Layer by Layer Three-dimensional Tissue
Epitaxy by Cell-Laden Hydrogel Droplets
SangJun Moon, Ph.D.,1,*Syed K. Hasan, M.D.,1,*Young S. Song, Ph.D.,1
Feng Xu, Ph.D.,1Hasan Onur Keles, B.Sc.,1Fahim Manzur, B.Sc.,1Sohan Mikkilineni,1
Jong Wook Hong, Ph.D.,2Jiro Nagatomi, Ph.D.,3Edward Haeggstrom, Ph.D.,4
Ali Khademhosseini, Ph.D.,5,6and Utkan Demirci, Ph.D.1,5,6
The ability to bioengineer three-dimensional (3D) tissues is a potentially powerful approach to treat diverse
diseases such as cancer, loss of tissue function, or organ failure. Traditional tissue engineering methods, how-
ever, face challenges in fabricating 3D tissue constructs that resemble the native tissue microvasculature and
microarchitectures. We have developed a bioprinter that can be used to print 3D patches of smooth muscle cells
(5mm?5mm?81mm) encapsulated within collagen. Current inkjet printing systems suffer from loss of cell
viability and clogging. To overcome these limitations, we developed a system that uses mechanical valves to
print high viscosity hydrogel precursors containing cells. The bioprinting platform that we developed enables (i)
printing of multilayered 3D cell-laden hydrogel structures (16.2mm thick per layer) with controlled spatial
resolution (proximal axis: 18.0?7.0mm and distal axis: 0.5?4.9mm), (ii) high-throughput droplet generation (1s
per layer, 160 droplets=s), (iii) cell seeding uniformity (26?2cells=mm2at 1 million cells=mL, 122?20cells=mm2
at 5 million cells=mL, and 216?38cells=mm2at 10 million cells=mL), and (iv) long-term viability in culture
(>90%, 14 days). This platform to print 3D tissue constructs may be beneficial for regenerative medicine ap-
plications by enabling the fabrication of printed replacement tissues.
dimensional (3D) tissues as an alternative treatment for
various diseases such as loss of tissue function or organ
failure.1–5Often in tissue engineering, two-dimensional (2D)
or 3D scaffolds are employed to generate tissues in vitro.6,7
However, engineered tissues generated in 2D cultures do not
mimic the complex microarchitecture of native tissues. Also,
current 3D polymer scaffolding approaches are not suitable
for fabricating complex tissue structures due to lack of spatial
and temporal control during cell seeding.8–10In the past
decade, deposition of polymers=metals=cells by printing has
gained momentum in electronic circuit board printing,
printing of transistors, and tissue printing.11,12Printing
technology shows promise in overcoming the limitations
associated with seeding cells on scaffolds. For example, bio-
and tissue engineering present bioengineered three-
printing methods, such as inkjet13–15and laser printing16–19
techniques, have been employed to control cell placement in
2D or 3D. However, some challenges still remain in existing
tissue printing systems such as low cell viability, loss of
cellular functionality, and clogging.20–22Cell printing also
requires extracellular matrix (ECM) to build 3D structures for
long-term culture. However, the current piezo-based inkjet
printing system is not easily adapted for high viscosity so-
lutions such as collagen ECM, since it requires high impact
force to generate droplets. To overcome these limitations,
alginate-based cell printing23,24and 3D fiber deposition25
approaches were used to encapsulate cells in ECM. Alginate-
based cell printing is adapted to the conventional piezo-
based bioprinter to prevent the rapid clogging issues by
printing a low viscosity calcium chloride as crosslinking
agent. However, for gelation the calcium must diffuse into
alginic acid, which limits the droplet placement resolution.
During the diffusion process, a change in pH also affects cell
1Bio-Acoustic MEMS in Medicine (BAMM) Laboratory, Center for Biomedical Engineering, Brigham and Women’s Hospital, Harvard
Medical School, Cambridge, Massachusetts.
2Department of Mechanical Engineering, Materials Research and Education Center, Auburn University, Auburn, Alabama.
3Department of Bioengineering, 313 Rhodes Engineering Research Center, Clemson University, Clemson, South Carolina.
4Department of Physics, University of Helsinki, Helsinki, Finland.
5Harvard-Massachusetts Institutes of Technology Health Sciences and Technology, Cambridge, Massachusetts.
