Bioreactor for modulation of cardiac microtissue phenotype by combined static stretch and electrical stimulation

Abstract

We describe here a bioreactor capable of applying electrical field stimulation in conjunction with static strain and on-line force of contraction measurements. It consisted of a polydimethylsiloxane (PDMS) tissue chamber and a pneumatically driven stretch platform. The chamber contained eight tissue microwells (8.05 mm in length and 2.5 mm in width) with a pair of posts (2.78 mm in height and 0.8 mm in diameter) in each well to serve as fixation points and for measurements of contraction force. Carbon rods, stimulating electrodes, were placed into the PDMS chamber such that one pair stimulated four microwells. For feasibility studies, neonatal rat cardiomyocytes were seeded in collagen gels into the microwells. Following 3 days of gel compaction, electrical field stimulation at 3-4 V cm(-1) and 1 Hz, mechanical stimulation of 5% static strain or electromechanical stimulation (field stimulation at 3-4 V cm(-1), 1 Hz and 5% static strain) were applied for 3 days. Cardiac microtissues subjected to electromechanical stimulation exhibited elevated amplitude of contraction and improved sarcomere structure as evidenced by sarcomeric α-actinin, actin and troponin T staining compared to microtissues subjected to electrical or mechanical stimulation alone or non-stimulated controls. The expression of atrial natriuretic factor and brain natriuretic peptide was also elevated in the electromechanically stimulated group.

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Bioreactor for modulation of cardiac microtissue phenotype by combined static stretch and
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2014 Biofabrication 6 024113
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Bioreactor for modulation of cardiac
microtissue phenotype by combined static
stretch and electrical stimulation
Jason W Miklas
1
, Sara S Nunes
2
, Aarash Sofla
1
, Lewis A Reis
1
,
Aric Pahnke
1,4
, Yun Xiao
1,4
, Carol Laschinger and Milica Radisic
1,2,3,4
1
Institute of Biomaterials and Biomedical Engineering, University of Toronto, Toronto, Ontario, Canada
2
Toronto General Research Institute, University Health Network, Toronto, Ontario, Canada
3
The Heart and Stroke/Richard Lewar Centre of Excellence, Toronto, Ontario, Canada
4
Department of Chemical Engineering and Applied Chemistry, University of Toronto, Toronto, Ontario,
Canada
E-mail: m.radisic@utoronto.ca
Received 13 September 2013, revised 27 April 2014
Accepted for publication 30 April 2014
Published 30 May 2014
Abstract
We describe here a bioreactor capable of applying electrical field stimulation in conjunction with
static strain and on-line force of contraction measurements. It consisted of a
polydimethylsiloxane (PDMS) tissue chamber and a pneumatically driven stretch platform. The
chamber contained eight tissue microwells (8.05 mm in length and 2.5 mm in width) with a pair
of posts (2.78 mm in height and 0.8 mm in diameter) in each well to serve as fixation points and
for measurements of contraction force. Carbon rods, stimulating electrodes, were placed into the
PDMS chamber such that one pair stimulated four microwells. For feasibility studies, neonatal
rat cardiomyocytes were seeded in collagen gels into the microwells. Following 3 days of gel
compaction, electrical field stimulation at 3–4Vcm
−1
and 1 Hz, mechanical stimulation of 5%
static strain or electromechanical stimulation (field stimulation at 3–4Vcm
−1
, 1 Hz and 5% static
strain) were applied for 3 days. Cardiac microtissues subjected to electromechanical stimulation
exhibited elevated amplitude of contraction and improved sarcomere structure as evidenced by
sarcomeric α-actinin, actin and troponin T staining compared to microtissues subjected to
electrical or mechanical stimulation alone or non-stimulated controls. The expression of atrial
natriuretic factor and brain natriuretic peptide was also elevated in the electromechanically
stimulated group.
SOnline supplementary data available from stacks.iop.org/BF/6/024113/mmedia
Keywords: cardiomyocyte, myocardium, tissue engineering, contraction force
(Some figures may appear in colour only in the online journal)
1. Introduction
Recent advances in the fields of stem cell biology [1–3] and
cardiac tissue engineering [4–6] enable us to create human
cardiac tissues in vitro [7,8]. These tissues can potentially be
used as platforms for drug testing or studies of cardiac phy-
siology and pathophysiology. However, to enable correct
utilization of these tissues in discovery studies, we need to
find a way to mature cardiac tissues in vitro, induce desired
disease phenotypes and enable on-line monitoring and
recording of functional outputs such as force of contraction.
Additionally, the resulting tissues should be composed of
relatively small numbers of cells to reduce costs associated
with high-throughput screening using cardiomyocytes derived
from human pluripotent stem cells (hPSC).
We have recently described a new platform for the
generation of mature cardiac tissues termed biological wire or
biowire [5]. The starting cell population were cardiomyocytes
1758-5082/14/024113+14$33.00 © 2014 IOP Publishing Ltd Printed in the UK1
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Biofabrication 6(2014) 024113 (14pp) doi:10.1088/1758-5082/6/2/024113

