Biowire: A platform for maturation of human pluripotent stem cell-derived cardiomyocytes

Article (PDF Available)inNature Methods 10(8) · June 2013with 337 Reads
DOI: 10.1038/nmeth.2524 · Source: PubMed
Abstract
Directed differentiation protocols enable derivation of cardiomyocytes from human pluripotent stem cells (hPSCs) and permit engineering of human myocardium in vitro. However, hPSC-derived cardiomyocytes are reflective of very early human development, limiting their utility in the generation of in vitro models of mature myocardium. Here we describe a platform that combines three-dimensional cell cultivation with electrical stimulation to mature hPSC-derived cardiac tissues. We used quantitative structural, molecular and electrophysiological analyses to explain the responses of immature human myocardium to electrical stimulation and pacing. We demonstrated that the engineered platform allows for the generation of three-dimensional, aligned cardiac tissues (biowires) with frequent striations. Biowires submitted to electrical stimulation had markedly increased myofibril ultrastructural organization, elevated conduction velocity and improved both electrophysiological and Ca(2+) handling properties compared to nonstimulated controls. These changes were in agreement with cardiomyocyte maturation and were dependent on the stimulation rate.
© 2013 Nature America, Inc. All rights reserved.
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1
Directed differentiation protocols enable derivation of
cardiomyocytes from human pluripotent stem cells (hPSCs) and
permit engineering of human myocardium in vitro. However,
hPSC-derived cardiomyocytes are reflective of very early
human development, limiting their utility in the generation
of in vitro models of mature myocardium. Here we describe
a platform that combines three-dimensional cell cultivation
with electrical stimulation to mature hPSC-derived cardiac
tissues. We used quantitative structural, molecular and
electrophysiological analyses to explain the responses of
immature human myocardium to electrical stimulation and
pacing. We demonstrated that the engineered platform allows
for the generation of three-dimensional, aligned cardiac
tissues (biowires) with frequent striations. Biowires submitted
to electrical stimulation had markedly increased myofibril
ultrastructural organization, elevated conduction velocity
and improved both electrophysiological and Ca
2+
handling
properties compared to nonstimulated controls. These changes
were in agreement with cardiomyocyte maturation and were
dependent on the stimulation rate.
As adult human cardiomyocytes are essentially postmitotic, the
ability to differentiate cardiomyocytes from human embryonic
stem cells (hESCs) and human induced pluripotent stem cells
(hiPSCs)
1–4
represents an exceptional opportunity to create
in vitro models of healthy and diseased human cardiac tissues that
can also be patient-specific
5
and useful in screening new thera-
peutic agents for efficacy. However, differentiated cells exhibit
a low degree of maturation
6
and are appreciably different from
adult cardiomyocytes.
hESC-derived cardiomyocytes exhibit immature sarcomere
structure characterized by the absence of H zones, I bands
and M lines (day 40 embryoid bodies
7
), high proliferation rates
Biowire: a platform for maturation of human
pluripotent stem cell–derived cardiomyocytes
Sara S Nunes
1,2
, Jason W Miklas
1,13
, Jie Liu
3,13
, Roozbeh Aschar-Sobbi
3
, Yun Xiao
1
, Boyang Zhang
1
,
Jiahua Jiang
4
, Stéphane Massé
5
, Mark Gagliardi
6
, Anne Hsieh
1
, Nimalan Thavandiran
1
, Michael A Laflamme
7
,
Kumaraswamy Nanthakumar
5
, Gil J Gross
4,8–10
, Peter H Backx
3,10,11
, Gordon Keller
6
& Milica Radisic
1,10,12
(~17% proliferating cells for day 37 embryoid bodies
8
(EBd37)
and ~10% for day 21–35 embryoid bodies
7
), immature action
potentials
9
and Ca
2+
handling properties
9–13
with contraction
shown to be, in many cases, dependent on trans-sarcolemmal
Ca
2+
influx and not on sarcoplasmic reticulum Ca
2+
release
10
.
hESC-based engineered cardiac tissues also exhibit characteris-
tics of immature cells, including immature sarcomere structure
14
,
high proliferation rates (15–45% proliferating cells in ref. 14 and
10–30% proliferating cells in ref. 15) and expression of the fetal
gene program
16–18
. This is an important caveat when using these
cells as models of adult human tissue
6
.
During embryonic development, cardiac cells are exposed to
environmental cues such as extracellular matrix, soluble factors,
mechanical signals and electrical fields that may determine the
emergence of spatial patterns and aid in tissue morphogenesis
19,20
.
Exogenously applied electrical stimulation has also been shown
to influence cell behavior
21–23
.
We created a platform that combines architectural and electrical
cues to generate a microenvironment conducive to maturation of
three-dimensional (3D) hESC-derived and hiPSC-derived cardiac
tissues, termed ‘biowires’. We seeded cells in collagen gel around
a template suture in a microfabricated well and subjected them
to electrical field stimulation with a progressive increase in fre-
quency. Consistent with maturation, stimulated biowires exhib-
ited cardiomyocytes with a remarkable degree of ultrastructural
organization, improved conduction velocity and enhanced Ca
2+
handling and electrophysiological properties.
RESULTS
Engineering of human cardiac biowires
We generated 3D, self-assembled cardiac biowires by seeding
cardiomyocytes, derived from hPSCs using a directed differ-
entiation protocol in embryoid bodies
2
, and supporting cells
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
Department of Physiology and Medicine, University of Toronto, Toronto, Ontario, Canada.
4
Cardiology Division, Hospital for
Sick Children, Toronto, Ontario, Canada.
5
The Toby Hull Cardiac Fibrillation Management Laboratory, Toronto General Hospital, Toronto, Ontario, Canada.
6
McEwen
Centre for Regenerative Medicine, University Health Network, Toronto, Ontario, Canada.
7
Department of Pathology, University of Washington, Seattle, Washington,
USA.
8
Physiology and Experimental Medicine Program, Hospital for Sick Children Research Institute, Toronto, Ontario, Canada.
9
Department of Pediatrics, University of
Toronto, Toronto, Ontario, Canada.
10
The Heart and Stroke/Richard Lewar Centre of Excellence, Toronto, Ontario, Canada.
11
Division of Cardiology, University Health
Network, Toronto, Ontario, Canada.
12
Department of Chemical Engineering and Applied Chemistry, University of Toronto, Toronto, Ontario, Canada.
13
These authors
contributed equally to this work. Correspondence should be addressed to M.R. (m.radisic@utoronto.ca).
Received 6 FebRuaRy; accepted 17 May; published online 23 june 2013; doi:10.1038/nMeth.2524
© 2013 Nature America, Inc. All rights reserved.
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(fibroblasts, endothelial cells and smooth muscle cells) into a
template poly(dimethylsiloxane) (PDMS) channel, around a
sterile surgical suture in type I collagen gels (Fig. 1 and
Supplementary Fig. 1a). The biowire suture remained anchored
to the device platform during matrix remodeling. Seeded cells
remodeled and contracted the collagen gel matrix during the
first week after seeding (Fig. 1a and Supplementary Fig. 1a)
with ~40% gel compaction (Fig. 1b; final width, ~600 µm).