6Center for Biomedical Engineering, Brigham and Women’s Hospital, Harvard Medical School, Cambridge, Massachusetts.
*These authors contributed equally to this work.
TISSUE ENGINEERING: Part C
Volume 15, Number 00, 2009
ª Mary Ann Liebert, Inc.
viability.23The other approach uses the squeezing of ECM
precursors from the nozzle to eliminate clogging. This ap-
proach may be limited in terms of low resolution and
An emerging approach to enhance bioprinting is to use a
nozzle-free acoustic ejector, which prevents clogging during
droplet generation.26–28Another approach features a me-
chanical valve ejector that uses a pressure source to over-
come the surface tension of high viscosity liquids.29–31This
mechanical ejector was applied for cryopreservation of cells
in droplets and for cell printing. In this article, we built on
the system by creating a cell-laden hydrogel droplet depo-
sition system that can create 3D structures made of collagen,
a temperature-sensitive gel. We adopted the system to
evaluate a model structure using bladder smooth muscle
cells (SMCs) to engineer tissues. We demonstrate that this
bioprinting system can be used to (i) pattern cell-laden hy-
drogel droplets with microscale resolution, (ii) print hydro-
gel droplets containing cells in a rapid and uniform manner,
and (iii) maintain long-term cell viability.
Materials and Methods
SMC collagen encapsulation
Primary bladder SMCs from Sprague Dawley rat were
harvested according to a previously established protocol.32
SMC culture medium was prepared by mixing 445mL Dul-
becco’s modified Eagle’s medium (Gibco, Carlsbad, CA,
11965-092), 50mL fetal bovine serum (Gibco, 10439-024), and
Strep (Sigma, St. Louis, MO, P4333) through a sterile filter
(500mL, Express Plus 0.22mm membrane, SCGPU05RE).
SMCs were cultured under standard conditions (378C, 5%
CO2) in a humidified incubator (Forma Scientific, Waltham,
MA, CO2 water jacketed incubator). After the culture
reached 80% confluency, cells were trypsinized (10?, 0.5
trypsin–EDTA; Gibco, 15400), washed, and resuspended in
SMC medium to be mixed with collagen. Collagen solution
was prepared by mixing 250mL type I bovine collagen (MP
Biomedicals, Solon, OH) with 50mL sterile H2O, 50mL 10?
phosphate-buffered saline (PBS) (DPBS, Carlsbad, CA,
14190), 50mL fetal bovine serum, 50mL SMC medium, and
50mL NaOH (0.1M, Sigma, 55881) and kept at 48C before
being mixed with SMCs (1:1 ratio).
3D printing using a droplet ejector
The droplet generation process was adjusted by control-
ling nitrogen gas pressure, valve opening duration, and cell
concentration (Fig. 1). To fabricate a collagen-coated sub-
strate, agarose (10% v=v mixture with distilled water and
agarose powder; Fisher, Pittsburgh, PA, BP1360-100) was
poured on the bare Petri dish (Falcon, Pittsburgh, PA,
sterile field (Cleanroom International, Grand Rapids, MI, 13202). (b) Schematic of droplet ejector shows cells and collagen
mixture flowing into the valve driven by constant air pressure. Mixture of cells and collagen solution was loaded into a 10mL
syringe reservoir. (c) Signal flow chart shows that the xyz stage is controlled by a controller that was synchronized with a
pulse generator and a control PC. With programmed sequences to build a three-dimensional (3D) structure, the apparatus can
control ejection conditions, that is, stage speed, pressure, valve on=off frequency, and valve opening duration. Color images
available online at www.liebertonline.com=ten.
Illustration of cell encapsulating droplet printing onto a substrate. (a) Image of the cell printing setup enclosed in a
2 MOON ET AL.
35-3002) to enhance adhesion between the Petri dish and
collagen. Collagen solution was then manually spread on the
agarose surface and gelled. The cell-laden collagen droplets
were printed onto the collagen-coated substrate. To maintain
the droplet size, we kept the valve opening duration at 60ms
and nitrogen gas pressure at 34.4kPa. To control the cell
density in droplets, we used three different cell concentra-
tions, 1?106, 5?106, and 10?106cells=mL. The cell viability
before and after printing was evaluated using a Live=Dead
kit (Invitrogen, Carlsbad, CA, L3224). The staining solution
was prepared with 0.5mL of (1mg=mL) calcein AM and 2mL
of (1mg=mL) ethidium homodimer solution in 1mL of PBS
for 1min. The staining solution was poured onto printed
structures and incubated for 10min at 378C. The stained cells
in the patch were manually counted under a florescent mi-
croscope (Eclipse Ti-s; Nikon, Melville, NY).