derived from either human embryonic stem cells or human
induced pluripotent stem cells. The combination of three-
dimensional (3D) cultivation in a micro-well and gel com-
paction around a suture enabled the creation of cardiac
organoids with remarkably well developed structural proper-
ties with cells aligned in parallel with one another containing
pronounced registers of sarcomeres. In addition, application
of electrical field stimulation with a progressive frequency
increase over 7 days of culture induced electrophysiological
changes consistent with cell maturation including up regula-
tion of I
K1
and hERG currents. However, terminal differ-
entiation was not achieved as demonstrated by the absence of
T-tubules and M-lines at the ultrastructural level.
These findings motivated us to explore additional options
that could result in terminal cell differentiation. Following
biomimetic principles, we aimed to recreate the environment
of the native heart tissue in a bioreactor in order to enable the
cells to acquire and maintain a differentiated adult-like phe-
notype. One of the stimuli that was most strikingly lacking in
our previous work was mechanical stimulation. In previous
studies, mechanical stimulation led to physiological hyper-
trophy but induced elongation of action potential duration and
did not seem to mature the calcium handling of the cardio-
myocytes [9]. Interestingly, electrical stimulation improved
cardiomyocyte calcium handling and also elicited a physio-
logical hypertrophic response, albeit at incomplete levels [5].
Mechanical stretch has been shown to induce an active force-
length relationship [4], similar to Frank–Starling curves
generated from intact hearts, which was not present in car-
diomyocytes derived from hPSC that were not subjected to
mechanical stimulation [10]. Additionally, as the levels of
load per cardiomyocyte are increased in the heart due to cell
death in disease states such as in hypertension, pathological
phenotypes of cardiomyocytes are induced and acquired.
Therefore, we hypothesized that developing a platform which
combined electrical and mechanical stimulation of cardiac
microtissues would enable us to modulate the cardiomyocyte
microenvironment to promote different levels of maturation
or pathological conditions in vitro.
Currently, there are a variety of costly and time-con-
suming assays that can be employed to determine the phe-
notype of cardiomyocytes and function of the cardiac tissue as
a whole. These include, confocal microscopy, transmission
electron microscopy, real time quantitative polymerase chain
reaction (RT-qPCR), immunohistochemistry, calcium tran-
sients mapping, optical mapping and patch clamping. In order
to obtain a complete picture of the state of an individual
cardiomyocyte in the tissue, all of these assays are required.
However, besides being time-consuming, some of these
assays (e.g. calcium transient recording and patch clamping)
require the tissue to be digested at the end time point with
plating of cells overnight which can possibly introduce errors.
As a result, a quick method that uses the bulk tissue to
determine changes consistent with maturation of hypertrophy
is needed. To address this, the platform developed here
included a method to measure functional properties, specifi-
cally through the presence of posts that can deflect under the
contractile force of the tissue as described in previous studies
[9,11–13]. Using two-dimensional (2D) analytical solutions
for a bending cantilever, the force of contraction of the tissue
can be calculated from images acquired using bright filed
microscopy and a camera. Additionally, the ability to acquire
the force on-line can provide an effective way for screening
different electromechanical regiments. Moreover, for the
cardiac microtissue to beat as a whole and to beat faster in
response to faster pacing the tissue must have functioning
excitation–contraction coupling machinery. Consequently, if
one assumes that as contractile force increases so does the
maturation state of the cardiac microtissue then, screening for
the force of contraction increase will also mean screening for
the conditions that bring about the highest degree of
maturation in the tissue. However, it should be noted that
there are instances where high force can be associated with an
immature, disease phenotype as found in pathological cardiac
hypertrophy.
We describe here a bioreactor that enables con-current
application of electrical and mechanical stimuli. Specifically,
electrical field stimulation was combined with static strain.
Cardiac microtissues were created by seeding cardiomyocytes
in a well created from polydimethylsiloxane (PDMS), using a
CNC (computer numerically controlled) machined mould,
followed by gel compaction around two built-in posts. After 3
days of gel compaction a regiment of mechanical and/or
electrical stimulation was applied for an additional 3 days.
This initial study was focused on designing and proving
feasibility of the bioreactor’s use, thus all experiments were
performed with cells derived from neonatal rat hearts. Future
studies will focus on the use of hPSC-derived
cardiomyocytes.
2. Methods
2.1. Neonatal rat heart isolation
Neonatal (1–2 day old) Sprague-Dawley rats were euthanized
according to the procedure approved by the University of
Toronto Committee on Animal Care. The hearts were
removed, quartered and the cells were isolated by an over-
night treatment with trypsin (4 °C, 6120 units mL
−1
in Hank’s
Balanced Salt Solution, HBSS) followed by serial collagenase
digestion (220 units mL
−1
in HBSS) as described in previous
work [14]. The supernatant from five collagenase digests of
the tissues was collected and centrifuged at 750 RPM (94 × g)
for 4 min, resuspended in culture medium and pre-plated into
T75 flasks (Falcon) for two 1 h intervals to separate the non-
adherent cells (enriched cardiomyocytes) from the adherent
cells (non-myocytes). The cells were cultivated in culture
medium consisting of Dulbecco’s modified Eagle medium
(DMEM) with 4.5 g L
−1
glucose, 4 mM L-glutamine, 10%
certified fetal bovine serum (FBS), 100 U mL
−1
penicillin,
100 μgmL
−1
streptomycin, and 10 mM 4-2-hydroxyethyl-1-
piperazineethanesulfonic acid buffer (HEPES, Gibco/
Invitrogen).
Biofabrication 6(2014) 024113 J W Miklas et al
2

2.2. Bioreactor platform design and manufacture
The bioreactor consisted of a PDMS tissue chamber capable
of housing eight microtissues, each in its own well and two
pairs of stimulating electrodes. Four microtissues were sti-
mulated by a pair of the stimulating electrodes. The PDMS
tissue chamber was then placed within a custom built stretch
platform. To fabricate the PDMS tissue chamber, a 3D
computer generated drawing of the mould was created using
AutoCad, figures 1(A), (B). Each microtissue well had a pair
of posts to serve as fixation points for the tissue and for
measurements of contraction force. The posts were designed
to have a high aspect ratio of 3.5, where the height of each
post was 2.78 mm and the diameter of each post was 0.8 mm.
The distance between the centers of the two posts was set at
6.80 mm. This value was chosen so that the length of a car-
diac microtissue would be about 7 mm. The AutoCad drawing
was loaded into a milling machine that created a negative
mould of the PDMS tissue chamber out of aluminum
figure 1(C). To create the final device, the aluminum negative
mould was filled with PDMS and allowed to cure at 80 °C for
2 h at which point the PDMS tissue chamber was removed
from the aluminum mould figure 1(D).
To provide mechanical stimulation, the PDMS tissue
chamber was inserted into a pneumatically driven stretch
device as shown in figure 1(E). The ends of the PDMS
platform were sandwiched between two metal plates that
served to anchor the PDMS tissue chamber platform to the
stretch device. Compressed air was pumped into the pneu-
matic pistons to move the two plates of the stretch device
apart stretching the entire PDMS platform. Finally, 3 mm
(0.12 inch) diameter carbon rods (Ladd Research Industries)
were inserted into the mould to provide the electrical field
stimulation and connected to a commercially available Grass
s88x stimulator as in our previous studies [5]. The displace-
ment of the posts upon application of different levels of strain
by the stretch platform was measured from microscopically
acquired images using ImageJ to validate the bioreactor
operating parameters.
2.3. Cell seeding
Cardiac microtissues were generated using enriched cardio-
myocytes derived from neonatal rat hearts as described above.
60 million cells mL
–1
were suspended in 3.0 mg mL
−1
col-
lagen type I gels [15–17] (BD Biosciences) with 2 mM
NaOH, 2.62 mM NaHCO
3
, in 1X M199 media with 15% of
the collagen gel solution consisting of growth factor reduced
Matrigel (BD Biosciences). 25 μL of the cell gel suspension
(1.5 × 10
6
cells) was pipetted into each of the PDMS micro-
tissue wells and placed into the incubator at 37 °C for 30 min
to allow the collagen hydrogel to gel. Microtissues were then
cultured for 3 days with culture media changes every other
day. Culture medium was the same as used in neonatal rat
heart isolation. Subsequent, mechanical stimulation, electrical
stimulation or a combination of electrical and mechanical
stimulation was applied for three additional days. The elec-
trical stimulation group had a pair of carbon electrodes placed
in the PDMS tissue chamber at a distance of 2 cm from one
another. The carbon electrodes were connected to a cardiac
stimulator (Grass s88x) with platinum wires (Ladd Research
Industries). Cardiac microtissues were situated perpendicular
to the electrodes and were submitted to electrical stimulation
(rectangular, biphasic, 1 ms, 3–4Vcm
−1
) at 1 Hz for 3 days.
These electrical field stimulation parameters have been shown
previously to work well in maturing rat neonatal cardio-
myocyte patches [14]. For the group where mechanical and
electrical stimulation was combined, the same electrical sti-
mulation regiment as described above was used. In addition,
to apply mechanical stimulation, the PDMS tissue chamber
was placed inside a custom made stretch platform figure 1(E).
The PDMS tissue chamber was clamped into the metal base
and by using compressed air to move a pair of pistons the
PDMS tissue platform was stretched by 5%. This 5% stretch
was transferred to the cardiac microtissues as they were
anchored to two posts in each well. The 5% stretch was held
constant for the duration of the 3 days of culture.
2.4. Post-deflection measurements
To determine the force of contraction of each EHT, a canti-
lever beam-partial uniform load equation was used. At the
end of cultivation, the PDMS tissue chamber was transferred
into a controlled environment, at 37 °C and 5% CO
2
,
microscope. To determine the force of contraction of each
cardiac microtissue, the top surface of a post for one micro-
tissue at a time was video recorded using CellSens software
(Olympus). Post deflection for each microtissue was deter-
mined at increasing rates of stimulation frequency, starting at
0.5 Hz and increasing to 3 Hz at 0.5 Hz increments. Each
tracking video was taken for 30 s with 5 s in between starting
a new tracking video at a higher frequency of pacing the
microtissues. Each video was then exported as a series of Tiff
images and opened in ImageJ. Using the ImageJ plugin
SpotTracker [18], the distance the post moved was deter-
mined. Multiple measurements were used and then averaged
to determine the average maximum post deflection, δmax, for
each cardiac microtissue at each frequency of pacing. The
elastic modulus of the PDMS was determined prior the post-
deflection measurements and an average obtained. The height
of the post and the height of the tissue on the post were
individually measured for each construct to ensure a precise
calculation for each cardiac microtissue could be determined.
Since the EHTs were not situated at the bottom of the
post, the method of superposition was implemented to
determine the distributed load on the cantilever beam
figure 2(B). To determine a partial uniform load at a specific
distance along the beam’s length, the subtraction of two dif-
ferent partial uniform distributed loads that all originate at the
fixed end are used to determine the ‘floating’distributed load.
The formula governing the method of superposition for a
cantilever beam-partial uniform deflection is shown in [19],
schematic of the cantilever beam is shown in figure 2(B), and
Biofabrication 6(2014) 024113 J W Miklas et al
3