This allowed us to remove the biowire from the PDMS template
(Supplementary Figs. 1 and 2).
Histological analysis revealed cell alignment along the axis of
the suture (Fig. 1c and Supplementary Fig. 2b). Biowires beat
synchronously and spontaneously between 2 d and 3 d after
seeding and kept beating after gel compaction, demonstrat-
ing that the setup enabled electromechanical cell coupling
(Supplementary Video 1). Biowires could be electrically paced
and responded to physiological agonists such as epinephrine
(β-adrenergic stimulation) by increasing frequency of spontane-
ous beating (Fig. 1d).
After preculture for 1 week, we either submitted the biowires to
electrical field stimulation or cultured them without stimulation
(nonstimulated controls) for 7 d. To assess whether effects were
dependent on stimulation rate, we used two different protocols:
(i) low-frequency ramp-up regimen, where stimulation started
at 1 Hz, increased to 3 Hz (1 Hz, 1.83 Hz, 2.66 Hz and 3 Hz on
days 1–4, respectively) and maintained at 3 Hz for the remainder
of the week (Supplementary Fig. 1b; referred to as low-frequency
regimen or 3-Hz regimen) or (ii) high-frequency ramp-up regi-
men, where stimulation started at 1 Hz and increased to 6 Hz
throughout the week (1 Hz, 1.83 Hz, 2.66 Hz, 3.49 Hz, 4.82 Hz,
5.15 Hz and 6 Hz; Supplementary Fig. 1c; referred to as high-
frequency regimen or 6-Hz regimen).
Physiological hypertrophy in stimulated biowires
After 2 weeks in culture, cells throughout the biowires strongly
expressed cardiac contractile proteins sarcomeric α-actinin,
actin and cardiac troponin T, as evidenced by immunostaining
(Fig. 2a and Supplementary Figs. 2 and 3). Sarcomeric banding
Day 0
a
200 µm
Day 1 Day 2 Day 4 Day 5 Day 7
d
Spontaneous activity
1 mm
Electrical stimulation
1 s
+ Epinephrine
b c
1,200
1,000
800
600
Width (µm)
400
200
0
0 1 2
Time in culture (d)
5 6 7
H&E MT
100 µm
Figure 1
|
Generation of human cardiac biowires. (a) Brightfield images
of Hes2 hESC-derived cardiomyocytes on indicated days of preculture in
biowire template. (b) Quantification of gel compaction on the indicated
days of culture (average ± s.d., n = 3 (day 0), n = 4 (days 1–7)).
(c) Hematoxylin and eosin (H&E) and Masson’s trichrome (MT) staining
of biowire sections (double-headed arrows represent suture axis).
(d) Representative picture (left) of a biowire being imaged with
potentiometric fluorophore (DI-4-ANEPPS), which shows spontaneous
electrical activity, with impulse propagation recording (left trace
recording), response to electrical stimulation (middle trace recording,
stimulation frequency is depicted in red trace below) and increase in
frequency of spontaneous response under pharmacological stimulation
(epinephrine, right trace recording).
Sarcomere Desmosomes
m
c
Control3 Hz6 Hz
m
I bands (per Z disc)
Control 3 Hz 6 Hz
d
H zones (per sarcomere)
*
*
2.5
2.0
1.5
1.0
0.5
0
P = 0.01
P = 0.003
0.8
0.7
0.6
0.5
Zero
*
0.4
0.3
0.2
0.1
0
Control 3 Hz 6 Hz
P = 0.005
*
8
7
6
5
4
3
2
1
0
Desmosomes (number
per nm membrane × 10
4
)
Control
3 Hz
6 Hz
P = 0.0003
α-Actinin, actin
α-actinin, DAPI
Cardiac troponin T
Control
a b
EBd34 Control
3 Hz 6 Hz
3 Hz6 Hz
100
80
60
Cell shape (%)
40
20
0
Control 3 Hz 6 Hz EBd34
P = 0.01
P = 0.03
*
*
Round
Rod-like
Figure 2
|
Cultured biowires in combination with electrical stimulation
promoted physiological cell hypertrophy and improved cardiomyocyte
phenotype. (a) Representative confocal images of nonstimulated (control)
and electrically stimulated biowires (3-Hz and 6-Hz regimens) showing
cardiomyocyte alignment and frequent Z disks (double-headed arrows
represent suture axis). (b) Analysis of cardiomyocyte cell shape in
indicated conditions (average ± s.d., n = 3 per group). EBd34 versus cells
subjected to 3-Hz regimen, P = 0.01 for both rod-like and round; EBd34
versus 6 Hz, P = 0.03 for both round and rod-like. DAPI, 4,6-diamidino-2-
phenylindole. (c) Representative ultrastructural images of nonstimulated
(control) and electrically stimulated biowires showing sarcomere structure
(sarcomere, white bar; Z disks, black arrows; H zones, white arrows;
m, mitochondria) and presence of desmosomes (desmosomes, white
arrows). Scale bars, 20 µm (a), 50 µm (b) and 1 µm (c). (d) Morphometric
analysis (average ± s.d.; n = 4 per group) showing ratio of H zones to
sarcomeres (control versus 6 Hz, P = 0.005) ratio of I bands to Z disks
(control versus 3 Hz, P = 0.01; control versus 6 Hz, P = 0.003) and number
of desmosomes per membrane length (control versus 6 Hz, P = 0.0003).
*, significant difference between group and control. In normal adult cells,
the ratio of H zones to sarcomeres is 1 and of I bands to Z disks is 2.
Data in ad were obtained with Hes2 hESC-derived cardiomyocytes.
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of the contractile apparatus (Fig. 2a and Supplementary Figs. 2c,
3a and 4) and myofibrillar alignment along the suture axis was
qualitatively similar to the structure in the adult heart
22
. Biowires
kept in culture for 3 weeks and 4 weeks maintained cell alignment
and their contractile apparatus structure, as evidenced by confocal
and transmission electron microscopy (Supplementary Fig. 5).
Early in cardiac development, cardiomyocytes are round cells
and differentiate into rod-like cells after birth
24
. Adult human
cardiomyocytes have a structurally rigid architecture, retain-
ing a rod-like shape
25
immediately after dissociation, whereas
hESC-derived cardiomyocytes remain round. We dissociated
age-matched, EBd34 and biowires, and seeded the cells into
Matrigel-coated plates (Supplementary Fig. 1d). Although ~80%
of cardiomyocytes from EBd34 exhibited a round phenotype, this
percentage was significantly lower (~50% less) in electrically
stimulated samples (Fig. 2b). The percentage of rod-like car-
diomyocytes was significantly higher (about fourfold) in electri-
cally stimulated biowires (Fig. 2b and Supplementary Fig. 6) as
compared to EBd34.
