Inverted colloidal crystals as three-dimensional microenvironments forcellular co-cultures
Jungwoo Lee,a Sachin Shanbhagb and Nicholas A. Kotov*abc
Received 24th April 2006, Accepted 4th July 2006
First published as an Advance Article on the web 25th July 2006
DOI: 10.1039/b605797g
Cellular scaffolds made on the basis of inverted colloidal crystals (ICC) provide a unique system
for investigation of cell–cell interactions and their mathematical description due to highly
controllable and ordered 3D geometry. Here, we describe three new steps in the development of
ICC cell scaffolds. First, it was demonstrated that layer-by-layer (LBL) assembly with clay/PDDA
multilayers can be used to modify the surface of ICC scaffolds and to enhance cell adhesion.
Second, a complex cellular system made from adherent and non-adherent cells co-existing was
created. Third, the movement of non-adherent cells inside the scaffold was simulated. It was
found that floating cells are partially entrapped in spherical chambers and spend most of their
time in the close vicinity of the matrix and cells adhering to the walls of the ICC. Using this
approach one can efficiently simulate differentiation niches for different components of
hematopoietic systems, such as T-, B- and stem cells.
Introduction
Colloidal crystals represent an exceptionally dynamic area of
research capitalizing on the unique spatial organization and
diffraction characteristics of sub-micron scale lattices.1–4 These
structures are designed, having in mind, primarily, applica-
tions in optics,5 sensors6 and catalysis.7 Inverted colloidal
crystals (ICC) also open an interesting opportunity for a rather
unexpected, but tremendously important area of science
related to cell communication. We recently introduced the
use of micron-scale ICC systems for three-dimensional (3D)
cell cultures, which mimic the microenvironment of three-
dimensionally organized native tissues.8–10 Unlike other
disordered cell supports, the ICC geometry affords systematic
study of cell signaling in 3D, which has been proven to be
fundamentally essential for proper development of tissues.11–16
Adequate understanding and proper methods of control of
cell signaling are particularly important for stem cell research.
For instance, the rate and direction of the differentiation of
stem cells are strongly affected by their 3D microenvironment
and soluble signaling molecules.17–22 Recent studies have
shown that a 3D culture environment significantly promotes
the efficiency of stem cell differentiation.23,24 Intense cell–cell
and cell–matrix interactions have been distinguished as key
factors that determine the fate of individual cells by serving
as important communication channels.23,25,26 In order to
reproduce the complexity and dynamics of cellular environ-
ments, various scaffold fabrication techniques have been
developed.27–31 However, the geometry of these scaffolds
mainly depends on the process, and usually they have a poorly
ordered or chaotic structure. Recently, rapid prototyping and
3D deposition techniques, assisted by computer-aided design
and complex robotic equipment, were developed to construct
more controlled 3D architectures.27,32 These techniques allow
researchers to design 3D scaffolds with desired properties such
as porosity, interconnectivity and pore size. Nevertheless,
besides being heavily equipment-dependent, they suffer from
limited material selection and inadequate resolution. From a
manufacturing standpoint, the fabrication procedure of ICC
scaffolds is simple and flexible. Any precursor solution capable
of undergoing a liquid-to-solid transition may potentially be
used as a scaffolding material. An ordered structure, with a
high degree of uniformity, can be achieved without the need
for complex computer design programs and facilities.
Beyond that, several unique characteristics of ICC used as
cell scaffolds make them particularly convenient for the use
with stem cell cultures,8–10,33 which can help uncover methods
for successful tissue engineering from them. In this respect,
ICC systems possess high surface areas with a void fraction of
76% and a regularly spaced network of pores which provides a
mechanically strong, well-connected open porous geometry.33
These features enhance cell seeding efficiency, transport of
nutrients and metabolites, and the rapid and uniform
distribution of soluble signaling molecules. The exceptionally
uniform and three-dimensionally ordered structure of ICC
scaffolds enables the development of computational models to
systematically study the effect of signaling molecules, cell–cell
and cell–matrix interactions, and other processes.34 Until now,
only single cell culture studies have been reported for ICC
scaffolds.8–10,33 Considering that this system could be a
convenient discovery tool for research on cell–cell interactions,
achieving the next level of complexity, i.e. the construction of
a system with two or more different cell types co-populating
the ICC matrix, appears to be the most essential step in
this area.
aDepartment of Biomedical Engineering, University of Michigan, 3074H.H. Dow Building, 2300 Hayward Street, Ann Arbor, MI 48109, USAbDepartment of Chemical Engineering, University of Michigan, 3074H.H. Dow Building, 2300 Hayward Street. Ann Arbor, MI 48109, USAcDepartment of Material Science and Engineering. University ofMichigan, 3074 H.H. Dow Building, 2300 Hayward Street, Ann Arbor,MI 48109, USA. E-mail: [email protected]; Fax: +1-734-764-7453;Tel: +1-734-763-8768
PAPER www.rsc.org/materials | Journal of Materials Chemistry
3558 | J. Mater. Chem., 2006, 16, 3558–3564 This journal is � The Royal Society of Chemistry 2006
In this paper, we introduce a model system combining two
types of cells co-existing in an ICC scaffold, which paves the
way for future systematic studies of cell evolution mechanisms.