Biofabrication 6(2014) 024113 J W Miklas et al
4
Figure 1. Bioreactor platform for combined electrical field and static stretch stimulation. (A) CAD drawing of a bioreactor well capable of
housing four cardiac microtissues and two pairs of stimulating electrodes. Each cardiac tissue microwell contains a pair of posts to be used for
monitoring of tissue contractions. (B) Three-dimensional CAD rendering of the bioreactor well used in the milling machine for production of
aluminum mold. (C) Aluminum mold produced by the milling process. (D) PDMS bioreactor wells produced using aluminum mold. (E)
PDMS bioreactor well placed in the pneumatically driven stretch device capable of providing static stretch.

variables for equation (1) are defined in table 1.
δ=−−
′−′
() ( )
qa La qa La
24EI 424EI 4(1)
max
33
In this case a=2L/3 and a′=L/3 [19].
To determine the force per cross-sectional area of each
cardiac microtissue, the force per length q, was converted into
a point load and divided by the cross-sectional area of the
cardiac microtissue. Each cross-sectional area of the cardiac
microtissue was individually measured using microscopy in
conjunction with ImageJ. Four points along the length of the
cardiac microtissue were measured and averaged to obtain the
diameter of the cardiac microtissue, assuming circular cross-
section, to determine each specific cardiac microtissue cross-
sectional area. Equation (2) was used to calculate force per
Biofabrication 6(2014) 024113 J W Miklas et al
5
Figure 2. Validation of the strains and force of contraction measurements in the PDMS well of the bioreactor platform. (A) Each cardiac
tissue microwell contains a pair of posts that deflects as the tissue contracts. The tissue is generated by gel compaction of cardiomyocytes in a
collagen gel (arrow). (B) To calculate force of contraction, a beam deflection analytical model can be used to correlate imaged deflection of
the post during a contraction cycle to force of contraction. If the tissue is not situated at the bottom of the post, a method of superposition can
be implemented to determine the distributed load on the post, where centrally positioned load (a) is modelled as a combination of two loads
(b) and (c). The images are redrawn based on a schematic presented in [19]. (C) Sensitivity analysis of post free end deflection with varying
tissue height along the post. A tissue of 0.4 mm thickness situated at the bottom of the post with an average point force of contraction of
0.2 mN was the base scenario (0% height change) in this graph. The tissue was then moved up the post and the deflection at the free end was
calculated again based on the same force of contraction. (D) Sensitivity analysis of post free end deflection with varying distributed load. A
tissue of 0.4 mm thickness situated at the bottom of the post with an average point force of contraction of 0.2 mN (0.5 N m
−1
) was the base
scenario at 0% change in distributed load. The distributed load was then varied while assuming the tissue remained at the bottom of the post.

unit area.
π
=qa
r
FPA (2)
2
Sensitivity analysis was performed to determine at which
magnitudes different factors influence the calculated force per
area measurements.
2.5. Validation of the force of contraction measurements
Validation of the calculated force required to move the PDMS
post was performed using direct measurements on the Refined
Myograph System (Kent Scientific). The PDMS post and part
of the base was excised from the PDMS platform and placed
on a glass slide. Since the Myograph has a caliper attached to
the force transducer, specific displacements were measured
while recording a change in voltage from the transducer,
which correlated to the force needed to displace the tip of a
single PDMS post. A displacement versus force graph was
then generated.
2.6. Measurements of elastic modulus of the PDMS
To determine the elastic modulus of the PDMS, a rectangular
strip of PDMS was created for tensile testing. Using an
ElectroForce 5200 BioDynamic Test Instrument (Bose) ten-
sile testing machine, a constant rate of displacement (1
mm min
−1
) and stretch distance was set. The software,
WinTest, was used to collect the displacement and the force
generated. Finally, the stress was calculated and a stress strain
curve was generated where the slope of the data set repre-
sented the elastic modulus. The curves were linear over the
entire range tested (R
2
= 0.99), up to 35% strain, thus at the
given composition our PDMS behaved as a linear elastic
material.
2.7. Excitation threshold (ET) and maximum capture rate
(MCR) measurements
After 6 days of culture, cardiac microtissues were assessed for
two functional parameters: ET and MCR. ET and MCR were
measurements performed in a controlled environment of
37 °C and 5% CO
2
. ET was determined as the lowest voltage
the cardiac microtissues could be continuously paced at 2 Hz.
MCR was determined as the highest frequency of continuous
pacing the microtissues could exhibit at 5–6Vcm
−1
.
2.8. Confocal microscopy
At the final time point of 6 days, cardiac microtissues were
fixed in 4% paraformaldehyde. Immunostaining was per-
formed as described previously [5] using the following anti-
bodies: mouse anti-cardiac troponin T (cTNT) (Abcam;
1 : 100), mouse anti-α-actinin (Abcam, 1 : 20), rabbit anti-
connexin 43 (Cx43) (Abcam, 1 : 500), goat anti-mouse-Alexa
Fluor 488 (Jackson ImmunoResearch; 1 : 400) and donkey
anti-rabbit-TRITC (Invitrogen; 1 : 400). Phalloidin (Invitro-
gen 1 : 100) was used to detect actin fibers. Cells were
visualized using a fluorescence confocal microscope (Zeiss
LSM-510).
2.9. qPCR
RT-qPCR was performed as previously described [5]. Total
RNA was prepared from cardiac microtissues after 6 days of
culture following the manufacturer’s protocol for TRIzol
Reagent (Invitrogen). RNA was reverse transcribed into
cDNA using random hexamers and Oligo (dT) with Super-
Script VILO (Invitrogen). RT-qPCR was performed on a
LightCycler 480 (Roche) using LightCycler 480 SYBR Green
I Master (Roche). Expression levels were normalized to the
housekeeping gene Glyceraldehyde 3-phosphate dehy-
drogenase (GAPDH). The oligonucleotide sequences are
summarized in supplemental table 1 (available at stacks.iop.
org/BF/6/024113/mmedia).
2.10. Western blotting
Cardiac microtissue protein was isolated after 6 days of cul-
ture following the manufacturer’s protocol for TRIzol
Reagent (Invitrogen). Once the protein was isolated, the
protein pellet was solubilized in a 1% SDS solution in de-
ionized water. Proteins were separated by electrophoresis in
Novex Tris-Glycine gels (Life technologies) and dry trans-
ferred using the iBlot (Life technologies) to a nitrocellulose
iBlot Transfer Stack (Life technologies). Membranes were
probed for Phospho-p44/42 MAPK (pERK1/2) (Cell Signal-
ing Technology), p44/42 MAPK (ERK1/2) (Cell Signaling
Technology), or GAPDH (Millipore) antibodies. Secondary
antibodies were peroxidase conjugated (DAKO). Membranes
were developed with ECL reagent LuminataClassico Sub-
strate (Millipore).
2.11. Statistical analysis
Statistical analysis was performed using SigmaPlot 12.0.
Differences between experimental groups were analyzed by
one or two-way ANOVA. For one-way ANOVA, Kolmo-
gorov–Smirnov was used for the normality test. For a failed
normality test during one-way ANOVA, Kruskal–Wallis one-
way ANOVA on Ranks was used. For two-way ANOVA, the
Shapiro–Wilk normality test and Holm–Sidak method for
Biofabrication 6(2014) 024113 J W Miklas et al
6
Table 1. Definition of variables used in post-deflection
measurements and force calculation.
Symbol Description Units
qForce per length N
m
aDistance of uniform load m
EElastic modulus N
m2
IArea of moments of inertia =π
(
)
circle r
4
4m
4
LLength of post m
δMaximum deflection at free end of beam m
FPA Force per area N
m2
rRadius m