During development, cardiomyocytes undergo physiological
hypertrophy characterized by an increase in cell size followed
by changes in sarcomere structure and downregulation of fetal
genes
26
. We observed a significant increase in cardiomyocyte
size (area of plated cells) in biowire conditions compared to
cardiomyocytes from age-matched embryoid bodies (EBd34)
(Supplementary Table 1; EBd34 versus control, P = 0.034; EBd34
versus 3-Hz regimen, P = 0.003; EBd34 versus 6-Hz regimen,
P = 0.01). Atrial natriuretic peptide (NPPA), brain natriuretic
peptide (NPPB) and α-myosin heavy chain (MYH6) are proteins
highly expressed in fetal cardiomyocytes and upregulated during
pathological hypertrophy in diseased adult ventricular cardiomyo-
cytes. Downregulation of the fetal cardiac gene program (NPPA,
NPPB and MYH6) in hESC-derived cardiomyocyte biowires
(Supplementary Fig. 7), compared to age-matched embryoid
bodies, in concert with cell-size increase, suggested physiologi-
cal hypertrophy and a more mature phenotype. Although we
observed downregulation of mRNA encoding structural pro-
teins in biowires compared to embryoid bodies, we observed
no changes in the amounts of these protein (Supplementary
Results). Potassium inwardly rectifying channel gene (KCNJ2),
that has important roles in cell excitability and K
+
homeosta-
sis
27
, was upregulated in cells from biowires compared to EBd34
(Supplementary Fig. 7).
hESC-derived cardiomyocytes cultured in biowires also
exhibted lower proliferation rates than those of embryoid bodies
(Supplementary Fig. 8; EBd20 versus EBd34, P = 0.002; EBd34
versus control, P = 0.019; EBd34 versus 3-Hz regimen, P = 0.016;
EBd34 versus 6-Hz regimen, P = 0.015) and the percentage of
cardiomyocytes in each condition remained unchanged after cul-
ture for 2 weeks (48.2% ± 10.7% average ± s.d.; Supplementary
Fig. 9). After cultivation, cell composition in biowires was compa-
rable to that in EBd20, specifically CD31
+
(2.4% ± 1.5%, endothe-
lial cells
28
), CD90
+
(34.4% ± 23%, fibroblasts
28
), calponin
+
(35 ±
22%, smooth muscle cells) or vimentin
+
(80% ± 22%, nonmyo-
cytes) cells. This suggests that the observed improvements were
not related to the induction of a particular cell type in biowires.
Maturation of contractile apparatus
Cells in nonstimulated biowires exhibited well-defined Z discs
and myofibrils (Fig. 2c and Supplementary Figs. 3c and 4) but
no signs of alignment of Z discs. In contrast, biowires stimu-
lated under the high-frequency regimen showed signs of mat-
uration, such as organized sarcomeric banding with frequent
myofibrils that converged and displayed aligned Z discs (6-Hz
regimen; Fig. 2c and Supplementary Figs. 3c and 4), many mito-
chondria (6-Hz regimen; Fig. 2c and Supplementary Fig. 3c
and 4) and desmosomes, a molecular complex of cell-cell adhesion
proteins (Fig. 2c). In the 6-Hz regimen condition, mitochondria
were positioned closer to the contractile apparatus than in con-
trol or 3-Hz regimen conditions (Fig. 2c and Supplementary
Figs. 3c and 4b).
Electrically stimulated samples exhibited a sarcomeric organiza-
tion more compatible with mature cells than with nonstimulated
controls as shown by a significantly higher presence of H zones
per sarcomere (Fig. 2d, control versus 6-Hz regimen, P = 0.005;
Supplementary Fig. 3d, control versus 6-Hz regimen, P = 0.001)
and I bands per Z disc (Fig. 2d, control versus 3-Hz regimen,
P = 0.01; control versus 6-Hz regimen, P = 0.003; Supplementary
Fig. 3d, control versus 6-Hz regimen, P = 0.0004). Biowires stim-
ulated using the 6-Hz regimen also had a significantly higher
number of desmosomes per membrane length than did both
nonstimulated controls and biowires subjected to the 3-Hz regi-
men (Fig. 2d, P = 0.0003). In hiPSC-derived cardiomyocyte bio-
wires, we frequently saw areas with nascent intercalated discs
(Supplementary Fig. 3c and 4b). However, the lack of M lines
and T tubules, consistent with previous reports
29,30
, indicated an
absence of terminal differentiation.
Functional assessment of engineered biowires
Electrical stimulation using the 6-Hz regimen improved biowires
electrical properties, leading to a significant decrease in the exci-
tation threshold (Fig. 3a; control versus 6-Hz regimen, P = 0.03)
and an increase in the maximum capture rate (MCR) (Fig. 3b,
d
Control 3 Hz 6 Hz
200
0
Local activation
time (ms)
1 mm
4.0
a
3.5
3.0
2.5
2.0
1.5
1.0
0.5
Excitation threshold
(V cm
–1
)
0
Control
3 Hz
6 Hz
P = 0.03
*
b
6
5
4
3
2
1
0
Maximum capture rate
(Hz)
P = 0.022
*
Control
3 Hz
6 Hz
c
25
20
15
10
5
0
Conduction velocity
(cm s
–1
)
P = 0.014
P = 0.011
*
*
Control
3 Hz
6 Hz
Figure 3
|
Functional assessment of engineered biowires. (ac) Excitation
threshold (a; control (n = 4) versus cells subjected to 6-Hz regimen (n = 3),
P = 0.03; measured by field stimulation and videomicroscopy; 3 Hz,
n = 3), maximum capture rate (b; control versus 6 Hz (n = 4 per group),
P = 0.022; measured by point stimulation and optical mapping; 3 Hz,
n = 3) and electrical impulse propagation rates (c; control (n = 13) versus
3 Hz (n = 10), P = 0.014; control versus 6 Hz (n = 5), P = 0.011; measured
by point stimulation and optical mapping) after electrical stimulation
(average ± s.d.). (d) Representative conduction velocity activation maps
in biowires. *, significant difference between group and control. Data in
ad were obtained with hESC-derived cardiomyocytes from Hes2 cell line.
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control versus 6-Hz regimen, P = 0.022; Supplementary Figs. 3
and 4) as analyzed by point stimulation at the end of the cultiva-
tion in conjunction with optical mapping of impulse propaga-
tion (Supplementary Fig. 10a and Supplementary Videos 25).
Optical mapping demonstrated higher MCR with field stimula-
tion (5.2 Hz) than with point stimulation (4 Hz) (Supplementary
Fig. 10b; we observed an MCR of 5.2 Hz with field stimulation
and intermittent capture at 6 Hz, Supplementary Videos 69).
During field stimulation, all cells received the stimulus at the same
time, and response was not limited by each cell’s propagation limi-
tations. Conduction velocity, assessed upon point stimulation at
the end of cultivation, was ~40% and ~50% higher in the samples
electrically stimulated during culture (3 Hz and 6 Hz, respec-
tively), than nonstimulated controls (Fig. 3c,d, control versus
3 Hz, P = 0.014; control versus 6 Hz, P = 0.011). Improvements
in electrical properties (excitation threshold, MCR and conduc-
tion velocity) were more pronounced with the high-frequency
regimen compared to the low-frequency one. Improvement in
conduction velocity directly correlated with the average number
of desmosomes (Supplementary Fig. 11, R
2
= 0.8526).