Since these interactions are of particular importance for the
development of hematopoietic stem cells, the cell cultures
were chosen having in mind the recreation of the 3D
microenvironment of bone marrow and thymus differentiation
niches.26,35,36 The selection of particular model cell cultures
was also aided by the fact that the characteristic geometry of
the ICC scaffolds resembles that of bone marrow and thymus
niches (i.e. stromal cells cover the surface and well intersticed
sinus cavities). Human thymus epithelial cells (Hs202.Th)
and human monocytes (HL-60) were used as anchorage-
dependent feeder cells and suspension cells mimicking pro-
genitors, respectively. Before using hematopoietic stem cells
in our 3D culture system, we tried to use the HL-60 cell line
because it is easier to deal with and has been proven a unique
in-vitro model system for studying the cellular and molecular
events involved in the proliferation and differentiation of
normal and leukemic cells.37 Cell–cell interactions within
ICC scaffolds were evidenced by simplified Brownian
Dynamics (BD) simulations taking advantage of the unique
3D morphology.
Experimental
Colloidal crystal construction
An aqueous suspension of polystyrene (PS) spheres with a
diameter of 100 mm (Duke Scientific, 3 6 104 particles per
milliliter and 1.4% size distribution) was changed with
isopropanol solution before use. A 0.5 ml plastic centrifuga-
tion tube was glued on the plastic dish and the top of the
centrifuge tube was cut and connected with a long Pasteur
glass pipette. The complex unit was attached to the bottom
of a glass beaker, and the glass beaker was placed on an
ultrasonic bath (VWR). Two drops of the solution were
released through a long Pasteur glass pipette at 15 minute
intervals (25 intervals total) under gentle agitation generated
by the ultrasonic bath. To reduce thermal motions of the
spheres, the bath temperature was maintained below 20 uC.
After the dropping was finished, the isopropanol was
evaporated off overnight at 60 uC.
Scaffold fabrication
Prepared colloidal crystals were heat treated at 120 uC for
4 hours, which caused partial melting of the beads’ surface. As
a result, the PS microspheres fused together and the free
standing colloidal crystals were easily extracted from the mold.
As scaffolding materials, a poly(acrylamide) hydrogel com-
posed of a 30 wt% acrylamide (Sigma) precursor containing
5 wt% of N,N-methylenebisacrylamide (NMBA) cross-linker
was used. The precursor was infiltrated into the colloidal
crystal by centrifugation at 5800 rpm for 10 min. An initiator,
1 wt% of potassium peroxide solution, and an accelerator,
N,N,N9,N9-tetramethylethylenediamine (TEMED), were
added. Polymerization occurred in a glass vial. After the
polymerization was complete, the colloidal crystal containing
the hydrogel part was cut out and soaked in tetrahydrofuran
(THF) for 24 hours to remove PS beads. Finally the ICC
hydrogel scaffolds were equilibrated in deionized water.
Fluorescent ICC hydrogel scaffolds were prepared by adding
0.05 wt% of fluorescent monomer, Polyfluor 511 (Polyscience
Inc.), to the hydrogel precursor solution.
Layer-by-layer surface coating
ICC hydrogel scaffold surfaces were coated with the sequential
deposition of positively charged 0.5 wt% poly(diallyldimethyl-
ammonium chloride) (PDDA, Sigma, MW = 200 000) solution
for 15 min, and a negatively charged 0.5 wt% clay platelet
(average 1 nm thick and 70–150 nm in diameter, Southern
Clay Products) dispersion for 15 min. Each adsorption
step was followed by rinsing in deionized water for 15 min,
and all processes were performed under a gentle flow
generated by a stirrer. Cyclic repetition of the polymer
adsorption/rinsing/clay adsorption/rinsing process was carried
out 10 times.38
Mechanical property testing
Compressive moduli of hydrated ICC scaffolds, where the
surface was coated with 10 layers of clay/PDDA, were
measured at a constant strain rate (10 mm s21) using a
mechanical properties tester and a 1.1 lb load cell
(TestReciurces Inc., MN).
Dynamic co-culture
Human thymus cell line Hs202.Th (CRL-7163) and human
premyeloblast cell line HL-60 (CCL-240) were purchased
from ATCC (Manassas, VA). Hs202Th cells were grown in
Dulbecco’s Modified Eagle’s Medium (DMEM) supplemented
with 10% Fetal Bovine Serum (FBS) (GIBCO, CA). HL-60
cells were cultured in Iscove’s Modified Dulbecco’s Medium
(IMDM) containing 20% FBS. The cells were maintained at
37 uC with 5% CO2 and the medium was changed twice a week
until they reached a confluent or desired population on T-75
culture flasks.