pairwise multiple comparison procedures were used. P< 0.05
was considered significant for all statistical tests.
3. Results
3.1. Validation of the bioreactor platform operation
In order to have a uniform and controllable amount of static
stress applied to the cardiac microtissues, we needed to ensure
that the strain distribution within the PDMS microtissue wells
was uniform and equal in all the microtissue wells. Addi-
tionally, we needed to ensure that as the PDMS well was
stretched using the bioreactor platform, the distance between
the two posts in the cardiac microtissues increased equally in
different microtissue wells, and proportionally to the amount
of strain applied to the entire PDMS platform. This validation
was achieved by direct measurements from microscopically
acquired images.
As seen in table 2, the actual displacement of the posts
was shown as a percent change in the distance. For the 5%
and 10% strain applied to the PDMS well by the bioreactor
platform, the achieved displacement between the anchoring
posts was 4.6% and 10.5%, resulting in a relatively small
error. As the applied strain increased to 15% and 20%, the
actual displacement between the posts began to falter to
13.3% and 16.9% resulting in unacceptably large percent
error for the highest strain applied. Thus, we concluded that
the developed platform should be used in conjunction with
strains of up to 15%.
It was also necessary for us to confirm that the forces of
contraction calculated using the analytical solution of the
beam-deflection model was correct. This was achieved by
measuring, using an independent instrument, the amount of
force needed to bend the PDMS post by a certain amount.
Percent error in evaluation was then calculated based upon the
difference between the Myograph testing and the analytical
solution as seen in table 3with an average percent error
of 7.4 ± 4.1%.
Finally, a sensitivity analysis was performed to determine
the relationship between tissue height and deflection of the
post at the free end and to also examine the relationship
between a tissue’s force of contraction and the deflection of
the post at the free end. Figure 2(C) shows how the deflection
of the post at the free end changes based on how the height of
the tissue varies from the bottom of the post to the top of the
post. In this scenario it is assumed that the tissue thickness
was 0.4 mm and the percentage change in height was the
difference between the tissue’s new height along the post and
the ideal position of the tissue, which is when the tissue is
sitting at the bottom of the post. Clearly, the force of con-
traction calculation is very sensitive to the position of the
tissue along the post as the post deflects much easier under the
same load as the tissue is positioned at higher positions
(moving away from the base and towards the tip of the post).
Consequently, it is important to determine the tissue’s posi-
tion along the post each time post deflection imaging is per-
formed by determining the Z-displacement from the top of the
post to the tissue. On average, the height of the tissues
measured were 1.72 ± 0.36 mm above the base of the post
when force of contraction was assessed on day 7. This cor-
responds to a percent change in height from the base of the
post of 330% when referring to figure 2(C). For the post to be
deflected 1 μm at this height, assuming an average tissue
thickness of 0.4 mm (as in figures 2(C) and (D)), the average
distributed load would need to be 0.02 N m
−1
which corre-
sponds to a point load of 0.008 mN or 0.064 mN mm
−2
. The
smallest stress that we measured in our system was 0.093
mN mm
−2
. Furthermore, each tissue’s width needs to be
measured to determine the cross-sectional area to calculate
force per area as shown in figure 2(A).
Figure 2(D) shows how the force of contraction of a
cardiac microtissue, or the distributed load along the post,
affects the maximum deflection at the free end. In this sce-
nario, the original distributed load was set to be 0.5 N m
−1
,
which is a point-load of 0.2 mN. At 0.2 mN the free end will
deflect about 1 μm, which is the lower limit of the
Biofabrication 6(2014) 024113 J W Miklas et al
7
Table 2. Validation of the relationship between the measured strain
in cardiac microtissue wells (i.e. distance between posts) and strain
applied to the entire PDMS well. The PDMS well was stretched
using the constructed stretching bioreactor. Images were taken of the
PDMS platform at each of the four strain settings of 5, 10, 15 and
20%. The actual distance between the posts was measured and
percentage error between the desired stretch and the average actual
stretch was calculated as follows:
= ∗
−
Percent error 100
Measured strain Applied strain
Applied strain .
Average measured
strain (%)
Applied
strain (%)
Percent
error (%)
4.7 ± 0.1 5 9.8 ± 2.9
10.5 ± 0.9 10 7.7 ± 4.1
13.3 ± 0.2 15 12.5 ± 2.0
16.9 ± 0.6 20 18.4 ± 4.4
Table 3. Comparison between direct measurements of the force
required to deflect a PDMS post and calculations based on the beam
deflection model. The final column shows the percent error between
the two methods calculated as follows:
= ∗
−
Percent error 100
Measured force Calcualted force
Calculated force . Displacement is the
distance the tip of a PDMS post was moved and the myograph force
is the corresponding force recorded from the device required to move
the PDMS post.
Displacement
(μm)
Myograph force
(mN)
Analytical
solution
force (mN)
Percent
error
(%)
0 0.00 0.00 0.0
84 0.24 0.26 9.5
98 0.31 0.31 1.3
112 0.35 0.35 0.9
179 0.63 0.56 12.1
206 0.71 0.65 8.7
238 0.82 0.75 9.9