Stimulation improves Ca
2+
-handling properties
Either all
10
or the majority
12
of hESC-derived cardiomyocytes rely
on Ca
2+
influx from sarcolemma rather than on Ca
2+
release from
sarcoplasmic reticulum for contraction, which differs markedly
from the case in the adult myocardium. We tested the effect of caf-
feine, which opens sarcoplasmic reticulum ryanodine channels,
on cytosolic Ca
2+
transients in single cells isolated from biowires
(Supplementary Fig. 1d). In accordance with previous work
10
,
none of the hESC-derived cardiomyocytes in nonstimulated
controls were responsive to caffeine (Fig. 4a), whereas electri-
cally stimulated cells using both 3-Hz and 6-Hz regimen condi-
tions responded to caffeine by inducing an increase in cytosolic
Ca
2+
(Fig. 4b,c). Quantification of Ca
2+
transient amplitudes
showed that electrically stimulated cells exhibited significantly
higher amplitude intensity in response to caffeine than nonstim-
ulated controls, in a stimulation frequency–dependent manner
(Fig. 4d,e). Blockade of L-type Ca
2+
channels in cells from bio-
wires subjected to the 6-Hz regimen with either verapamil or
nifedipine (Fig. 4f,g) led, as expected in mature cells, to cessa-
tion of Ca
2+
transients. Addition of caffeine after blockade of
L-type Ca
2+
channels led to Ca
2+
release into the cytosol (Fig. 4f,g).
Blockade of the ion-transport activity of sarcoplasmic reticulum
Ca
2+
ATPase (SERCA) by addition of thapsigargin (Fig. 4h) led to
the cessation of calcium transients with time because of the deple-
tion of Ca
2+
from the sarcoplasmic reticulum. Cardiomyocytes
from the 6-Hz regimen condition also demonstrated a faster ris-
ing slope and time to peak, parameters that represent the kinetics
of Ca
2+
release into the cytosol and faster τ decay and time to base,
parameters that represent the kinetics of clearance of Ca
2+
from
the cytosol (Supplementary Table 2). Taken together, these data
indicated that cardiac biowires stimulated using the 6-Hz regi-
men during culture exhibited Ca
2+
-handling properties compat-
ible with a functional sarcoplasmic reticulum.
Stimulation alters electrophysiological properties
To assess maturity, we measured action potentials, human ERG
(hERG) currents and inward rectifier currents (I
K1
)
11
in cardio-
myocytes derived from biowires and embryoid bodies (Fig. 5).
hERG currents were larger (P = 0.0434) in biowires subjected to
the 6-Hz regimen (0.81 pA pF
−1
± 0.09 pA pF
−1
) than nonstimu-
lated controls (0.52 pA pF
−1
± 0.10 pA pF
−1
) (Fig. 5a) without dif-
ferences in their biophysical properties (Supplementary Fig. 12).
Cardiomyocytes from both biowire groups had higher hERG levels
compared to those from day-20 or day-44 embryoid bodies (Fig. 5a).
Similarly, I
K1
densities were greater (P = 0.0406) in biowires
subjected to the 6-Hz regimen (1.53 pA pF
−1
± 0.25 pA pF
−1
)
than in controls (0.94 pA pF
−1
± 0.14 pA pF
−1
), and I
K1
levels
in both biowire groups were greater (P = 0.0005) than those
recorded in embryoid body–derived cardiomyocytes (Fig. 5b).
Caffeine, 5 mM
f
Verapamil, 1 mM
20 s
1.0 F/F0
[Ca
2+
]i transients
Caffeine, 5 mM
g
Nifedipine, 10 µM
20 s
1.0 F/F0
[Ca
2+
]i transients
Caffeine, 5 mM
h
Thapsigargin, 2 µM
20 s
1.0 F/F0
[Ca
2+
]i transien
Caffeine, 5 mM
e
a
Caffeine, 5 mM
20 s
1.0 F/F0
[Ca
2+
]i transients
b
Caffeine, 5 mM
20 s
1.0 F/F0
[Ca
2+
]i transients
c
Caffeine, 5 mM
20 s
1.0 F/F0
[Ca
2+
]i transients
d
Control
3 Hz
6 Hz
*
*
#
60
40
20
0
–20
–40
Caffeine-induced change
of peak intensity (F, %)
Figure 4
|
Electrical stimulation promoted
improvement in Ca
2+
handling properties.
(ac) Representative traces of Ca
2+
release
in response to caffeine in nonstimulated
control cells (a), cells subjected to the
3-Hz regimen (b) and 6-Hz regimen (c).
(d) Caffeine-induced change of peak
fluorescence intensity among experimental
groups (mean ± s.e.m. after normalizing the
peak fluorescence intensity before administration
of caffeine; control versus 3 Hz, P = 1.1 × 10
−6
; control versus 6 Hz, P = 2.1 × 10
−7
; 3 Hz versus 6 Hz, P = 0.003; n = 10 (control), n = 6 (3 Hz) and
n = 9 (6 Hz)). (e) Representative fluorescence recording of Ca
2+
transients before and after administration of caffeine at 5 mM (arrow) in cells subjected
to the 6-Hz regimen. (fh) Inhibition of L-type Ca
2+
channels with verapamil (f) or nifedipine (g) and blockade of SERCA channels with thapsigargin (h)
in cells subjected to the 6-Hz regimen before addition of caffeine. *, significant difference between group and control.
#
, significant difference between
3-Hz regimen group and 6-Hz regimen group. Data in ah were obtained with hESC-derived cardiomyocytes obtained from Hes2 cell line and represent
measurements performed in single cell cardiomyocytes after dissociation from biowires. F, fluorescence intensity; F0, fluorescence intensity at baseline;
F/F0, F normalized by F0.
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Cell capacitance, a measure of cell size, was larger (P = 0.0052)
in biowires subjected to the 6-Hz regimen (19.59 pF ± 1.41 pF)
compared to control biowires (14.23 pF ± 0.90 pF) and smaller
(P = 0.0041) in embryoid body–derived cardiomyocytes (Fig. 5c).
Resting membrane potentials (V
rest
) of the cardiomyocytes from
biowires were more negative than in embryoid body–derived car-
diomyocytes (P < 0.0001; Fig. 5d). After correcting for the liquid
junction potential, which was ~16 mV, the values of V
rest
recorded
in biowire cardiomyocytes with the patch-clamp method were
well below the equilibrium potential for Nernst potential for K
+
(E
K
= −96 mV), suggesting that hyperpolarizing currents, possibly
those generated by the Na
+
pump
31,32
, strongly influenced V
rest
.
Consistently, we found that the cardiomyocytes from biowires
had a very low resting membrane conductance, which correlated
with V
rest
(R = 0.5584, P < 0.0001), and I
K1
values had negative
correlations with V
rest
(R = 0.2267, P = 0.0216; Supplementary
Fig. 13). Maximum depolarization rates (Fig. 5e) and peak volt-
ages of the action potentials (Fig. 5f) did not differ between the
two biowire groups. However, both properties were improved in
these biowire groups compared to embryoid bodies (P = 0.5248
and P = 0.0488, respectively). Action potential durations were
longer (P = 0.0021) with greater variation in embryoid body–
derived cardiomyocytes than biowire-derived cardiomyocytes
(Fig. 5g and Supplementary Fig. 14), suggesting less electrophys-
iological diversity and more maturation in biowires. Automaticity
(spontaneous beating activity) was greater (P = 0.0414) in embry-
oid body–derived cardiomyocytes compared to control biowires
(Fig. 5h), which was comparable to that in biowires subjected
to the 6-Hz regimen. Taken together, these results support the
conclusion that biowires and electrical stimulation using the 6-Hz
regimen promoted electrophysiological maturation.