Using the valve-based droplet ejector setup that was pre-
viously described,29,30cells were ejected on the prepared
substrate. Using 1?106, 5?106, or 10?106cells=mL, the
10mL syringe attached to the ejector was filled with the
desired cell=collagen suspension. The ejector and collagen
were kept cool with liquid nitrogen (LN2, *58C in gas
phase) vapor to minimize viscosity changes of collagen that
can solidify at room temperature. Each printed layer was
gelled by incubation at 378C for 5min. Subsequently, another
layer of collagen was printed onto the first layer. This process
of layering was repeated to create 3D tissue structures.
Staining and microscopy
Printed SMC patches were gelled at 378C for 5min before
SMC medium was added and incubated overnight. After
24h, medium was aspirated off, and printed patches were
washed three times with PBS at room temperature and fixed
in 2mL of 4% paraformaldehyde (Sigma). These patches
were then rinsed with PBS three times and permeabilized
with 1mL of detergent solution (mixture of 4% bovine serum
albumin and 0.1% TritonX-100 in PBS solution; Sigma). The
specimens were incubated with primary antibody (actin,
connexin-43, and mouse monoclonal immunoglobulin G
[IgG], 1:50 dilution in PBS; Santa Cruz Biotechnology, Santa
Cruz, CA) and 5mg=mL nuclear stain 40,6-diamidino-2-phe-
nylindole (Invitrogen) at 378C for 40min. Secondary anti-
bodies (goat anti-mouse IgG fluorescein isothiocyanate and
IgG R, 1:50 dilution in PBS; Santa Cruz Biotechnology) were
also incubated at 258C for 40min. After each incubation
process, excess antibody was washed off, and stained SMC
patches were imaged under the florescent microscope
(Eclipse Ti-s; Nikon). The number of cells per square milli-
meter was plotted using SigmaPlot?that depicted cell dis-
tribution as a contour plot of an entire patch.
Results and Discussions
Uniform cell seeding density is critical for tissue engi-
neering, since it controls the average cell-to-cell distances
that influence cell-to-cell communication. The overall mor-
phological characteristics of a tissue construct depend on this
uniformity. To achieve 3D tissue structures with spatial
control of cell seeding, we characterized (i) the number of
cells per droplet as a function of cell loading concentration,
(ii) droplet printing precision, (iii) overlapping cell-laden
collagen droplets to fabricate seamless line structures, and
(iv) number of cells per unit area in a printed patch.
The mechanical valve was attached to a micrometer pre-
cision xyz stage that enabled 3D spatial motion. The move-
ment of the stage was synchronized with droplet generation
signal resulting in 3D patterning capability. The platform
spatially and temporally controlled the droplet placement
(Fig. 1). First, we evaluated the position and density of cells
in the biomaterial by printing cell-laden droplets in multiple
layers. The cell-laden collagen droplets landed onto a Petri
dish surface that was coated with collagen gel (Fig. 2a). This
controlled placement allowed the system to deposit a cell-
laden hydrogel droplet epitaxially in 2D and 3D using
droplets with 650?18mm spread diameter on the surface.
Uniform cell seeding was investigated by characterizing
where droplets land onto a surface during droplet generation
and xyz stage movement along a temporal line (distal axis,
Fig. 2a). The landing locations and placement variation (dx
and dy) of droplets determine the overlap between droplets
when patterning lines and patches in 3D. The droplet ejection
directionality was the major determinant of this variation.
The system achieves 0.5?4.9 and 18.0?7.0mm variation in
the x (distal) and y (proximal) directions, respectively. These
variations were negligible compared to the 650?18mm
spread droplet diameter. To create layered structures using
an intermediate collagen layer was printed between the first
layer of droplets and second layer of droplets (Fig. 2b). The
adjacent droplets gel together and form a single seamless
layer. Further, a secondary droplet array was printed on top
of the gelled layers to pattern droplets in a 3D micro-
architecture (Fig. 2c). The cell-laden collagen droplet in the
first layer was printed at a lower cell concentration on the
substrate than the collagen droplet printed in the secondary
layer to depict a layered structure.