displacement that could be measured using this set-up due to
the optical transparency limitations of the PDMS and the
tissue.
3.2. Cardiac microtissues generated in the bioreactor platform
Our main goal here was to determine if the newly developed
bioreactor platform could support survival of cardiomyocytes
and their assembly into contractile tissue, as well as to
determine if the differences between electrical field stimula-
tion alone compared to the combined electromechanical sti-
mulation regimen could be detected. We therefore used
cardiomyocytes derived from neonatal rat hearts to generate
cardiac microtissues in non-stimulated, non-stretched con-
trols; electrical field stimulation alone, mechanical stimulation
alone, or in the presence of both electrical field stimulation
and static stretch. To allow cells to elongate and remodel the
matrix first, stimulation regimens were initiated after 3 days
of stimulation-free pre-culture. Initiating stimulation too early
was found detrimental in our previous work [14] and would
likely lead to rupture of the tissue here. At the end of culture,
there were no significant differences in electrical excitability
parameters, ET and MCR, between the three groups
(figures 3(A), (B)). However, force of contraction was
increased in the group that was subject to con-current elec-
tromechanical stimulation and found to be statistically sig-
nificant at 0.5 Hz (P= 0.003), 1.0 Hz (P= 0.004), 1.5 Hz
(P= 0.006), 2.0 Hz (p= 0.0017) and 2.5 Hz (P= 0.028) in
comparison to the 5% strain group (figure 3(C)). Repre-
sentative traces of each condition being paced at frequencies
between 0.5–3 Hz are shown in supplemental figure 1
(available at stacks.iop.org/BF/6/024113/mmedia).
Immunostaining demonstrated that cardiac microtissues
subjected to concurrent electromechanical stimulation exhib-
ited well developed sarcomeric structures in comparison to
those subjected to electrical field stimulation alone, mechan-
ical strain alone or to the control cardiac microtissues. Double
staining for sarcomeric α-actinin and actin revealed well
developed registers of sarcomeres in the 5% strain +1 Hz
group, whereas this organization was not as apparent in the
other three groups (figure 4(A)). Furthermore, the 5% strain
group displayed poor myofibril formation with some myofi-
brils aligning perpendicular to the static strain direction in
comparison to previous cardiac tissue engineered studies
showing cyclic stretching (10%, 2 Hz) [20] or auxotonic loads
[21] (supplemental figure 2, available at stacks.iop.org/BF/6/
024113/mmedia). This poor response is typically found in 2D
static and cyclic strain cultures as cardiomyocytes rearrange
themselves to elongate in a transverse direction to the strain,
i.e. to experience less strain [22,23]. This result may also
account for the lower force produced by the 5% strain con-
dition. Double staining for cTnT and Cx43, revealed well
defined cross-striation of cTnT in the 5% strain +1 Hz group.
The gap-junctional protein, Cx43, was present in all condi-
tions (figure 4(B)).
Sarcomeric proteins α-myosin heavy chain (MHC) and β-
MHC are differentially regulated in mammalian hearts during
development. In rodents, α-MHC is upregulated compared to
β-MHC during maturation while, the opposite holds true for
humans [12]. Quantitative PCR demonstrated similar α-
MHC/β-MHC ratios in all four conditions with the 5% strain
group being the lowest, which indicated that at the current
conditions, no appreciable shift towards maturation was
occurring over 6 days of total culture of neonatal rat cardiac
microtissues (figure 5(A)). The lower average α/βMHC ratio
in the 5% strain group may point towards a pathological state
[24]. Sarco/endoplasmic reticulum Ca
2+
ATPase (SERCA) is
an ion pump found in the membrane of the sarcoplasmic
reticulum that is responsible for pumping Ca
2+
back from the
cytoplasm into the sarcoplasmic reticulum at the completion
of a contraction cycle in mature cardiomyocytes [10,25].
There were no significant differences in SERCA expression
amongst the three groups (figure 5(B)) in cardiac micro-
tissues. Atrial natriuretic factor (ANF) and brain natriuretic
peptide (BNP) are molecules highly expressed in ventricular
cardiomyocytes in the late stages of rodent and human heart
development [26]. In the post-natal and adult stages, the re-
expression of these proteins is consistent with pathological
hypertrophy [27]. The fact that ANF was highly expressed in
the 5% strain + 1 Hz and BNP was highly expressed in the
5% strain and 5% strain +1 Hz groups might indicate that
these cells were at the on-set of the pathological hypertrophy
process (figures 5(C), (D)) however no clear signs of sarco-
mere destruction were observed in the immunostaining sam-
ples. To gauge the levels of cell apoptosis versus survival in
different groups we examined the expression levels of
mRNAs for proteins in the B-cell lymphoma family (Bcl)
[12]. There were no significant differences in the ratio of pro-
apoptotic Bax to anti-apoptotic Bcl-2 in the four groups,
indicating that electrical field stimulation, mechanical stimu-
lation, concurrent electromechanical stimulation and the
control non-stimulated condition were all equally conducive
to survival of the cells in PDMS wells.
To further investigate the induction of a hypertrophic
response with stimulation in the bioreactor platform we
investigated the levels of ERK1/2 and phosphorylation levels
of ERK1/2 as phospho ERK1/2 has been previously shown to
be associated with physiological hypertrophy [28]. As shown
in figure 6, total levels of ERK1/2 were comparable amongst
the three groups and the levels of phospho ERK1/2 were
slightly although not significantly elevated in either 1 Hz, 5%
strain or the 5% strain +1 Hz condition compared to the
control condition. The ratio of phospho ERK/ERK was
comparable in all conditions except 5%, which was statisti-
cally significant in comparison to control (P= 0.032).
4. Discussion
Native heart muscle consists of aligned cardiomyocytes that
contract synchronously as a result of rapid action potential
propagation between adjacent cardiomyocytes via gap junc-
tions. This coordinated action of cardiomyocytes results in
pumping and expulsion of blood from the ventricles. As the
ventricles relax, they are filled with blood causing the ven-
tricular volume to expand and the ventricular wall to stretch.
Biofabrication 6(2014) 024113 J W Miklas et al
8

Thus, at each contraction cycle in vivo, cardiomyocytes
contract against the load (i.e. blood filling the ventricles) and
the electrical stimulus arises when the ventricular wall is in
the stretched state. Ventricular cardiomyocytes have a number
of stretch activated ion channels that are important regulators
of cardiomyocyte functionality [29]. The importance of
physical stretch was realized in the earliest cardiac tissue
engineering studies [12,20,30–32] and early cardiac tissues
based on neonatal rat cardiomyocytes cultivated in the pre-
sence of cyclic mechanical stretch. Zimmermann and
Eschenhagen demonstrated that when neonatal rat cardio-
myocytes were stimulated at 10% stretch and a frequency of
2 Hz, they could be pushed to a mature phenotype similar to
the one found in the native adult heart as evidenced by the
presence of Z, I, A and H bands in the sarcomeres as well as
T-tubules [30]. Additionally, cyclic stretching led to physio-
logical hypertrophy as evidenced by increased
cardiomyocytes size, increased percentage of binucleated
cardiomyocytes and upregulation of the ventricular isoform of
myosin light chain [12]. Engineered cardiac tissues that were
cultivated under auxotonic load were implanted in a rat model
of myocardial infarction mitigating heart failure [21]. This
study also conclusively demonstrated integration of rat engi-
neered heart tissues with the native rat myocardium [21].
Mechanical loading was also used to stimulate cardio-
myocytes derived from hPSC. Since cyclic stretch was able to
mature neonatal rat cardiomyocytes to an adult like stage,
Tulloch et al [4] implemented a similar set-up using cyclic
stretch to try and mature hPSC-derived cardiomyocytes.
Similarly, cyclic stretch promoted a pro-hypertrophic
response in these cells as illustrated by increased cell align-
ment parallel to the mechanical loading force, increased DNA
synthesis, increased cardiomyocyte area, and induction of
βMHC, cTnT, L-type calcium channel, ryanodine receptor,
Biofabrication 6(2014) 024113 J W Miklas et al
9
Figure 3. Functional properties of cardiac microtissues. (A) Excitation threshold determined at the end of cultivation as a minimum voltage
required to induce synchronous contraction. (B) Maximum capture rate determined at the end of cultivation as the maximum tissue beating
frequency. (C) Force of contraction. Control-cardiac microtissues cultivated in the PDMS wells without electrical or mechanical stimulation.
1 Hz cardiac microtissues cultivated in the PDMS wells in the presence of electrical field stimulation at 1 Hz 5% cardiac mictotissues
cultivated in the PDMS stretched at 5% static strain without electrical stimulation. 5% strain +1 Hz cardiac microtissues stretched at 5% static
strain and concurrently subjected to electrical field stimulation at 1 Hz. * denotes statistical significant by two-way ANOVA between 5% and
5% +1 Hz groups at specific pacing frequencies. Data represented as average ± standard deviation, N=3.