Co-culture was carried out in 10 ml rotary cell culture
vessels (RCCS-4D, Synthecon Inc.). Scaffolds were sterilized
by soaking in 70% EtOH for one hour followed by washing
in phosphate buffered saline (PBS) for 15 min twice. 2 6105 Hs202.Th cells were placed in a culture vessel, which
subsequently was filled with the medium. The rotation speed
was set at 12 rpm for the first 12 hours and later it was
decreased to 8 rpm, the normal speed. The medium was
replaced once every three days. On day six, both Hs202.Th and
HL-60 were stained with fluorescent dyes followed by a five
day co-culture period. The HL-60 cells were stained with 5 mM
chloromethyl derivatives fluorescent dye (CMRA, Molecular
Probes) diluted in PBS buffer following the protocol provided
by the vendor. Hs202.Th cells on the scaffold were stained with
carboxyfluorescein diacetate succinimidyl ester (CFDA-SE,
Molecular Probes). The culture medium was replaced with
10 ml of 5 mM CFDA-SE diluted in PBS buffer, and the
culture was incubated at 37 uC for 20 min. After that, the
medium was changed to IMDM supplemented with 20% FBS,
and pre-stained 1 6 106 HL-60 cells were seeded.
This journal is � The Royal Society of Chemistry 2006 J. Mater. Chem., 2006, 16, 3558–3564 | 3559
Cell culture characterization
Scaffolds were carefully removed from the culture vessel
together with the medium and moved to a glass bottom culture
dish (MatTek Corporation). The scaffolds were kept immersed
in the medium during the process. Co-cultured scaffolds were
investigated utilizing a Leica SP2 confocal microscope with
106 and 206 objective lenses. Thymic epithelial cells stained
with CFDA-SE were imaged using a 470 nm filter while
excited with 457 nm laser light (green). CMRA stained
monocytes were excited at 543 nm and fluorescent lights were
detected at 576 nm (red). Cross-sectional images were obtained
in the same manner by taking pictures at 1 mm depth
increments down to 160 mm.
Scanning Electron Microscope (SEM) observations were
performed with a Philips XL30 SEM at 5.0 KV. Before
imaging, hydrogel scaffold samples were first fixed in 2%
cacodylate-buffered glutaraldehyde for 2 hours and washed
three times in 0.1 M cacodylate buffer for 30 min. Fixed
hydrogel scaffolds were dehydrated through a series of ethanol
solutions concentrations of 50, 70, 90, 95, and 100% for 10 min.
Dehydrated samples were freeze dried overnight utilizing a
Labconco FreeZone (Labconco), and then were coated with
gold for 180 s using a sputter coater (Desktop 2, Denton
Vacuum Inc.). Cross-section images of the internal architec-
ture were obtained after cutting the sample with a razor blade.
Results and discussion
ICC scaffold construction
The diameter of the spheres commonly used as colloidal
crystals is around 100–1000 nm for the purpose of matching
the optical band-gap in the visible region. Various methods
such as electrophoretic deposition,39 solvent evaporation,40
dipping,41 agitation42,43 and most recently spin coating44 have
been developed to construct highly ordered colloidal crystal
structures. In order to utilize the unique geometry of the
inverted colloidal crystals as a scaffold, the sphere size has to
be increased to the 10–1000 mm range. However, it is difficult
to obtain the same degree of order with micron scale beads
using the above methods of synthesis, mainly due to their
larger volume and heavier mass. Fortunately, micron-sized
beads offer two advantages over nanosize spheres. First, the
agitation of beads by shear force works more effectively
because of their larger volume. Previously, we reported the
construction of colloidal crystals by employing a gentle
agitation method.9 The second advantage is a faster sedimen-
tation rate due to their greater mass. However, the sedimenta-
tion rate was often too fast for them to self-assemble into a
closely-packed ordered array. The opposite problem, viz., how
to retard the sedimentation rate, was solved by introducing a
Pasture glass pipette before the beads entered into the mold.
The pipette extended the sedimentation distance and worked
as a thin funnel, which caused a bottleneck effect for
precipitating beads (Fig. 1a).
Once beads precipitated at the bottom of the mold, gentle
agitation generated by an ultrasonic bath assisted the move-
ment of beads and positioned them at the lowest energy spots.
This led to a highly packed and ordered array of spheres.
When the bottom area was covered with beads, their rugged
surface served as a template for the formation of the second
layer. Since structural defects accumulated from the bottom
area, incomplete layers and less ordered arrays were usually
observed on the top area.