Second, we characterized the number of cells per droplet
at three cell loading densities and the cell viability of the
printing platform (Fig. 2d). It showed 6?1cells per droplet
at 1?106cells=mL, 29?5cells per droplet at 5?106cells=mL,
and 54?8cells per droplet at 10?106cells=mL. The number
of cells per droplet was repeatable over ejected droplets at
various cell loading concentrations. Further, the number of
cells per droplet increased with increasing cell loading den-
sity in the ejector reservoir. The number of cells that can be
packed in a single droplet does not increase linearly with the
loading density. Consequently, it is harder to pack more cells
into a fixed droplet volume. To better understand cell seed-
ing density, the mean and standard deviation for number of
cells per droplet were investigated. Smaller standard devia-
tion can be translated into a more uniform seeding density as
cells are patterned to create 3D constructs. The platform also
printed cells with high viability 94.8?0.8% compared to the
culture flask viability. The viability was calculated by the
ratio of pre-ejection cell viability (96.1?1.9%) and post-
ejection cell viability (91.1?2.3%) by counting 250 printed
cells (Fig. 2d). The results showed that system precision,
printing cell viability, and cells per droplet uniformity suf-
ficed to establish controlled cell seeding density with high
The third step was to print overlapping collagen droplets
to pattern cell-laden collagen lines as we build a 3D structure.
LAYER BY LAYER 3D TISSUE EPITAXY3
valve that is operated by a controlled pulse width (open period of the valve) and a frequency (on=off time of the valve) to
generate required volume and timed placement of droplets onto a substrate, respectively (Fig. 1). Droplets are printed to form
multiple layers of collagen; smooth muscle cell (SMC)–laden collagen droplet array (gray color sphere), intermediate collagen
layer, and top SMC-laden droplet layer (blue color sphere). Image of a printed array of collagen droplets (b) and image of a
multilayered array on a slide glass (c). A gray-colored droplet indicates the bottom layer of collagen shown in (c). dx and dy
are measured between centers of each droplet in different layers. Mean and standard deviation values of x (distal axis) and y
(proximal axis; moving axis) directional variations were 0.5?4.9 and 18.0?7.0mm, respectively. (d) Number of cells per
droplet and cell viability as a function of loading concentrations. Mean and standard deviation values of encapsulated cells
were 6?1, 29?5, and 54?8cells per droplet in 1?106, 5?106, and 10?106cells=mL, respectively. The cell printing platform
showed 94.8?0.8% average cell viability for three different concentrations compared to the culture flask. Each cell loading
concentration had 94.9?1.7%, 95.8?1.3%, and 93.5?3.0% cell viability. Scale bar: 200mm. Color images available online at
Printing platform for 3D cell-laden droplet printing. (a) Cell-laden hydrogel droplets are generated by a mechanical
4MOON ET AL.
An illustration describing placement of droplets in a printed
line pattern is shown by overhanging printed cell-line
bridges in separate layers (Fig. 3a). The overlap between the
adjacent droplets was maintained at 50% by the temporally
controlled ejection. To test the system operation, two colla-
gen lines were printed side by side in a single layer (Fig. 3b),
and multiple lines were printed within separate layers of a
3D structure in a crossover pattern (Fig. 3c). These cell-laden
collagen lines were placed on top of each other in the z di-
rection by printing a cell-less collagen layer within between
two layers. The magnified images of the cross-pattern
bridges of printed cell lines are shown in Figures 3d and e.
Finally, native tissue comprises multiple cell layers. To
mimic such tissue architecture, the bioprinting system
the line pattern form a 3D structure like a bridge separated by a spacing layer of hydrogel. (b, c) Dot and solid lines represent
the edge of bottom and top collagen lines; dried collagen line pattern in (b) and multilayered line pattern in (c). (d, e)
Magnified images show cross-patterned lines on separate layers. The top and bottom layers are shown with two focused
images: bottom focused image in (d) and top focused image in (e). Scale bar: 200mm. Color images available online at
Printing of cells in lines of hydrogel microstructures. (a) Illustration of printed droplets in a line pattern. Top layer of
LAYER BY LAYER 3D TISSUE EPITAXY5
employs a 3D printing capability using an epitaxial method
(layer by layer) (Fig. 4a). To print smooth muscle tissue con-
structs, cell-laden collagen droplets were patterned on top of
earlier printed layers. The challenge of 3D patterning was
overcome by first gelling the initial printed layer and then
depositing additional cell-laden hydrogel droplets on top of
the previously printed layer like in layer-by-layer epitaxy.