and SERCA mRNA compared to the constructs that were
cultivated in the absence of loading [4].
Interestingly, Kensah et al [33] found that cyclic stretch
(10%; 1 Hz for 7 days) did not improve contractile function or
morphology of their cardiac tissue engineered constructs in
comparison to static stretch. Instead of cyclic stretch, they
generated a protocol that gradually increased the static strain
of their constructs over 14 days with increases in static strain
occurring every second day in an attempt to recapitulate the
increasing systolic and diastolic pressure in the developing
heart. Similar to our findings, they did not see a statistically
significant increase in maximum active force of their gradu-
ally increasing static strain group in comparison to their
control. They did not see an increase in BNP or ANF gene
expression in their gradually increasing static strain group
[33]. Yet, in their gradually increasing static strain group, they
did have aligned sarcomeres parallel to the stretching force
while we found that our large single increase in static stress
resulted in cardiomyocytes elongating perpendicular to the
stretching direction, most likely in an attempt to reduce the
strain on their system. This could also account for the
decreased force of contraction that was observed, albeit not
statistically significant to control.
While these results with cyclic stretch alone were
encouraging, there was scarce evidence that mechanical sti-
mulation alone was sufficient to mature certain aspects of the
calcium handling machinery and induce appropriate expres-
sion and function of diverse ion channels required for cardiac
function. Engineered heart tissues generated from hPSC
derived cardiomyocytes displayed abnormally long action
potential durations (up to 1200 ms) and a resting membrane
potential of −49.1 mV [9] which is less negative than the
resting membrane potential of comparable 7–8 week old
embryoid bodies that resulted in cardiomyocytes with resting
membrane potential of −60.7 mV.
Interestingly, mechanical stimulation could also be pro-
vided by a compressive fluid flow as we [34] and others [35]
have demonstrated previously. When mechanical compres-
sion was provided together with fluid shear instead of
stretching in a static vessel to stimulate neonatal rat cardio-
myocytes, an intermittent compression regiment was able to
preserve α-actinin and N-cadherin expression and improve
Cx43 expression compared to non-compressed controls [35].
Fluid shear could also induce a physiological hypertrophic
response, mediated through the ERK1/2 signaling pathway,
as evidenced by upregulation of protein synthesis [28].
One of the other major parameters that has been shown to
affect functionality of engineered heart tissues is cell align-
ment. Many cardiac tissue engineering studies relied upon gel
compaction, of either fibrin or collagen gels, to generate
engineered heart tissues with aligned cardiomyocytes. Passive
tensile force was usually generated in these systems by two
anchoring posts or using a cylindrical non-adhesive mold.
Black et al [36] systematically showed that engineered heart
tissues which undergo gel compaction contain well aligned
cells. This increased cell alignment in turn prompted an
increased phosphorylated state of Cx43. Both of these factors
resulted in higher forces of contraction in the aligned engi-
neered heart tissues in comparison to those that had randomly
oriented cells [36]. This finding shows the importance of
anisotropy in engineered myocardium and the importance of
cardiomyocyte elongation with tension for proper maturation.
Optimizing and enhancing mechanical stimulation platforms
Biofabrication 6(2014) 024113 J W Miklas et al
10
Figure 4. Immunostaining of cardiac microtissues for sarcomeric and
gap junctional proteins. (A) Double staining for sarcomeric α-actinin
(green) and actin red. (B) Double staining for cardiac troponin T
(green) and connexin-43 red. Control-cardiac microtissues cultivated
in the PDMS wells without electrical or mechanical stimulation.
1 Hz: cardiac microtissues cultivated in the PDMS wells solely in the
presence of electrical field stimulation at 1 Hz. 5% strain: cardiac
microtissues cultivated in the PDMS wells solely in the presence of
5% static strain. 5% strain +1 Hz: cardiac microtissues stretched at
5% static strain and concurrently subjected to electrical field
stimulation at 1 Hz.

is an area of active research as evidenced by recent reports of
novel bioreactors capable of mechanical stimulation [37,38].
We first started using electrical field stimulation in car-
diac tissue engineering using cardiomyocytes derived from
neonatal rat hearts. The cells were seeded into porous col-
lagen scaffold and subjected to field stimulation for 5 days
after a pre-culture for 3 days allowed the cells to elongate in
the matrix. Electrically stimulated samples were able to form
well aligned registers of sarcomeres and exhibit defined M
and Z lines and H, I, and A bands [14]. Recently, we used
electrical stimulation of progressive frequency increase to
mature cardiomyocytes derived from hPSC cultivated in
microstructures termed biowire [5].
Our goal here was to develop a platform that can provide
combined electromechanical stimulation and prove the fea-
sibility of its operation. Electromechanical stimulation of
scaffold-based engineered tissues have recently been reported
by Wang et al [39] and Morgan et al [40]. However, the
platform designed by Wang et al was only validated using
mesenchymal stem cells as the cell source and both afore-
mentioned designs did not utilize on-line force readout. Liao
et al created an electromechanical stimulation platform that
had online force of contraction measuring, however, only one
cardiac construct could be created and measured per bior-
eactor. The platform described here provides a higher
throughput system, eight cardiac microtissues per PDMS
platform, in comparison to other platforms that usually can
only accommodate one or two constructs per platform
[37,38,41]. While our platform is similar to Boudou et al
[13] they did not mechanically stretch their cardiac constructs
to observe the effects of combined mechanical strain and
electrical stimulation.
One of the main aims of this work was to demonstrate
that cardiomyocytes can survive, function and form a
Biofabrication 6(2014) 024113 J W Miklas et al
11
Figure 5. Quantitative polymerase chain reaction analysis of gene expression in cardiac microtissues. (A) Ratio of α-myosin heavy chain
(MHC) to β-myosin heavy chain, (B) Sarco/endoplasmic reticulum Ca
2+
ATP-ase (SERCA), (C) Atrial natriuretic factor (ANF), (D) Brain
natriuretic peptide (BNP), (E) Ratio of pro-apoptotic Bax to anti-apoptotic Bcl-2. Data represented as averages ± standard deviation, N=3.

contractile tissue in the developed bioreactor platform. We
therefore focused on the use of readily available neonatal rat
heart cells, while the use of cardiomyocytes derived from
hPSC will be reserved for future studies. In future studies,
electromechanical stimulation protocols will be optimized
with these cells and the possible parameter space may include
intermitted application of electrical stimulation on stretched
samples, application of auxotonic or cyclic mechanical sti-
mulation in conjunction with electrical stimulation as well as
switching from field to point stimulation. We have previously
extensively utilized electrical field stimulation in cardiac tis-
sue engineering [5,42], thus we wanted to evaluate here if the
application of electromechanical stimulation will result in
additional changes in the cardiac microtissues compared to
the application of electrical or mechanical stimulation alone.
The values of electrical excitability parameters obtained
here (figures 3(A), (B)) were in the same range as those we
previously reported [42], however the differences between the
control and stimulated groups were not apparent, likely due to
the short stimulation time (3 days) in comparison to longer
stimulation times (5–21 days) we used in previous studies
[5,42,43]. As expected, force of contraction increased with
the presence of electromechanical stimulation (figure 3(C)).
While the force measurements we obtained were not as high
as previously reported groups [44], when culture time is taken
into consideration our force values (0.05–0.15 mN at day 6)
match well with previous reports of engineered heart tissue
Biofabrication 6(2014) 024113 J W Miklas et al
12
Figure 6. Western blotting for extracellular signal regulated kinases (ERK1/2) expression and phosphorylation. * denotes statistical
significance (P= 0.032) between 5% strain and control group using one-way ANOVA. Data represented as averages ± standard
deviation, N=3.