The sedimentation rate was controlled further by adjusting
the concentration of beads in the solution and the time interval
between injections. For example, decreasing the amount of
beads and increasing the interval period provided more time
for the repositioning of precipitated beads. The use of
isopropanol guaranteed that the agitation was not too violent
to destroy the whole structure, while its buoyancy made it
easier for the PS beads to rearrange. Generated colloidal
crystal structures were 8 mm diameter and 1–1.5 mm in
thickness and SEM investigations revealed a highly ordered
hexagonally close-packed structure (Fig. 1b–d).
Following sedimentation, the colloidal crystals were heat
treated which resulted in partial melting of the spheres.29 This
step allowed the beads to stick together and on subsequent
cooling (re-solidification), junctions were created between the
spheres setting the structure in place. The resulting free
standing colloidal crystals were strong enough to be easily
handled and removed from the mold. The formation of the
junctions later prevented breakage of the crystal lattice during
the infiltration of scaffolding material and ensured the
connectivity between spheres and continuity of the chain of
pores in the final scaffold. The channel diameter was
determined at this stage, because the size of the melted
area depended on the annealing temperature. Increasing
Fig. 1 (a) Schematic diagram of the experimental setup used for
assembling micron-range polystyrene beads in 3D ordered structure.
(b–d) SEM images of the colloidal crystal structure made from 100 mm
polystyrene beads, showing the bottom (d), and internal structure at
different magnifications (b, c). Internal images were taken after cutting
the colloidal crystal with a razor blade. The small white spots on each
PS sphere in (b) are contact points between beads, which later become
channels.
3560 | J. Mater. Chem., 2006, 16, 3558–3564 This journal is � The Royal Society of Chemistry 2006
temperature enlarged the melting spot, but it caused shrinkage
of the colloidal crystal. Usually it led to the cracking of the
crystal structure and/or incomplete precursor solution infiltra-
tion. For 100 mm diameter PS beads, heat-treatment at 120 uCfor 4 hours gave the optimal result.
The diameter of pores was dictated by the details of the co-
culture system. Although monocytes and trypsinized thymic
epithelial cells have similar dimensions, epithelial cells stretch
out after attachment to the surface. Based on 2D charac-
terization, the size of the elongated thymic epithelial cells was
around 80–160 mm. Kotov et al. studied the pore size effect of
ICC scaffolds on a 3D cell culture utilizing three different sizes
of beads: 10 mm, 75 mm and 160 mm. The 75 mm pore diameter
favored bone marrow stromal cells nesting, while the 10 mm
pore size was too small for even a single cell, and 160 mm
diameter pores were too large to effectively entrap cells.10
Also, Zinger et al. investigated osteoblast-like cell cultures
on well-defined 2D cavities which were analogous to ICC
scaffolds, and found that 100 mm cavities favored osteoblast
attachment and growth.45 For the entrapment and transport of
suspension cells, the channel diameter, which is determined by
the size of the microspheres, was the most important para-
meter. PS beads which had a 100 mm diameter made 25–30 mm
diameter channels after annealing at 120 uC. The diameter of
the suspension cells (approximately 15–20 mm) was small
enough to enable them to pass through the channels.
As a scaffolding material, we selected poly(acrylamide)
hydrogel. Hydrogel is a broadly used scaffolding material
because of its biocompatibility, mechanical strength, and
transparency.46,47 The transparency of the hydrogel makes it
easier to monitor cell migration and growth deep inside the
scaffold using optical microscopy. Recently the observation of
cell growth at a depth greater than 250 mm 9 and real time cell
migration via a channel33 were reported. In addition, the
hydrogel exhibited another feature that facilitated its use in
ICC work. At low viscosity of the precursor solution, it
completely infiltrated to the colloidal crystal, and the whole
structure of the crystal template was transferred intact (Fig. 2).
The monomer concentration was set low enough to prevent
incomplete infiltration due to increased viscosity, and simulta-
neously to prevent deformation of the geometry during solvent
extraction.
The compressive moduli of hydrated and LBL coated ICC
scaffolds were 189.4 ¡ 5.89 KPa (Fig. 3). Compared to the
mechanical strengths of other porous hydrogel substrates, it
showed stronger mechanical stability.33,48 This was mainly
due to the higher content of polymer and the highly ordered
structure of the hydrogel ICC scaffolds. The achieved
compressive modulus was within the range of normal articular
cartilage.49 This degree of mechanical property was adequate
to construct artificial supports of targeted soft tissues.
Layer-by-layer surface modification
Hydrogel matrices rarely support adherent cell adhesion
without surface modification, because acrylamide polymer
chains do not have cell adhesion receptors, and the hydrophilic
nature of the hydrogel inhibits adsorption of cell binding
proteins on the gel surface.47 To render the surface bioactive,
we selected a layer-by-layer (LBL) deposition technique
instead of the commonly used covalent coupling of specific
peptide sequences such as RGD or an entire ECM protein to
the polymer.47 The driving forces for LBL coating are the
electrostatic, van der Waals, and hydrogen bonding interac-
tions between oppositely charged polyelectrolytes dissolved in
aqueous solution.50 This unique feature of the LBL technique
allows a complex porous 3D geometry, such as the intricate
and convoluted ICC surface, to be coated as long as fluid
transport in and out of the sample is not severely constrained.