First, a bottom cell-less collagen layer was placed in agarose.
Then, on top of this layer a cell-laden collagen layer was
printed. This process was repeated creating five cell-less
and two cell-laden collagen layers (81mm thick). To observe
the multiple layers, a motorized system was created that
steps the microscope focus (Fig. 5). Images were taken at
each focus point with 16.2mm steps (Fig. 4b–e). The printed
3D multilayer SMC-laden collagen construct was stained
with 40,6-diamidino-2-phenylindole. Focal images show prin-
ted layers with stained cells and without cells. The cell-laden
the cell-less collagen layers only show background due
to staining of the gel (Fig. 4b, d). The described epitaxial
patch imaging.Thedistance betweeneachimagedlayeris16.2mmwhich iscontrolledbytimedimagingandmovingspeed ofa
z-axis knob (Fig. 5). (b–e) Focal images of 3D patch layers; top layer of printed collagen in (b), second layer of SMC patch in (c),
intermediate collagen layer in (d), and first layer of SMC patch in (e). (f) Cell distribution of two-dimensional patch of 1, 5, and
10million cells=mLconcentration after printing(day0).Each patch sizeis 5?5mm.Average numberand standarddeviation of
printed cells for each patch were 26?2cells=mm2(average?standard deviation) at 1?106cells=mL, 122?20cells=mm2at
5?106cells=mL, and 216?38cells=mm2at 10?106cells=mL. The number of cells is represented in log scale for comparison
between 1?106and 10?106cells=mL. Scale bar: 100mm. Color images available online at www.liebertonline.com=ten.
Focalimagesofaprinted 3DSMCtissueconstructandtwo-dimensionalcellseedingdistribution. (a)Illustrationof3D
6 MOON ET AL.
knob of a fluorescence microscope body by a timing belt. Each image was taken at a scheduled time by a charge-coupled
device camera control software. The distance of each layer was calculated by the reference index of the microscope
(65mm=3608), motor speed (1808=s), and imaging time control (0.5s=image). These conditions gave a resolution of 16.2mm
separation between each image for an 81-mm thick patch (five layers). Color images available online at www
Focal 3D imaging method using a motorized microscope. A direct current motor was connected to control the z-axis
within a single layer of printed SMC patch: day(s) 1 in (a), 2 in (b), 4 in (c), and 7 in (d) for 5?106cells=mL. Each patch size is
5?5mm (xy-axis index). The cell distribution of printed cells for each patch was 289?47cells=mm2(average?standard
deviation) in (a), 489?48cells=mm2in (b), 897?125cells=mm2in (c), and 1183?236cells=mm2in (d). Color images available
online at www.liebertonline.com=ten.
Cell distribution of printed SMC patch in culture. (a–d) Quantification of cell distribution and cell proliferation
LAYER BY LAYER 3D TISSUE EPITAXY7
period of time in collagen patches for three initial cell concentrations (Cinit), that is, 1?106, 5?106, and 10?106cells=mL. (a)
The total number of cells per square millimeter in three different initial printing concentrations were measured from day 0 to
7. Inset represents an enlarged figure of 1?106cells=mL initial cell loading density. After 7 days of culturing (Csat), 270?25,
1183?236, and 2097?287cells=mm2were observed for 1?106, 5?106, and 10?106cells=mL, respectively. The inflection time
(tinflection) of sigmoid regression curves was 2.6 days for 5?106cells=mL and 3.2 days for 10?106cells=mL. In case of
26?1.7cells=mm2initial cell loading density, proliferation rate of cells showed an exponential increment. The unknown
factor for cell proliferation b is a factor of each exponent and sigmoid regression functions, 0.2 for 1?106cells=mL, 1.3 for
5?106cells=mL, and 1.7 for 10?106cells=mL. (b–e) Stained SMC patch images for 1?106cells=mL concentration after day(s)
in culture: day 4 culture of SMC patch stained with 40,6-diamidino-2-phenylindole (DAPI) (blue) and actin (green) under a
light microscope (10?) in (b), day 7 SMCs stained with DAPI and actin in (c), SMCs stained with DAPI (blue) at day 14 in
culture in (d), SMCs stained with DAPI and connexin-43 (red) at day 14 in culture in (e). Scale bar: 100mm. Color images
available online at www.liebertonline.com=ten.