cultured for similar times (0.1 mN at day 8) [9,11,45]. It is
possible that after two weeks of culture the force generated by
the cardiac micro tissues presented in this paper may match
the force generated by Hirt et al [11]. We did not observe a
positive force-frequency relationship (figure 3(C)). We are
convinced that this is due to a short total cultivation time (6
days) and a short stimulation time (3 days). Previous
mechanical stimulation studies have also been unable to
observe a positive force frequency relationship due to
increased pacing rate that is found in adult mouse ventricular
cardiomyocytes [46], however, they do show a positive active
force-length relationship during increasing strain (Frank–-
Starling mechanism) at time points of 9–10 days [30] and 14
days [20].
Further analysis is required to determine if the improved
sarcomere structure observed in the 5% strain +1 Hz condi-
tion (figure 4) is truly a result of physiological hypertrophy
alone or if the pathological hypertrophy process is being
initiated in this group based on the upregulation of ANF and
BNP (figures 5(C), (D)). Since the onset of pathological
hypertrophy takes longer than 3 days, future studies of longer
stretch time periods will explore this disease state. Moreover,
the increased ratio of pERK1/2 to ERK1/2 in the 5% strain
group contrasted to the low force of contraction, possibly due
to poor alignment of myofibrils (figures 3and 6). The
developed bioreactor platform enables us to optimize the
modalities of electromechanical stimulation to achieve either
physiological or pathological hypertrophy for future studies.
Importantly, none of the stimulation modalities applied here
were detrimental in terms of enhancing cell apoptosis as
evidenced by the unchanged ratio of Bax/Bcl2 in the three
groups (figure 5(E)). Overall, we described here a new bior-
eactor that can be used to study biological phenomena, such
as cardiomyocyte hypertrophy, with further studies needed to
delineate mechanistic effects of electro-mechanical
stimulation.
5. Conclusion
We developed a bioreactor that enabled us to apply con-
current electrical and static strain stimulation to cardiac
microtissues. The operation of the bioreactor was validated by
the cultivation of cardiac microtissues based on neonatal rat
cardiomyocytes. The tissue compacted over 3 days of culture
and an additional 3 days of stimulation resulted in the
improved sarcomere structure and increased force in the
group subjected to electromechanical stimulation. Future
studies are required to optimize the bioreactor for either
pathological or physiological hypertrophy in cardiomyocytes
derived from hPSC cells.
Acknowledgements
The authors would like to thank the University of Toronto
MIE Machine shop for their assistance in building the stretch
platform. This work was funded by grants from Ontario
Research Fund–Global Leadership Round 2 (ORF-GL2),
National Sciences and Engineering Research Council of
Canada (NSERC) Strategic Grant (STPGP 381002-09),
Canadian Institutes of Health Research (CIHR) Operating
Grant (MOP-126027), NSERC-CIHR Collaborative Health
Research, Grant (CHRPJ 385981-10), NSERC Discovery
Grant (RGPIN 326982-10), NSERC Discovery Accelerator
Supplement (RGPAS 396125-10) and National Institutes of
Health grant (2R01 HL076485).
References
[1] Yang L et al 2008 Human cardiovascular progenitor cells
develop from a KDR+ embryonic-stem-cell-derived
population Nature 453 524–8
[2] Zhang J, Wilson G F, Soerens A G, Koonce C H, Yu J,
Palecek S P, Thomson J A and Kamp T J 2009 Functional
cardiomyocytes derived from human induced pluripotent
stem cells Circ. Res. 104 e30–41
[3] Didie M et al 2013 Parthenogenetic stem cells for tissue-
engineered heart repair J. Clin. Invest. 123 1285–98
[4] Tulloch N L, Muskheli V, Razumova M V, Korte F S,
Regnier M, Hauch K D, Pabon L, Reinecke H and
Murry C E 2011 Growth of engineered human myocardium
with mechanical loading and vascular coculture Circ. Res.
109 47–59
[5] Nunes S S et al 2013 Biowire: a platform for maturation of
human pluripotent stem cell-derived cardiomyocytes Nat.
Methods 10 781–7
[6] Caspi O, Lesman A, Basevitch Y, Gepstein A, Arbel G,
Habib I H, Gepstein L and Levenberg S 2007 Tissue
engineering of vascularized cardiac muscle from human
embryonic stem cells Circ. Res. 100 263–72
[7] Chiu L L, Iyer R K, Reis L A, Nunes S S and Radisic M 2012
Cardiac tissue engineering: current state and perspectives
Front Biosci. (Landmark Ed) 17 1533–50
[8] Nunes S S, Song H, Chiang C K and Radisic M 2011 Stem
cell-based cardiac tissue engineering J. Cardiovasc. Transl.
Res. 4592–602
[9] Schaaf S et al 2011 Human engineered heart tissue as a
versatile tool in basic research and preclinical toxicology
PLoS One 6e26397
[10] Binah O, Dolnikov K, Sadan O, Shilkrut M, Zeevi-Levin N,
Amit M, Danon A and Itskovitz-Eldor J 2007 Functional
and developmental properties of human embryonic stem
cells-derived cardiomyocytes J. Electrocardiol. 40 S192–6
[11] Hirt M N et al 2012 Increased afterload induces pathological
cardiac hypertrophy: a new in vitro model Basic Res.
Cardiol. 107 307
[12] Tiburcy M et al 2011 Terminal differentiation, advanced
organotypic maturation, and modeling of hypertrophic
growth in engineered heart tissue Circ. Res. 109
1105–14
[13] Boudou T, Legant W R, Mu A, Borochin M A, Thavandiran N,
Radisic M, Zandstra P W, Epstein J A, Margulies K B and
Chen C S 2012 A microfabricated platform to measure and
manipulate the mechanics of engineered cardiac
microtissues Tissue Eng. A18 910–9
[14] Radisic M, Park H, Shing H, Consi T, Schoen F J, Langer R,
Freed L E and Vunjak-Novakovic G 2004 Functional
assembly of engineered myocardium by electrical
stimulation of cardiac myocytes cultured on scaffolds Proc.
Natl. Acad. Sci. USA 101 18129–34
[15] Nunes S S, Greer K A, Stiening C M, Chen H Y, Kidd K R,
Schwartz M A, Sullivan C J, Rekapally H and Hoying J B
Biofabrication 6(2014) 024113 J W Miklas et al
13