It has been reported that 2D polyelectrolyte multilayers
Fig. 2 SEM images of the hydrogel scaffold after dehydration: (a)
bottom structure image, (b) internal structure image taken after cutting
the scaffold with a razor blade. The dehydration process caused
shrinkage of the ICC hydrogel scaffold, which led to some deformation
of the structure. Confocal images of a fluorescent hydrogel scaffolds:
(c) a 3D reconstruction of serial z-section images taken in 0.5 mm steps
showing the organization of main pores and interconnected channels
of a hydrogel ICC scaffold without shape deformation, and (d) 3D
overlapping images of serial z-sectional images of 160 mm interval with
5 mm step size.
Fig. 3 A compressive stress–strain curve from the mechanical
property test.
This journal is � The Royal Society of Chemistry 2006 J. Mater. Chem., 2006, 16, 3558–3564 | 3561
supported anchorage dependent cell attachment without using
adhesive proteins.51
In our system, we used clay nanoparticles/poly(diallyl-
dimethylammonium chloride) (PDDA) multilayers.38 The clay
particles are biocompatible and their flat shape effectively
covered the hydrogel surface. Coated clay nanoparticles
created nanoscale roughness, increased charging on the
surface, and created much stiffer films than hydrogel. An
increase of Young’s modulus was shown to be the primary
factor determining the adhesion of cells to materials.52,53 These
synchronous effects promoted cell adhesion.54,55 Ten layers of
PDDA/clay easily changed the surface property from cell
repulsive to cell adhesive, and thymic epithelial cells could
attach to the hydrogel scaffold.
Dynamic co-culture
Human thymic epithelial cells and human premyelote mono-
cytes were co-cultured in a rotary cell culture bioreactor.
Rotary motion induced convective flow, and the scaffold
geometry utilized this flow as a continuous driving force for
the cell movement.56 After five days of co-culture, the hydrogel
ICC scaffolds were imaged through a confocal microscope.
Different emission ranges of fluorescent dyes were used to
stain the thymus cells and monocytes with green and red,
respectively. Thymic epithelial cells attached to the cavity were
observed as green circles, and floating monocytes were imaged
as red spots (Fig. 4). Although many monocytes were diffused
out of the scaffold during the sample preparation, some of
them remained entrapped inside the pores. Confocal cross-
sectional images reveal that suspended cells were distributed
uniformly throughout the scaffold (Fig. 4b).
Co-cultured hydrogel ICC scaffolds were dehydrated and
observed under an SEM. The dehydration process deformed
the structure, which is the reason for the dimensional
differences between the two rows of images in Fig. 4. It was
found that the scaffold exterior was covered densely with
thymic epithelial cells, and their population reduced the
inward movement of other epithelial cells (Fig. 4a, b, d).
Secondly, epithelial cells migrated between pores through
interconnected channels, and some colonies expanded over
several pores (Fig. 4f). Thirdly, a few suspension cells were
trapped inside when they were observed at the interior of the
scaffold in SEM cross-sectional images (Fig. 4e). It suggests
that monocytes travel deep into the ICC scaffolds.
Modeling
To study the interaction of a floating cell with the scaffold
quantitatively, we constructed a simplified Brownian
Dynamics model. The cell was treated as a hard sphere of
radius acell that is suspended in an ICC geometry, composed of
hollow spherical chambers of nominal radius R connected by
channels of radius b.34 To simplify the treatment, we assumed
that the fluid inside the scaffold was quiescent and that the
motion of the cells was purely diffusive. Under the assump-
tions stated above, the motion of the cell results from a balance
between the drag force and the random Brownian force:
fdr/dt = FB (1)
where f = 6pgacell is the hydrodynamic drag exerted by the
solvent of viscosity g on a cell of radius acell, and r is the
position vector of the center of mass of the cell. The Brownian
Fig. 4 Confocal images (a–c) of co-cultured ICC hydrogel scaffolds; thymic epithelial cells (green) and monocytes (red). (a) Bottom area image
shows the surface of the scaffold was densely covered with thymic epithelial cells. Most of the monocytes around the edge of the scaffold were
released. (b) A cross-sectional image after cutting the co-cultured ICC scaffold with a razor blade shows decreasing thymic epithelial cell density
moving into the inside of the ICC scaffold. Monocytes were distributed through the whole ICC scaffold and a similar number of cells were
entrapped at each pore. (c) A lateral section image of 80 mm in depth. SEM images (d–f) of co-cultured hydrogel scaffolds. (d) Cross-sectional image
of the scaffold’s interior. (e) Entrapped monocytes. (f) Thymic epithelial cells covering pores and channels.