DISCUSSION
Normal human fetal heart rate varies but is maintained at ~3 Hz
for most of the time
33
whereas adult resting heart rate is ~1 Hz
33
.
The rate change is associated with changes in expression of con-
tractile proteins and suggests a possible dependence of cardiac
maturation on stimulation rate. The fact that the progressive
increase from 1 Hz to 6 Hz was the best stimulation condition
tested in biowires was a surprise to us as 3 Hz is the average fetal
heart rate
33
. This could be a compensatory mechanism for the
lack of other important cells types and cell-cell developmental
guidance in the in vitro setting. As we increased frequency of field
stimulation over 7 d in culture, the group subjected to the 6-Hz
regimen might only lose capture (exceed the rate of 5.2 Hz) at
the very last day of stimulation. Therefore, it may be the stimula-
tion at the highest possible rate, and not the rate itself, that is the
governing cue for cardiomyocyte maturation in vitro.
Mechanical stimulation has been reported to lead to a robust
induction of structural proteins such as myosin heavy chain and
induce proliferation of hPSC-derived cardiomyocytes
14,34
, sug-
gesting that electrical stimulation of the biowire at 6 Hz did not
simply provide a better mechanical stimulation environment.
Previously, mechanical stimulation did not lead to electrophysio-
logical maturation
34
. The use of electrical stimulation in conjunc-
tion with stretch as a mimic of cardiac load
14
, concurrently or
sequentially, might be required to induce terminal differentiation
in hPSC-derived cardiomyocytes and upregulate the expression of
myofilament proteins. Other strategies might include cultivation
in the presence of T3 thyroid hormone
35
, insulin-like growth
factor-1 (ref. 36), addition of laminin or native decellularized
heart extracellular matrix into the hydrogel mixture
37
and
cultivation on stiffer substrates
38,39
.
It is well accepted that some human stem cell lines are more
cardiomyogenic than others
12,16
, and these differences could
also be related to the maturity of the produced cells. In previous
reports
10,11,40
, many and usually most cells were irresponsive to
caffeine at the end of differentiation. Therefore, differences in Ca
2+
-
handling properties could also be due to cell line variability. Here we
demonstrated that in a given cell line, culture in biowires and electri-
cal-field stimulation enhanced Ca
2+
-handling properties of cardio-
myocytes consistent with a functional sarcoplasmic reticulum.
Biowire cardiomyocytes were clearly more mature than
cardiomyocytes obtained from embryoid bodies cultivated for
20 d (EBd20) or for 40–44 d (EBd44), which exhibited a greater
propensity for automaticity, more depolarized membrane
f
Action potential
Peak voltage (mV)
6 Hz Control EBd44 EBd20
n = 17
n = 13 n = 13 NA
P = 0.0200
P = 0.0488
100
50
0
b
3
2
1
0
I
K1
at –100 mV
current density (pA/pF)
6 Hz Control EBd44 EBd20
n = 25 n = 14
n = 28 n = 15
P = 0.0406
P = 0.0471
a
1.0
P = 0.0434
6 Hz
hERG current density
(pA/pF)
Control EBd44 EBd20
n = 25 n = 27 n = 16
n = 56
P = 0.0387
P = 0.0355
0.5
0
d
Resting membrane
potential (mV)
6 Hz Control EBd44 EBd20
n = 36 n = 51
n = 16
n = 13
P < 0.0001
P < 0.0001
0
–50
–100
–150
c
Cell capacitance (pF)
6 Hz Control EBd44 EBd20
n = 62 n = 60n = 42 n = 16
P = 0.0052
P = 0.0048
30
20
10
0
g
APD90 (ms)
6 Hz Control EBd44 EBd20
n = 17 n = 13 n = 13 NA
P = 0.0021
3,000
2,000
200
100
0
e
Maximum depolarization
rate (mV ms
–1
)
6 Hz Control EBd44 EBd20
n = 17
n = 13 n = 13 NA
P = 0.0107
200
100
0
h
Automaticity No automaticity
Ratio (%)
6 Hz Control EBd44 EBd20
NA
n = 5 n = 5 n = 11
P = 0.0414
100
50
0
n = 12 n = 8 n = 2
Figure 5
|
Electrophysiological properties in single cardiomyocytes isolated from biowires or embryoid bodies and recorded with patch clamp. (a) hERG
tail current density. (b) I
K1
current density measured at −100 mV. (c) Cell capacitance. (d) Resting membrane potential. (e) Maximum depolarization
rate of action potential. (f) Action potential peak voltage. (g) Action potential duration measured at 90% repolarization (APD90). (h) Ratio of cells
displaying spontaneous beating (automaticity) or no spontaneous beating (no automaticity). Control, control biowire. Data in ah were obtained with
hESC-derived cardiomyocytes obtained from Hes2 cell line (average ± s.e.m.).
© 2013 Nature America, Inc. All rights reserved.
6
|
ADVANCE ONLINE PUBLICATION 
|
NATURE METHODS
ARTICLES
potentials, lower cell capacitance and less hERG currents and I
K1
.
Electrophysiological measurements of the EBd20 cardiomyocytes
represented the cell properties before their incorporation into
biowires, whereas EBd44 cardiomyocytes were cultured for peri-
ods slightly longer than the biowire culture time, allowing assess-
ment of the independent effect of culture time on maturation
11,41
.
We acknowledge that biowire maturation is clearly incomplete,
as evidenced by the relatively low membrane conductance.
Nevertheless, it is intriguing to speculate that the combination
of low membrane conductance with V
rest
below E
K
may represent
an ‘intermediate’ phenotype as cardiomyocytes undergo matura-
tion from the embryonic state.
Correlating the properties of hPSC-derived cardiomyocytes in
biowires with mouse or human development could help to gauge
maturation stage, but mouse and rat cardiomyocytes are physi-
ologically distinct, and age-defined healthy human heart samples
are scarce. Additionally, in vitro maturation might not be compat-
ible with embryo development.
The small size (radius of ~300 µm) of biowire upon gel com-
paction was selected to be close to the diffusional limitations
for oxygen supply to ensure that the biowires can be maintained
in culture without perfusion. Addition of vascular cells will be
imperative for improving survival and promoting integration
with the host tissue in future in vivo studies
14
. We generated a
unique platform that enables generation of human cardiac tissues
of graded levels of maturation that can be used to determine, in
future in vivo studies, the optimal maturation level that will result
in the highest ability of cells to survive and integrate in adult
hearts with the lowest side effects (such as arrhythmias).
METHODS
Methods and any associated references are available in the online
version of the paper.
Note: Supplementary information is available in the online version of the paper.
