthaloyl chloride in
THF and Boltorn H40 in THF were combined in the presence of
triethylamine (TEA) in a poly(propylene) vial, which was then
sealed. The overall molar ratio in the final mixture was 1:1:0.25
OH/COCl/TEA, with the catalyst concentration chosen to induce
the rapid formation of a homogeneous, colorless, opaque gel. The
gel was allowed to stand for 24 h at room temperature before
being dried in a vacuum oven at 60 8C for 48 h.
Sample 2: identical to Sample 1, except that adipoyl chloride
was used instead of terephthaloyl chloride. The overall molar ratio
in the final mixture was 1:1:0.5.
Sample 3: identical to Sample 1, except that the total solid
content of the precursor solution was 15% w/v.
Materials analyses were performed without further treatment,
with the exception of the cell culture studies. In this case, the dried
xerogels were immersed in boiling water for 1 h to ensure the
deactivation and/or removal of any remaining monomer or catalyst
and to reduce the chances of contamination by microorganisms.
Scanning Electron Microscopy (SEM)
For the purpose of characterizing the pore structures of the various
xerogels, a JEOL JSM-7401F field emission scanning electron
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P. Borgaonkar, S. Sharma, M. Chen, S. Bhowmick, D. F. Schmidt
Figure 3. SEM images of polyester xerogels. Representative SEM images of Sample 1 (a), Sample 2 (b), and Sample 3 (c), all taken at amagnification of 1,000� (size bar corresponds to 10 mm) using uncoated samples imaged at a low accelerating voltage (1.0 kV) to minimizecharging.
204
microscope (FE-SEM) was used. Dried, uncoated samples were
imaged at an accelerating voltage of 1.0 kV to minimize charging.
For the purpose of imaging fibroblasts, samples were fixed in
4% glutaraldehyde (Aldrich) for 1 h at room temperature, rinsed
with distilled water (Sigma), and dehydrated in increasing
concentrations of ethanol (Aldrich). Dehydrated samples were
dried overnight in a desiccator, sputter coated with gold, and
observed using a JEOL M-5610 SEM with an accelerating voltage of
20 kV.
Mercury Intrusion Porosimetry
Porosities and pore size were determined using a PoreMaster 33
mercury intrusion porosimeter (Quantachrome Instruments). The
maximum intrusion pressure was set at 50 psi (corresponding to a
minimum pore size of �4.26 mm), while mercury surface tension
and intrusion and extrusion contact angles were set at 480
erg � cm�2, 1408, and 1408, respectively. The stem volume used for
all the tests was 0.5 cm3. Samples with masses of �50–100 mg
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were cut from the xerogels following drying, weighed using an
analytical balance, then analyzed. One analysis was performed for
each sample, except Sample 2, which was analyzed twice to
confirm the reproducibility of the measurement.
Mercury intrusion is one of the most commonly applied
porosimetric methods, thanks to the ease with which the
measurement is performed and the wide range of pore sizes
detected, and operation based on a very simple principle: the
intrusion of a non-wetting fluid into a porous body requires
greater applied pressure as the pore size decreases. This is
described mathematically by Washburn’s equation, Equation (1):
Pc ¼2g
rcos u (1)
where Pc is the capillary pressure, g the surface tension of the
liquid within the capillary, r the radius of the capillary, and u the
contact angle made by the liquid with the walls of the capillary.
Mercury is the most commonly used intrusion liquid because it
displays consistent non-wetting behavior (u�1408) on almost any
surface, the exceptions being compounds with which it may react
DOI: 10.1002/mabi.200600212
A Flexible Approach to the Preparation of . . .
(most notably gold, zinc, and selected other metals, sulfur, and
sulfur-rich compounds). While the use of the term ‘‘capillary’’
implies regular cylindrical pores, it is understood that real pores
have more complex geometries. The underlying physics remains
the same, however, in that it is always harder to infiltrate a
smaller pore than a larger pore, allowing the consistent derivation
of pore size distributions by monitoring mercury intrusion into
the sample as a function of pressure. The volume of mercury
intruded into the sample is monitored versus pressure and
converted into a plot of cumulative pore volume versus pore size.
The differential pore volume is then determined based on the
instantaneous slope (i.e., the first derivative) of the cumula-
tive intrusion volume versus pore size curve to give the familiar
differential pore size distribution, where pore volumes given at a
particular pore size are not cumulative but correspond to the
volume of pores of that size alone.