3562 | J. Mater. Chem., 2006, 16, 3558–3564 This journal is � The Royal Society of Chemistry 2006
force, FB, satisfies the fluctuation–dissipation theorem57 which
necessitates ,FB. = 0, and ,FBFB. = 2kBTfI. Here, I is the
unit tensor, kB = 1.38 6 10223 J K21 is Boltzmann’s constant,
and T is the absolute temperature.
The diffusivity, D, was obtained from the hydrodynamic
drag via the Einstein relation,57 D = kBT/f. In accordance with
microscopy measurements, we took R = 50 mm, b = 12.5 mm,
and acell = 7.5 mm. Thus, f = 6p(1 cP)(7.5 mm) = 1.414 61024 g s21, and D = kBT/f = 2.91 6 1022 mm2 s21. We used
the algorithm outlined by Larson57 to implement the BD
simulation, choosing the simulation time step, dt, so that
!6Ddt # 0.05acell. We employed reflecting boundary condi-
tions to model collisions between the cell and the scaffold.
Grigoriev et al. considered a dimensionless Brownian
particle trapped inside a spherical chamber of volume V.58
They estimated that the time, t*, that it takes for the particle to
escape from a small circular hole of radius b on the surface of
the chamber is given by t* = V/4bD, where D is the diffusivity
of the particle. We adapted the expression for t* to obtain a
crude estimate for the escape time of a Brownian particle of
finite size from an ICC scaffold as:
t*ICC = (p/3ZD)(R 2 acell)3/(b 2 acell) (2)
where Z = 12 is the co-ordination number of the ICC lattice.
From eqn (2), we obtained t*ICC y 5.5 6 105/12 s y 12 hours.
Thus, the ICC geometry provides very suitable geometry
for cell interactions due to partial entrapment of the cells in
the cavity.
We simulated the dynamics of the cell in the ICC scaffold
using BD, and recorded its trajectory from t = 0 to t =
1000 days. Over this period, the cell visited several chambers.
From the simulation, we observed that by the time the cell
vacated a chamber by escaping through the interconnecting
channel to another chamber, it thoroughly, and uniformly,
sampled the whole chamber. In other words, the amount of
time the cell spent in any region of the chamber was
proportional to the volume of that region. Fig. 5a shows a
cross section of a spherical chamber that has been divided into
shells of equal thickness, DR = acell. These shells do not have
the same volume. For illustration, if we approximate the
volume of a shell by DVshell = 4pRi2DR, where Ri is the inner
radius of the shell, we can see that the volume of the outer
shells is greater than that of the inner shells. As mentioned
previously, the center of mass of the cell resides in a shell, in
proportion to the volume of that shell. Thus, it spends a
significant fraction of time (about 41%, see Fig. 5a) in the
outermost shell, where the distance between the surface of the
cell and the inner surface of the chamber is less than or equal
to the radius of the cell. Thus the ICC geometry fosters contacts
between the cell and the matrix surface or between the
suspension and adherent cells in a co-culture.
Conclusions
We have demonstrated a 3D co-culture model system within a
LBL surface modified ICC scaffold in both experimental and
modeling works. Clay/PDDA multilayer successfully formed
on a complex porous ICC scaffold walls, and significantly
improved adhesion of epithelial cells. The unique geometry of
ICC scaffolds can accommodate two different types of cells
within a same chamber. Modeling results indicated that cells
were effectively trapped in the spherical chambers and that
entrapped suspension cells spent a significant fraction of time
in the vicinity of the ICC chamber wall or the epithelial coating
in co-cultures.
Well controlled multiscale structures which can build real-
size organ systems and generate the essential subcellular
morphology are a key factor for the successful investigation
of cell–molecule and cell–cell interactions.12,59 It is obvious
that the full function of the tissues and organs cannot be
recovered without rebuilding the ultrastructure of the tissue
itself. Proposed ICC scaffolds and surface modification
utilizing a LBL technique will be excellent approaches for this
purpose. ICC scaffold structure generates super- and cellular-
scale microenvironments for intense cell contacts with other
types of cell or matrix. On this surface, various insoluble
signaling molecules such as ECM components, membrane
bound receptors and ligands can be incorporated through a
LBL method which can produce a subcellular, nanoscale
resolution environment for cellular receptor–molecular inter-
actions. In particular, this could greatly facilitate the study of B
and T cell development from stem cells which requires under-
standing and controlling precise 3D molecular interactions.