Characterization of printed SMC patch in culture. The proliferation graph shows increasing number of cells over a
8 MOON ET AL.
method was used to observe cell seeding densities within a
single printed layer at three different cell densities, 1?106,
216?38cells=mm2at 10?106cells=mL, 122?20cells=mm2at
5?106cells=mL, and 26?2cells=mm2at 1?106cells=mL.
The patches were imaged after printing, and the number
of cells was averaged per square millimeter in each image for
an entire patch area of 25mm2. We validated the distribu-
tion, uniformity, and variation of cell seeding density by the
printing method. The topographic color coding of the top
view of these patches reveals the cell distribution over 1–7
days for 5?106cells=mL cell printing concentration (Fig. 6a–
d). The color coding indicates the cell concentration in that
area (see the legend). The increased cell seeding density
correlates with the increased number of cells per droplet (Fig.
7a). This characterization is crucial, since it builds the logical
tie between a cell-laden hydrogel droplet and a printed 3D
tissue construct. However, the proliferation rate is not linear
as a function of cell density and culture time. The rates show
a sigmoid tendency as a function of culture duration, which
indicates that initial high proliferation rates decrease as
the number of cells per unit area increases. The inflection
time, tinflection, of the sigmoid regression curves were 2.6 days
for 5?106cells=mL and 3.2 days 10?106cells=mL. In case of
26?1.7cells=mm2initial cell loading density, the prolifera-
tion rate of cells showed an exponential increment. The ex-
ponent andthe sigmoid regression
unknown factor, b, which is related to cell proliferation, 0.2
for 1?106cells=mL, 1.3 for 5?106cells=mL, and 1.7 for
10?106cells=mL. The number of cells per droplet and the
precise positioning of these droplets in a 3D architecture
determine the cell seeding density of the patch before the
long-term culture. Such high-throughput capability and cell
seeding control to create 3D tissue constructs allow poten-
tially rapid characterization and optimization of tissues.
Printing a 5?5mm patch takes 10s with 160Hz ejection
frequency. The total time becomes 10min including the ge-
lation time to build a secondary layer. This processing time
indicates the high-throughput aspect of the system compared
to the conventional scaffold methods that take 1–2h to build
a single patch. Cells are also observed to adhere and spread
within the printed cell-laden collagen layer (Fig. 7b–e). In
long-term culture, cells were observed to be viable as dem-
onstrated by histological stains. During days 4 and 7, the
printed cells expressed actin after the printing and culturing
steps (Fig. 7b, c). Patches on the 14th day of culture ex-
pressed connexin-43 (Fig. 7d, e). This marks a positive
turning point for the printed patches and indicates future
possibilities for tissue engineering by this 3D bioprinting
platform technology. This technology employed for tissue
engineering and regenerative medicine could create avenues
for functional tissues and could create a clinical impact by
enhancing the quality of life for patients.
Briefly, we presented a 3D cell patterning platform that
allows efficient cell–matrix deposition with microscale spa-
tial resolution and uniform initial cell seeding density, while
maintaining cell viability over long-term culture. This high-
throughput system to print tissue constructs from micro-
droplets has the potential to enable future therapies by
providing (i) uniform cell seeding, (ii) 3D cell patterning
layer by layer, and (iii) viability over long-term culture.
We would like to thank The Randolph Hearst Founda-
tion and the department of Medicine, Brigham and Women’s
Hospital for the Young Investigators in Medicine Award.
Y.S., F.X., and U.D. were also partially supported by R21
(EB007707). This work was performed at the BAMM Labs at
the HST-Brigham and Women’s Hospital Center for Bioen-
gineering, Harvard Medical School.
No competing financial interests exist.
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Address correspondence to:
Utkan Demirci, Ph.D.
Bio-Acoustic MEMS in Medicine (BAMM) Laboratory
Center for Biomedical Engineering
Brigham and Women’s Hospital
Harvard Medical School
Cambridge, MA 02139
Received: March 16, 2009
Accepted: July 8, 2009
Online Publication Date: August 17, 2009
10MOON ET AL.