ACKNOWLEDGMENTS
We thank P. Lai, C. Laschinger, N. Dubois and B. Calvieri for technical assistance,
C.C. Chang and L. Fu for assistance with biowire setup figure preparation. 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 and MOP-62954), NSERC-CIHR Collaborative Health
Research Grant (CHRPJ 385981-10), NSERC Discovery Grant (RGPIN 326982-10),
and NSERC Discovery Accelerator Supplement (RGPAS 396125-10) and National
Institutes of Health grant 2R01 HL076485.
AUTHOR CONTRIBUTIONS
S.S.N. developed biowire concept, designed and performed experiments, analyzed
data and prepared the manuscript. J.W.M. performed experiments and analyzed
data. J.L., R.A.-S. and P.H.B. performed patch clamping and microelectrode
recordings. Y.X. designed and validated initial device. B.Z. designed and fabricated
masters for device fabrication. J.J. and G.J.G. performed calcium transient
measurement and analysis. S.M. and K.N. performed optical mapping measurements
and analysis. M.G. and G.K. differentiated hESC-derived cardiomyocytes.
A.H. designed primers. N.T. developed initial collagen gel mixture. M.A.L. provided
training on hiPSC differentiation and cells. P.H.B. contributed to writing of the
manuscript. M.R. envisioned the biowire concept and electrical stimulation
protocol, supervised the work and wrote the manuscript.
COMPETING FINANCIAL INTERESTS
The authors declare competing financial interests: details are available in the
online version of the paper.
Reprints and permissions information is available online at http://www.nature.
com/reprints/index.html.
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© 2013 Nature America, Inc. All rights reserved.
doi:10.1038/nmeth.2524
NATURE METHODS
ONLINE METHODS
Human pluripotent stem cell culture and differentiation.
Cardiomyocytes derived from two different hESC lines (Hes2 and
Hes3) and two different hiPSC lines (CDI-MRB and HR-I-2Cr-2R)
were used. Both hESC lines and hiPSC line HR-I-2Cr-2R were
maintained as described
2,4
. Embryoid bodies (EBs) were differen-
tiated to the cardiovascular lineage as previously described
2,4
. In
brief, EBs were generated by culture in StemPro-34 (Invitrogen)
medium containing BMP4 (1 ng/ml). On day 1, EBs were col-
lected and suspended in induction medium (StemPro-34 contain-
ing basic fibroblast growth factor (bFGF; 2.5 ng/ml), activin A
(6 ng/ml) and BMP4 (10 ng/ml)). On day 4, the EBs were collected
from the induction medium and recultured in StemPro-34
supplemented with vascular endothelial growth factor (VEGF;
10 ng/ml) and DKK1 (150 ng/ml). On day 8, the medium was
changed again and the EBs were cultured in StemPro-34 contain-
ing VEGF (20 ng/ml) and bFGF (10 ng/ml) for the duration of
the experiment. Cultures were maintained in hypoxic environ-
ment (5% CO
2
and 5% O
2
) for the first 12 d and then transferred
into 5% CO
2
for the remainder of the culture period. EBs were
dissociated for seeding in biowires on day 20 (EBd20), day 34
(EBd34) and day 40–44 (EBd44) for specific cellular and electro-
physiological analyses. CDI-MRB hiPSC-derived cardiomyocytes
were purchased from Cellular Dynamics International (CMC-
100-110-001) and used for biowire production immediately
after thawing.
Device design and manufacture. The device was fabricated
using soft lithography technique. A two-layer SU-8 (Microchem
Corp.) master was used to mold PDMS. Briefly, device features
were printed on two film masks (CADART) corresponding to the
two-layer design. SU-8 2050 was spun onto 4-inch silicon wafer,
baked and exposed to UV light under the first-layer mask to cre-
ate the first layer including the suture channel and the chamber
with thickness of 185 µm. The second layer including only the
chamber with thickness of 115 µm was spun on top. After addi-
tional baking, the second-layer mask was aligned to the features
on the first layer and then exposed to UV light. Finally, the wafer
was developed using propylene glycol monomethyl ether acetate
(Doe & Ingalls Inc.). PDMS was then cast onto the SU-8 master
and baked for 2 h at 70 °C. PDMS templates were used to hold a
piece of surgical suture centrally in the channel (Supplementary
Fig. 1a) to which the cardiac cell suspension gel was added.
Biowire generation. EBd20 generated as described above were
incubated in collagenase type I (1 mg/ml; Sigma) and DNAse
(1 mg/ml, CalBiochem) in Hanks balanced salt solution (NaCl,
136 mM; NaHCO
3
, 4.16 mM; Na
3
PO
4
, 0.34 mM; KCl, 5.36 mM;
KH
2
PO
4
, 0.44 mM; dextrose, 5.55 mM and HEPES, 5 mM) for
2 h at 37 °C. EBs were centrifuged (107g, 5 min), incubated with
trypsin (0.25%, Gibco) for 5 min at 37 °C and pipetted gently
to dissociate the cells. After dissociation, cells were centrifuged
(167 x g, 5 min), counted and seeded at 0.5 × 10
6
cells/wire of
0.5 cm in length. This ratio was maintained for generation of longer
biowires. Cells were seeded in collagen type I gels (4 µl/0.5 cm wire
length; 2.1 mg/ml of rat tail collagen type I (BD Biosciences) in
24.9 mM glucose, 23.81 mM NaHCO
3
, 14.34 mM NaOH, 10 mM
HEPES, in 1× M199 medium plus 10% of growth factor reduced
Matrigel (BD Biosciences)) by pipetting the cell suspension into
the main channel of the PDMS template (Supplementary Fig. 1a).
CDI-MRB hiPSC-derived cardiomyocytes were thawed,
counted and seeded in same concentration as hESC-derived
cardiomyocytes. After seeding, cells were kept in culture for 7 d
to allow collagen matrix remodeling and assembly around the
suture (Fig. 1a,b).
Electrical stimulation setup and conditions. After preculture for
7 d, biowires were transferred to stimulation chambers fitted with
two 1/4-inch-diameter carbon rods (Ladd Research Industries)
placed 2 cm apart and connected to a cardiac stimulator (Grass
s88x) with platinum wires (Ladd Research Industries). Biowires
were placed perpendicular to the electrodes and were either
submitted to electrical stimulation (rectangular, biphasic, 1 ms,
3–4 V/cm) or cultured without electrical stimulation (nonstimu-
lated controls or control) for 7 d (Supplementary Fig. 1b,c). As
increased time in culture affects maturation
11,41
, age-matched EBs
(EBd34) were used as an additional control. For long-term stimu-
lation experiments, shown in Supplementary Fig. 5, the biowires
were precultured for 7 d as described above, followed by 7 d of
6-Hz protocol, at which point the frequency was decreased to
1 Hz (to mimic postnatal decrease in heart rate) and maintained
for an additional 14 d.