Acknowledgements
This work was supported by a grant from DARPA and
VaxDesign Inc. We thank Dr Michael Solomon and Tesfu
Fig. 5 Radial probability distribution of a finite-sized Brownian
particle of radius 7.5 mm diffusing in a spherical ICC chamber obtained
from BD simulations, when the chamber is divided into shells of the
same (a) thickness, and (b) volume. In (a), the dotted arc and the disc
represent the inner surface of the chamber, and the cell which is
modeled as a hard sphere, respectively. The thickness of each shell is
equal to the radius of the cell, acell. From (b), it can be seen that the cell
spends the same amount of time in each of the equi-volume shells,
whereas in (a), it spends more time in the exterior shells due to their
greater volume.
This journal is � The Royal Society of Chemistry 2006 J. Mater. Chem., 2006, 16, 3558–3564 | 3563
Solomon (University of Michigan, Ann Arbor) for assistance
with the laser scanning confocal microscope.
References
1 T. Prasad, R. Rengarajan, D. M. Mittleman and V. L. Colvin, Opt.Mater., 2005, 27, 1250.
2 D. Wang, V. Salgueirino-Maceira, L. M. Liz-Marzan andF. Caruso, Adv. Mater., 2002, 14, 908.
3 Y. Wang and F. Caruso, Adv. Funct. Mater., 2004, 14, 1012.4 C. E. Reese, A. V. Mikhonin, M. Kamenjicki, A. Tikhonov and
S. A. Asher, J. Am. Chem. Soc., 2004, 126, 1493.5 N. Tetreault, H. Miguez and G. A. Ozin, Adv. Mater., 2004, 16,
1471.6 Y. Y. Song, D. Zhang, W. Gao and X. H. Xia, Chem.–Eur. J.,
2005, 11, 2177.7 R. C. Schroden, C. F. Blanford, B. J. Melde, B. J. S. Johnson and
A. Stein, Chem. Mater., 2001, 13, 1074.8 Y. Liu, S. Wang, J. W. Lee and N. A. Kotov, Chem. Mater., 2005,
17, 4918.9 Y. Zhang, S. Wang, M. Eghtedari, M. Motamedi and N. A. Kotov,
Adv. Funct. Mater., 2005, 15, 725.10 N. A. Kotov, Y. Liu, S. Wang, C. Cumming, M. Eghtedari,
G. Vargas, M. Motamedi, J. Nichols and J. Cortiella, Langmuir,2004, 20, 7887.
11 A. Abbott, Nature, 2003, 424, 870.12 S. Kale, S. Biermann, C. Edwards, C. Tarnowski, M. Morris and
M. W. Long, Nat. Biotechnol., 2000, 18, 954.13 Y. Xie, S. T. Yang and D. A. Kniss, Tissue Eng., 2001, 7, 585.14 C. Trojani, P. Weiss, J. F. Michiels, C. Vinatier, J. Guicheux,
G. Daculsi, P. Gaudray, G. F. Carle and N. Rochet, Biomaterials,2005, 26, 5509.
15 H. J. Evans, J. K. Sweet, R. L. Price, M. Yost and R. L. Goodwin,Am. J. Physiol., 2003, 285, H570.
16 D. Ferrera, S. Poggi, C. Biassoni, G. R. Dickson, S. Astigiano,O. Barbieri, A. Favre, A. T. Franzi, A. Strangio, A. Federici andP. Manduca, Bone, 2002, 30, 718.
17 J. Zhang, C. Niu, L. Ye, H. Huang, X. He, W. Tong, J. Ross,J. Haug, T. Johnson, J. Q. Feng, S. Harris, L. M. Wiedemann,Y. Mishina and L. Li, Nature, 2003, 425, 836.
18 E. Fuchs, T. Tumbar and G. Guasch, Cell, 2004, 116, 769.19 A. Spradling, D. Drummond-Barbosa and T. Kai, Nature, 2001,
414, 98.20 T. Imamura, L. Cui, R. Teng, K. Johkura, Y. Okouchi,
K. Asanuma, N. Ogiwara and K. Sasaki, Tissue Eng., 2004, 10,1716.
21 S. Ding and P. G. Schultz, Nat. Biotechnol., 2004, 22, 833.22 A. W. Duncan, F. M. Rattis, L. N. DiMascio, K. L. Congdon,
G. Pazianos, C. Zhao, K. Yoon, J. M. Cook, K. Willert, N. Gaianoand T. Reya, Nat. Immunol., 2004, 6, 314.
23 H. Liu and K. Roy, Tissue Eng., 2005, 11, 319.24 M. C. Poznansky, R. H. Evans, R. B. Foxall, I. T. Olszak,
A. H. Piascik, K. E. Hartman, C. Brander, T. H. Meyer,M. J. Pykett, K. T. Chabner, S. A. Kalams, M. Rosenzweig andD. T. Scadden, Nat. Biotechnol., 2000, 18, 729.
25 E. K. F. Yim and K. W. Leong, Nanomedicine, 2005, 1, 10.26 P. Bousso, I. R. Bhakta, R. S. Lewis and E. Robey, Science, 2002,
296, 1876.