2010 Implanted microvessels progress through distinct
neovascularization phenotypes Microvasc. Res. 79 10–20
[16] Nunes S S, Krishnan L, Gerard C S, Dale J R, Maddie M A,
Benton R L and Hoying J B 2010 Angiogenic potential of
microvessel fragments is independent of the tissue of origin
and can be influenced by the cellular composition of the
implants Microcirculation 17 557–67
[17] Nunes S S, Rekapally H, Chang C C and Hoying J B 2011
Vessel arterial-venous plasticity in adult neovascularization
PLoS One 6e27332
[18] Sage D, Neumann F R, Hediger F, Gasser S M and Unser M
2005 Automatic tracking of individual fluorescence
particles: application to the study of chromosome dynamics
IEEE Trans. Image Process 14 1372–83
[19] Gere J M and Goodno B J 2012 Mechanics of Materials, SI
Edition 8th edn (Stanford, CT: Cengage Learning)
[20] Zimmermann W H, Schneiderbanger K, Schubert P, Didie M,
Munzel F, Heubach J F, Kostin S, Neuhuber W L and
Eschenhagen T 2002 Tissue engineering of a differentiated
cardiac muscle construct Circ. Res. 90 223–30
[21] Zimmermann W H et al 2006 Engineered heart tissue grafts
improve systolic and diastolic function in infarcted rat hearts
Nat. Med. 12 452–8
[22] Salameh A, Wustmann A, Karl S, Blanke K, Apel D,
Rojas-Gomez D, Franke H, Mohr F W, Janousek J and
Dhein S 2010 Cyclic mechanical stretch induces
cardiomyocyte orientation and polarization of the gap
junction protein connexin43 Circ. Res. 106 1592–602
[23] Matsuda T, Takahashi K, Nariai T, Ito T, Takatani T,
Fujio Y and Azuma J 2005 N-cadherin-mediated cell
adhesion determines the plasticity for cell alignment in
response to mechanical stretch in cultured cardiomyocytes
Biochem. Biophys. Res. Commun. 326 228–32
[24] Iemitsu M, Miyauchi T, Maeda S, Sakai S, Kobayashi T,
Fujii N, Miyazaki H, Matsuda M and Yamaguchi I 2001
Physiological and pathological cardiac hypertrophy induce
different molecular phenotypes in the rat Am. J. Physiol.
Regul. Integr. Comp. Physiol. 281 R2029–36
[25] Dolnikov K, Shilkrut M, Zeevi-Levin N, Gerecht-Nir S,
Amit M, Danon A, Itskovitz-Eldor J and Binah O 2006
Functional properties of human embryonic stem cell-derived
cardiomyocytes: intracellular Ca
2+
handling and the role of
sarcoplasmic reticulum in the contraction Stem Cells 24
236–45
[26] Razeghi P, Young M E, Alcorn J L, Moravec C S,
Frazier O H and Taegtmeyer H 2001 Metabolic gene
expression in fetal and failing human heart Circulation 104
2923–31
[27] Kong S W, Bodyak N, Yue P, Liu Z, Brown J, Izumo S and
Kang P M 2005 Genetic expression profiles during
physiological and pathological cardiac hypertrophy and
heart failure in rats Physiol. Genomics 21 34–42
[28] Dvir T, Levy O, Shachar M, Granot Y and Cohen S 2007
Activation of the ERK1/2 cascade via pulsatile interstitial
fluid flow promotes cardiac tissue assembly Tissue Eng. 13
2185–93
[29] Zipes D P and Jalife J 2009 Cardiac Electrophysiology: from
Cell to Bedside (Philadelphia, PA: Saunders Elsevier)
[30] Zimmermann W H, Fink C, Kralisch D, Remmers U,
Weil J and Eschenhagen T 2000 Three-dimensional
engineered heart tissue from neonatal rat cardiac myocytes
Biotechnol. Bioeng. 68 106–14
[31] Akhyari P, Fedak P W, Weisel R D, Lee T Y, Verma S,
Mickle D A and Li R K 2002 Mechanical stretch regimen
enhances the formation of bioengineered autologous cardiac
muscle grafts Circulation 106 I137–42
[32] Boublik J, Park H, Radisic M, Tognana E, Chen F, Pei M,
Vunjak-Novakovic G and Freed L E 2005 Mechanical
properties and remodeling of hybrid cardiac constructs made
from heart cells, fibrin, and biodegradable, elastomeric
knitted fabric Tissue Eng. 11 1122–32
[33] Kensah G et al 2013 Murine and human pluripotent stem cell-
derived cardiac bodies form contractile myocardial tissue
in vitro Eur. Heart J. 34 1134–46
[34] Brown M A, Iyer R K and Radisic M 2008 Pulsatile perfusion
bioreactor for cardiac tissue engineering Biotechnol. Prog.
24 907–20
[35] Shachar M, Benishti N and Cohen S 2012 Effects of
mechanical stimulation induced by compression and
medium perfusion on cardiac tissue engineering Biotechnol.
Prog. 28 1551–9
[36] Black L D 3rd, Meyers J D, Weinbaum J S, Shvelidze Y A and
Tranquillo R T 2009 Cell-induced alignment augments
twitch force in fibrin gel-based engineered myocardium via
gap junction modification Tissue Eng. A15 3099–108
[37] Nguyen M D, Tinney J P, Yuan F, Roussel T J, El-Baz A,
Giridharan G, Keller B B and Sethu P 2013 Cardiac cell
culture model as a left ventricle mimic for cardiac tissue
generation Anal. Chem. 85 8773–9
[38] Kensah G et al 2011 A novel miniaturized multimodal
bioreactor for continuous in situ assessment of bioartificial
cardiac tissue during stimulation and maturation Tissue Eng.
Part C Methods 17 463–73
[39] Wang B, Wang G, To F, Butler J R, Claude A,
McLaughlin R M, Williams L N, de Jongh Curry A L and
Liao J 2013 Myocardial scaffold-based cardiac tissue
engineering: application of coordinated mechanical and
electrical stimulations Langmuir 29 11109–17
[40] Morgan K Y and Black L D 3rd 2014 Mimicking isovolumic
contraction with combined electromechanical stimulation
improves the development of engineered cardiac constructs
Tissue Eng. A20 1654–67
[41] Feng Z G, Matsumoto T, Nomura Y and Nakamura T 2005 An
electro-tensile bioreactor for 3D culturing of cardiomyocytes
—A bioreactor system that simulates the myocordium’s
electrical and mechanical response in vivo IEEE Eng. Med.
Biol. Mag. 24 73–9
[42] Chiu L L, Iyer R K, King J P and Radisic M 2011 Biphasic
electrical field stimulation aids in tissue engineering of
multicell-type cardiac organoids Tissue Eng. A17 1465–77
[43] Iyer R K, Odedra D, Chiu L L, Vunjak-Novakovic G and
Radisic M 2012 Vascular endothelial growth factor secretion
by nonmyocytes modulates connexin-43 levels in cardiac
organoids Tissue Eng. A18 1771–83
[44] Zhang D, Shadrin I Y, Lam J, Xian H Q, Snodgrass H R and
Bursac N 2013 Tissue-engineered cardiac patch for
advanced functional maturation of human ESC-derived
cardiomyocytes Biomaterials 34 5813–20
[45] Naito H, Melnychenko I, Didie M, Schneiderbanger K,
Schubert P, Rosenkranz S, Eschenhagen T and
Zimmermann W H 2006 Optimizing engineered heart tissue
for therapeutic applications as surrogate heart muscle
Circulation 114 I72–8
[46] Lim C C, Apstein C S, Colucci W S and Liao R 2000 Impaired
cell shortening and relengthening with increased pacing
frequency are intrinsic to the senescent mouse
cardiomyocyte J. Mol. Cell Cardiol. 32 2075–82
Biofabrication 6(2014) 024113 J W Miklas et al
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