Cell Culture Studies
NIH 3T3 mouse fibroblast cells were cultured on the xerogels in
culture medium containing Dulbecco’s Modified Eagle’s Medium
(DMEM), 10% Bovine Calf Serum (BCS) and 1% penicillin/
streptomycin. The cell number was indirectly measured using
[3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-
sulfophenyl)-2H-tetrazolium] (MTS), a compound that is biore-
duced by cells into a colored formazan product that is soluble in
the tissue culture medium. This conversion is accomplished by
0
1
2
3
4
5
6
7
8
10101
Pore Si
Dif
fere
nti
al P
ore
Vo
lum
e (-
dV
/dlo
gD
, mL
/g)
Figure 4. Pore size distributions of polyester xerogels. Pore size distintrusion porosimetry; Samples 1 and 3 are based on Boltorn H40þSample 2 is based on Boltorn H40þ adipoyl chloride (20% w/v solid
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NADPH or NADH produced by dehydrogenase enzymes in
metabolically active cells, allowing for detection of living cells
by monitoring of the absorbance at 490 nm. A CellTiter 96
Aqueous One Solution Cell Proliferation Assay (Promega, Madison,
WI) was used to measure the cellular activity. On day 3, each
sample was put in a fresh well and rinsed twice with
phosphate-buffered saline (PBS) followed by incubation with
20% MTS reagent in a total of 1 ml of serum free culture medium
for 1 h at 37 8C and 10% CO2. Then, five 100 ml aliquots from each
well were pipetted into five wells of a 96 well plate. The 96 well
plates were placed into a Vmax microplate reader (Molecular
Devices, Sunnyvale, CA), with the absorbance of the contents of
each well at 490 nm measured to indicate metabolically active cell
number.
Results and Discussion
A schematic representation of the structure of the Boltorn
H40 hyperbranched polyester polyol is shown in Figure 1.
Demonstrating its biocompatibility and suitably for
medical applications, this aliphatic polyester has already
attracted interest as a precursor for antiviral drugs[15] and
as a basis for dental resins.[16] Figure 2 shows the step of
polymerization reaction by which gel formation occurs. As
with inorganic sol–gel chemistry, the reactions used are
straightforward and reaction speed is readily varied from
1000010000
ze (µm)
Sample 1
Sample 2
Sample 3
ributions of selected polyester xerogels as determined by mercuryterephthaloyl chloride (20 and 15% w/v solids, respectively), while
s). All samples were prepared with a 1:1 OH/COCl molar ratio.
www.mbs-journal.de 205
P. Borgaonkar, S. Sharma, M. Chen, S. Bhowmick, D. F. Schmidt
Figure 5. SEM images of fibroblast cells attached to Sample 2.Representative SEM images of fibroblast cells attached toSample 2 following cell culture studies. Samples were fixed withglutaraldehyde and sputter-coated with gold, with images takenat a magnification of 1,000� using an accelerating voltage of20 kV.
Table 1. Cell culture data for polyester xerogels. Results of MTScell proliferation assays performed on selected polyester xero-gels. As before, Samples 1 and 3 are based on BoltornH40þ terephthaloyl chloride (20 and 15% w/v solids, respect-ively), while Sample 2 is based on Boltorn H40þ adipoyl chloride(20% w/v solids). All the samples were prepared with a 1:1 OH/COCl molar ratio.
Sample Mean absorbance at 490 nm
1 0.1000� 0.0036
2 0.2630� 0.0092
3 0.0906� 0.0055
Background 0.1013� 0.0005
206
seconds to days by adjusting the catalyst concentration.
Still, many organic sol–gel reactions lead either to inhomo-
geneous gelation or precipitation due to early phase
separation, or the formation of dense bodies prone to
significant drying shrinkage due to little or no phase
separation. Careful control of the solvent quality as well as
the solid content is, therefore, critical for the successful
preparation of the macroporous xerogels reported here.
Changes in composition also result in changes in the
structure developed during gel formation, further empha-
sizing the need for proper control of solvation conditions in
order to prepare comparable systems in the untemplated
case. While pure solvents are used in the examples given,
mixed solvent systems can be used to fine-tune the sol–gel
process.
Scanning electron microscopy was performed on the
xerogels with representative images shown in Figure 3.