27 J. M. Williams, A. Adewunmi, R. M. Schek, C. L. Flanagan,P. H. Krebsbach, S. E. Feinberg, S. J. Hollister and S. Das,Biomaterials, 2005, 26, 4817.
28 X. Liu and X. P. Ma, Ann. Biomed. Eng., 2004, 32, 477.29 P. X. Ma and J. W. Choi, Tissue Eng., 2001, 7, 23.30 T. B. F. Woodfield, J. Malda, J. de Wijn, F. Peters, J. Riesle and
C. A. van Blitterswijk, Biomaterials, 2004, 25, 4149.31 S. Yang, K. F. Leong, Z. Du and C. K. Chua, Tissue Eng., 2002,
8, 1.32 X. Wang, Y. Yan, Y. Pan, Z. Xiong, H. Liu, J. Cheng, F. Liu,
F. Lin, R. Wu, R. Zhang and Q. Lu, Tissue Eng., 2006, 12, 83.33 A. N. Stachowiak, A. Bershteyn, E. Tzatzalos and D. J. Irvine,
Adv. Mater., 2005, 17, 399.34 S. Shanbhag, J. W. Lee and N. Kotov, Biomaterials, 2005, 26,
5581.35 I. R. Lemischka and K. A. Moore, Nature, 2003, 425, 778.36 M. M. Davis, M. Krogsgaard, J. B. Huppa, C. Sumen,
M. A. Purbhoo, D. J. Irvine, L. C. Wu and L. Ehrlich, Annu.Rev. Biochem., 2003, 72, 717.
37 S. J. Collins, Blood, 1987, 70, 1233.38 Z. Tang, N. A. Kotov, S. Magonov and B. Ozturk, Nat. Mater.,
2003, 2, 413.39 A. L. Rogach, N. A. Kotov, D. S. Koktysh, J. W. Ostrander and
G. A. Ragoisha, Chem. Mater., 2000, 12, 2721.40 J. P. Hoogenboom, C. Retif, E. de Bres, M. van de Boer,
A. K. Langen-Suurling, J. Romijn and A. van Blaaderen, NanoLett., 2004, 4, 205.
41 S. H. Im, M. H. Kim and O. O. Park, Chem. Mater., 2003, 15,1797.
42 O. Vickreva, O. Kalinina and E. Kumacheva, Adv. Mater., 2000,12, 110.
43 M. Sasaki and K. Hane, J. Appl. Phys., 1996, 80, 5427.44 P. Jiang and M. J. McFarland, J. Am. Chem. Soc., 2004, 126,
13778.45 O. Zinger, G. Zhao, Z. Schwartz, J. Simpson, M. Wieland,
D. Landolt and B. Boyan, Biomaterials, 2004, 26, 1837.46 Y. Luo and M. S. Shoichet, Nat. Mater., 2004, 3, 249.47 J. L. Drury and D. J. Mooney, Biomaterials, 2003, 24, 4337.48 S. H. M. Soentjens, D. L. Nettles, M. A. Carnahan, L. A. Setton
and M. W. Grinstaff, Biomacromolecules, 2006, 7, 310.49 P. Kiviranta, J. Rieppo, R. K. Korhonen, P. Julkunen, J. Toyras
and J. S. Jurvelin, J. Orthop. Res., 2006, 24, 690.50 G. Decher, Science, 1997, 277, 1232.51 S. Kidambi, I. Lee and C. Chan, J. Am. Chem. Soc., 2004, 126,
16286.52 M. C. Berg, S. Y. Yang, P. T. Hammond and M. F. Rubner,
Langmuir, 2004, 20, 1362.53 H. Zheng, M. C. Berg, M. F. Rubner and P. T. Hammond,
Langmuir, 2004, 20, 7215.54 G. B. Schneider, A. English, M. Abraham, R. Zaharias,
C. Stanford and J. Keller, Biomaterials, 2004, 25, 3023.55 S. Kay, A. Thapa, K. M. Haberstroh and T. J. Webster, Tissue
Eng., 2002, 8, 753.56 P. A. Plett, S. M. Frankovitz, R. Abonour and C. M. Orschell-
Traycoff, In Vitro Cell. Dev. Biol.: Anim., 2001, 37, 73.57 G. R. Larson, The Structure and Rheology of Complex Fluids,
Oxford University Press, New York, 1999.58 I. V. Grigoriev, Y. A. Makhnovskii, A. M. Berezhkovskii and
V. Y. Zitserman, J. Chem. Phys., 2002, 116, 9574.59 V. Vogel and M. Sheetz, Nat. Rev. Mol. Cell Biol., 2006, 7, 265.
3564 | J. Mater. Chem., 2006, 16, 3558–3564 This journal is � The Royal Society of Chemistry 2006