To verify that biowire-stimulated cardiomyocytes truly exhib-
ited maturation on a single-cell basis, assays were performed
in which single cells were used. With this goal, biowires were
digested with collagenase type I (1 mg/ml; Sigma) and DNAse
(1 mg/ml, CalBiochem) in Hanks balanced salt solution (NaCl,
136 mM; NaHCO
3
, 4.16 mM; Na
3
PO
4
, 0.34 mM; KCl, 5.36 mM;
KH
2
PO
4
, 0.44 mM; dextrose, 5.55 mM; and HEPES, 5 mM) for
4 h at 37 °C, centrifuged (107g, 5 min), incubated with trypsin
(0.25%, Gibco) for 5 min at 37 °C and pipetted gently to dissoci-
ate the cells as depicted in Supplementary Figure 1d. Isolated
single cells were seeded on Matrigel-coated or laminin-coated
glass coverslips as described below, and area, calcium transient
and patch-clamp measurements were performed.
Assessments. The progression of tissue assembly was assessed
at various levels after 2 weeks in culture (7 d of gel compaction
followed by 7 d of stimulation): functional (excitation threshold,
MCR, conduction velocity and Ca
2+
handling); ultrastructural
(sarcomere development, frequency (number/membrane length)
of desmosomes), cellular (cell size and shape, proliferation, dis-
tribution of cardiac proteins: actin, troponin T and α-actinin),
electrophysiological (hERG, I
K1
and I
Na
) and molecular (expres-
sion of cardiac genes and proteins).
Immunostaining and fluorescence microscopy. Immunostaining
was performed using the following antibodies: mouse anti
cardiac troponin T (1:100, Thermo Scientific; MS-295-P1), mouse
anti-α-actinin (1:200, Abcam, ab9465), anti-mouse–Alexa Fluor
488 (1:400, Invitrogen, A21202), anti-Ki67 (1:250, Millipore,
AB9260), anti-rabbit–TRITC (1:400, Invitrogen, 81-6114).
DAPI was used to counterstain nuclei. Phalloidin–Alexa Fluor
660 (1:1,000, Invitrogen, A22285) was used to detect actin fibers.
The stained cells were visualized using a fluorescence microscope
(Leica CTR6000) and images captured using the Leica Application
Suite software. For confocal microscopy, cells were visualized
using a fluorescence confocal microscope (Zeiss LSM-510).
© 2013 Nature America, Inc. All rights reserved.
doi:10.1038/nmeth.2524
NATURE METHODS
Transmission electron microscopy. Tissue was fixed with
4% paraformaldehyde, 1% glutaraldehyde in 0.1 M PBS for at
least 1 h and washed 3 times with PBS pH 7.2. Post-fixation was
done with 1% osmium tetraoxide in 0.1 M PBS, pH 7.2 for 1 h
and dehydrated using ethanol series from 25% to 100%. Tissue
was infiltrated using Epon resin and polymerized in plastic dishes
at 40 °C for 48 h. Tissue was stained with uranyl acetate and lead
citrate after sectioning. Imaging was performed at Hitachi H-7000
transmission electron microscope.
Optical mapping. Biowires were incubated with a voltage-
sensitive dye (Di-4-ANEPPS, 5 µM, Invitrogen) for 20 min at
37 °C in warm Tyrodes solution (118 mM NaCl, 4.7 mM KCl,
1.25 mM CaCl
2
, 0.6 mM MgSO
4
, 1.2 mM KH
2
PO
4
, 25 mM
NaHCO
3
and 6 mM glucose; oxygenated by bubbling carbogen
95% O
2
, 5% CO
2
for at least 20 min shortly before use). Dye
fluorescence was recorded on an MVX-10 Olympus fluorescence
microscope equipped with a high-speed complementary metal-
oxide semiconductor (CMOS) camera (Ultima-L, Scimedia)
42,43
.
The 1-cm sensor had 100 × 100 pixel resolution and the spa-
tial resolution was 50–100 µm/pixel. Imaging was performed at
200 frames/s. The fluorescence was excited using a mercury arc
source (X-Cite Exacte) with green filter (Olympus U-MWIG2
filter cube). The constructs were electrically point-stimulated
using a bipolar electrode made of two fine wires (AWG#32)
inserted in a stainless steel needle, which was mounted on a
micromanipulator (World Precision Instruments). For electri-
cal field stimulation, the chamber depicted in Supplementary
Figure 1 was used. The plate containing the biowires was placed
on a heated plate (MATS-U55S, Olympus) and temperature was
regulated at 38 °C. Data analysis was performed using BrainVision
software (Scimedia).
Intracellular recordings. Action potentials were recorded in bio-
wires with high-impedance glass microelectrodes (50–70 M,
filled with 3 M KCl) at 37 ± 0.5 °C. Biowires were superfused
with Krebs solution containing 118 mM NaCl, 4.2 mM KCl,
1.2 mM KH
2
PO
4
, 1.8 mM CaCl
2
, 1.2 mM MgSO
4
, 23 mM
NaHCO
3
, 20 mM glucose and 2 mM Na-pyruvate, equilibrated
with 95% O
2
and 5% CO
2
; final pH was 7.4. The microelectrodes
were connected to an Axopatch 200B amplifier (Axon Instrument)
current clamp. Signals were filtered at 1 kHz, sampled at 2 kHz and
analyzed with Clampfit 10 (Axon Instrument). Resting potential
was measured at I = 0 mode. For some experiments, biowires were
paced using field stimulation set at twice the excitation threshold.
Patch-clamp recordings. Single cells isolated from biowires
(Supplementary Fig. 1d) or EBs were seeded on laminin-coated
glass coverslips (laminin, Sigma-Aldrich, 10 µg/cm
2
) overnight
before patch-clamp experiments were performed. Whole-cell
patch-clamp recordings were made using an Axopatch 200B
amplifier at room temperature (23–25 °C). Data were analyzed
with Clampfit 8.0 (Axon Instrument). Amplifier was set at I = 0
when measuring resting potential of cells. Action potentials were
recorded by using the current-clamp mode method. Myocytes
were stimulated at 1 Hz, and the maximum rate of membrane
depolarization, the action potential peak and action potential
duration at 90% (APD90) of the 10th action potential were mea-
sured. The membrane potentials were not corrected for the liquid
junction potentials, which were estimated to be 15.9 mV (esti-
mated with Clampfit 8.0) for the solutions used. Na
+
current,
hERG current and I
k1
current were also recorded under voltage-
clamp conditions with 70–80% series resistance compensation.
Na
+
current was induced from holding potential of80 mV by
applying a series of test pulses ranging from −120 mV to +30 mV
for 500 ms with 10-mV increments followed by a test pulse to
−10 mV for 100 ms for measurement of steady-state inactiva-
tion. Although this protocol simultaneously activates overlapping
voltage-dependent Ca
2+
currents, these Ca
2+
currents were esti-
mated (using prepulse protocols) to be less than 3% of the evoked
Na
+
currents. hERG was assessed by measuring tail currents in
response to steps to −50 mV (for 500 ms) after depolarization to
voltage steps ranging from −45 mV to 60 mV with 15-mV incre-
ments for 2,000 ms. The peak amplitude of hERG tail current
was measured and compared. I
K1
was measured in two ways that
were found to be equivalent for our studies. For complete I-V
relationships, we assessed Ba
2+
-sensitive currents by subtracting
(trace-by-trace for voltage steps ranging from 120 to −10 mV in
10-mV increments from holding potential of −40 mV) the cur-
rents measured in the presence of 500 µM Ba
2+
from the current
measured in the absence of Ba
2+
. For the purposes of measuring
the I
K1
density, we subtracted the background current from that
measured in the absence of Ba
2+
at −100 mV.