While porosity is volumetric whereas microscopy gives
images of surfaces or thin sections, these results never-
theless offer a qualitative assessment of pore size and
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morphology. Smaller pores are observed in Sample 1 as
compared to Samples 2 and 3 but while Sample 1 and
Sample 2 display a granular morphology, some elongated,
almost fibrous features are observed in Sample 3,
indicating a morphological shift, thanks to the lower solid
content versus Sample 1.
Figure 4 shows the mercury intrusion pore size distri-
butions of the polyester xerogels. Based on these results,
the porosities and volume median pore sizes for each
sample are as follows: 79 vol.-%/12 mm (Sample 1), 77
vol.-%/27 mm (Sample 2), and 89 vol.-%/41 mm (Sample 3),
assuming rsolid � 1.2 g � cm�3 for Sample 2 and rsolid � 1.4
g � cm�3 for Samples 1 and 3 based on typical densities of
aliphatic and semi-aromatic polyesters, respectively. A
repeat analysis of Sample 2 (not shown) indicates some
variation in pore size (48 mm) but an almost identical level
of porosity (74 vol.-%), indicating a reasonable level of
homogeneity. From these data, it can be seen that the
sol–gel technique allows for the rapid production of
macroporous polyester xerogels produced via ambient
drying. In addition, these results demonstrate that the pore
structure may be tuned by altering the composition and/or
concentration of the precursor solutions, allowing for the
variation of composition with minimal structural varia-
tion or vice-versa.
The results of the cell culture studies are given in Table 1.
Sample 1 and Sample 3 display background level absor-
bances, indicating no cell growth on these scaffolds.
Sample 2 displays an absorbance significantly higher than
background level, however, implying metabolic activity of
3T3 fibroblasts on this scaffold following cell culturing.
This conclusion is supported by SEM images of Sample 2
following cell culture trials (Figure 5), in which numerous
fibroblast cells are seen to have attached to the polyester
xerogel.
From this, we can draw several conclusions. First, the
positive result in Sample 2 indicates a possible combination
of appropriate pore size and materials chemistry in the
Boltorn H40/adipoyl chloride system. Second, the negative
DOI: 10.1002/mabi.200600212
A Flexible Approach to the Preparation of . . .
result in Sample 3, despite having similar pores to those
found in Sample 2, implies that the Boltorn H40/terephtha-
loyl chloride polyester thermoset is incompatible with cell
growth. Accepting this argument, the lack of cell growth in
Sample 1 can be explained by a combination of structural
incompatibility as a result of the very small pore size versus
the other xerogels and materials incompatibility, given an
identical composition to Sample 3.
Conclusion
In this work, we show that porous polyester thermosets
may be produced via an organic sol–gel technique, and
that a range of pore structures may be realized by chang-
ing the composition, solid content, and reaction condi-
tions. The cytocompatibility studies performed on these
xerogels in turn indicate that large pore materials are
appropriate for the growth of mammalian cells, and show
that the replacement of an aliphatic diacid chloride by an
aromatic diacid chloride in otherwise similar polyester
xerogels may degrade cytocompatibility, emphasizing the
importance of materials choice as well as pore structure.
Finally, this work validates the synthesis of fully organic
sol–gel derived xerogels as a useful means to produce
porous polymers with significant cytocompatibility. In
addition to more detailed studies of these and similar
sol–gel systems, we are pursuing replica molding as a
means to ensure the desired pore structure and further
expand the range of scaffolds that may be produced via the
sol–gel technique.
Acknowledgements: We thank Dr. Earl T. Ada (Manager, UMLMaterials Characterization Laboratory) for the SEM images
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presented, as well as Hugh Lippincott (graduate student,Biomedical Engineering and Biotechnology Program, U. Mass.Lowell) and Shruti Shah (graduate student, Biomedical Engineer-ing and Biotechnology Program, U. Mass. Lowell) for assistance inmaterials preparation and analysis. Finally, we thank theNanomanufacturing Centers (NSF NSEC Center for High-RateNanomanufacturing/Nanomanufacturing Center of Excellence) atthe University of Massachusetts at Lowell for general support.
Received: September 25, 2006; Revised: December 1, 2006;Accepted: December 12, 2006; DOI: 10.1002/mabi.200600212
Keywords: biocompatibility; porous polymers; sol–gel chemis-try; surfaces; tissue engineering
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