Patch-clamp recordings were performed in bath solutions
containing 140 mM NaCl, 4 mM KCl, 1 mM MgCl
2
, 1.2 mM
CaCl
2
, 10 mM HEPES, 10 mM -glucose, at pH 7.35 adjusted
with NaOH. Pipette resistance was ~5.5–7.5 M when filled with
a solution containing 120 mM potassium aspartate, 20 mM KCl,
4 mM NaCl, 1 mM MgCl
2
, 5 mM MgATP, 10 mM HEPES and
10 mM EGTA, at pH 7.2 adjusted with KOH (calculated reversal
potential of K
+
was −95.6 mV after pH adjustment). Dofetilide
100 nM (ref. 44) and BaCl
2
500 µM
11
were used to block hERG
current and I
k1
, respectively.
Calcium transient measurements. Biowires were dissociated by
incubation with collagenase and trypsin as described above. The
dissociated cardiomyocytes were plated onto 25-mm microscope
glass coverslips coated with growth factor–free Matrigel (diluted
1:60 in RPMI medium) and cultured overnight. Cells were then
incubated with 5 µM of fluo-4 acetoxymethyl ester (fluo-4 AM)
in culture medium for 2 h at 37 °C. Subsequently, cardiomyocytes
were washed twice with dye-free medium and placed back into
the incubator for 30 min. A laser-scanning confocal microscope
(Zeiss LSM 510) was used to measure the fluorescence intensity
of fluo-4 AM. The coverslips containing the fluo-4 AM–loaded
cardiomyocytes were moved onto a special chamber and tightly
secured. Approximately 1.8–1.9 ml of culture medium was added
into the chamber, which was placed on a temperature-controlled
plate (37 °C) on the microscope. Fluo-4 AM was excited via an
argon laser (488 nm), and emitted fluorescence was collected
through a 505 nm emission filter. Changes in fluo-4 AM fluores-
cence intensity, which indicates transient fluctuation of cytosolic
calcium concentration, were recorded in frame and line-scan
model. The images and fluorescence data were acquired through
Zeiss software. The fluorescence data were analyzed with Origin
8.5 software. Fluorescence signals were normalized to baseline
fluorescence after loading fluo-4 AM. The rising phase of the
signals was fitted by linear model, and the decaying phase of the
© 2013 Nature America, Inc. All rights reserved.
doi:10.1038/nmeth.2524
NATURE METHODS
signals was fitted by ExpDecay with Offset model. Caffeine, vera-
pamil (Sigma) and fluo-4 AM (Invitrogen) were directly added
into the chamber that contained the cardiomyocytes during imag-
ing at concentrations indicated in the figures. Cells beating at
similar average beating frequency (9.4 ± 0.7 beats per minute
(bpm) for control, 9 ± 0.7 bpm for 3-Hz regimen and 10 ± 0.8 bpm
for 6-Hz regimen) were used for calcium transient measure-
ments to ensure that differences in beating rates would not affect
the measurements.
Quantitative RT-PCR. RT-PCR was performed as previously
described
28
. Total RNA was prepared with the High Pure RNA
Isolation Kit (Roche) and treated with RNase-free DNase (Roche).
RNA was reverse-transcribed into cDNA using random hexamers
and oligo(dT) with SuperScript 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 genes TATA box binding
protein (TBP) or glyceraldehyde 3-phosphate dehydrogenase
(GAPDH). The oligonucleotide sequences are summarized in
Supplementary Table 3.
Flow cytometry analysis. Cells were obtained from biowires or
EBs by dissociation with collagenase and trypsin as described
above and fixed with 4% paraformaldehyde for 10 min at room
temperature. For intracellular epitopes, cells were permeabilized
in PBS containing 5% FBS and 0.1% Triton X for 10 min on ice
before a blocking step of 5% FBS in PBS for 30 min. Cells were
incubated with the following antibodies in blocking buffer on
ice for 1 h: anti-CD31-PE (1:100) and anti-CD90-APC (1:500,
BD Biosciences, 553373 and 559879, respectively); anti–cardiac
troponin T (1:100, Thermo Scientific, MS-295-P1); anti-calponin
H1 (1:250, Abcam, ab46794); and anti-vimentin (1:100, Sigma-
Aldrich, V6630). Secondary antibodies used were anti-mouse–
Alexa Fluor 488 (1:400, Invitrogen, A21202) and anti-rabbit–Cy5
(1:500, Jackson ImmunoResearch, 111-175-144). Owing to the
intrinsic variability in the percentage of cardiomyocytes in each
assay, the percentage of cells positive for each marker (above the
secondary-antibody-only control) was normalized to the starting
cell population (EBd20) of each experiment to accurately evaluate
whether a change in cell population was occurring.
Western blotting. Biowires were solubilized in (2×) Novex Tris-
Glycine SDS sample buffer (Life technologies) and proteins were
separated by electrophoresis in Novex Tris-Glycine gels (Life
Technologies) and transferred to Biotrace NT (Nitrocellulose,
Pall Corp.). Membranes were probed with either anti-myosin
heavy chain (total, Abcam, ab15, 1:2,000), Phospholamban 1D11
(gift of A. Gramolini, University of Toronto, 1:5,000) or GAPDH
(Millipore, MAB374, 1:10,000) antibodies. Secondary antibod-
ies used were peroxidase-conjugated (DAKO, P0448 or P0447,
1:2000). Membranes were developed with ECL reagent Luminata
Classico Substrate (Millipore).
Statistical analysis. Statistical analysis was performed using
SigmaPlot 12.0. Differences between experimental groups
were analyzed by Student’s t-test except in Figure 2b and
Supplementary Table 1, where statistics was done using two-
way ANOVA. Normality test (Shapiro-Wilk) and pairwise mul-
tiple comparison procedures (Holm-Sidak method) were used
for two-way ANOVA tests. In Supplementary Figure 6, statistics
was done using one-way ANOVA on Ranks (pairwise multiple
comparison, Dunns method). In Figure 5 and Supplementary
Figures 1214, differences between experimental groups were
analyzed by Students t-test, chi-squared test, ANOVA (pairwise
multiple comparisons, Holm-Sidak method). P < 0.05 was con-
sidered significant for all statistical tests.
42. Nanthakumar, K. et al. Optical mapping of Langendorff-perfused human
hearts: establishing a model for the study of ventricular fibrillation in
humans. Am. J. Physiol. Heart Circ. Physiol. 293, H875–H880 (2007).
43. Witkowski, F.X., Clark, R.B., Larsen, T.S., Melnikov, A. & Giles, W.R.
Voltage-sensitive dye recordings of electrophysiological activation in a
Langendorff-perfused mouse heart. Can. J. Cardiol. 13, 1077–1082 (1997).
44. Snyders, D.J. & Chaudhary, A. High affinity open channel block by
dofetilide of HERG expressed in a human cell line. Mol. Pharmacol. 49,
949–955 (1996).

Supplementary resources

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