published: 09 March 2017
Frontiers in Cell and Developmental Biology | www.frontiersin.org 1March 2017 | Volume 5 | Article 19
Univercell Biosolutions, France
University of Cologne, Germany
Stanford University, USA
Joao Mario Martins Bigares,
Wales Heart Research Institute-School
of Medicine-Cardiff University, UK
Sara S. Nunes
This article was submitted to
Stem Cell Research,
a section of the journal
Frontiers in Cell and Developmental
Received: 01 December 2016
Accepted: 21 February 2017
Published: 09 March 2017
Sun X and Nunes SS (2017)
Bioengineering Approaches to Mature
Human Pluripotent Stem Cell-Derived
Front. Cell Dev. Biol. 5:19.
Bioengineering Approaches to
Mature Human Pluripotent Stem
Xuetao Sun 1and Sara S. Nunes 1, 2, 3*
1Toronto General Research Institute, University Health Network, Toronto, ON, Canada, 2Institute of Biomaterials and
Biomedical Engineering, University of Toronto, Toronto, ON, Canada, 3Heart & Stroke/Richard Lewar Centre of Excellence,
University of Toronto, Toronto, ON, Canada
Human pluripotent stem cell-derived cardiomyocytes (hPSC-CM) represent a potential
unlimited cell supply for cardiac tissue engineering and possibly regenerative medicine
applications. However, hPSC-CMs produced by current protocols are not representative
of native adult human cardiomyocytes as they display immature gene expression
proﬁle, structure and function. In order to improve hPSC-CM maturity and function,
various approaches have been developed, including genetic manipulations to induce
gene expression, delivery of biochemical factors, such as triiodothyronine and
alpha-adrenergic agonist phenylephrine, induction of cell alignment in 3D tissues,
mechanical stress as a mimic of cardiac load and electrical stimulation/pacing or a
combination of these. In this mini review, we discuss biomimetic strategies for the
maturation for hPSC-CMs with a particular focus on electromechanical conditioning
Keywords: cardiomyocytes, cardiac regeneration, stem cell, biomaterials, cell therapy, electrical stimulation,
Human embryonic stem cells (hESCs), ﬁrst isolated from inner cell mass of blastocysts, possess
the capacity to diﬀerentiate into cells of all three germ layers (Thomson et al., 1998). Similar
characteristics can also be found in human induced pluripotent stem cells (hiPSCs), which are
generated from terminally diﬀerentiated, adult cells by genetically reprogramming via expression
of a set of transcription factors (Takahashi et al., 2007; Yu et al., 2007). These cells circumvent the
ethical concerns associated with hESCs and allow a potential autologous approach without the need
for long-term immunosuppression. Cardiomyocytes can be diﬀerentiated from both hESCs and
hiPSCs using directed diﬀerentiation approaches, which are based on the stage-speciﬁc treatment
with cardiogenic-inducing signaling factors (Laﬂamme et al., 2007; Yang et al., 2008).
However, human pluripotent stem cell derived cardiomyocytes (hPSC-CMs) (including hESC-
CM and hiPSC-CM) display immature characteristics when compared to adult cardiomyocytes,
such as (Table 1):
1) Genetically, hPSC-CMs express much lower levels of cardiac contractile and cytoskeletal genes
(Cao et al., 2008; Xu et al., 2009). Early hPSC-CMs have high proliferation rates (Robertson
et al., 2013) while adult cardiomyocytes are considered non-proliferative (∼0.5% proliferation
per year) (Bergmann et al., 2009).
2) Morphologically, hPSC-CMs are small, disorganized, mononucleated, round/triangular in
shape; while adult human cardiomyocytes are large, highly organized, ∼25% binucleated
Sun and Nunes Bioengineering Pro-maturation Strategies for hPSC-CMs
TABLE 1 | Human pluripotent stem cell-derived cardiomyocytes (hPSC-CMs) vs. adult ventricular cardiomyocytes.
Criteria hPSC-CM Adult ventricular cardiomyocytes References
Structure Shape Round Rod Gerdes et al., 1992; Lundy et al., 2013
Cell surface area 10212–14418 µm2500–1294 µm2Li et al., 1996; Lundy et al., 2013; Ribeiro et al.,
Gene Expression MYH7 <MYH6 TNNI3 <TNNI1 MYH7 >MYH6 TNNI3 >TNNI1 Xu et al., 2009
Nuclei Mononuclear 25% binucleation Olivetti et al., 1996; Snir et al., 2003
Sarcomere ∼1.65 µm∼2.2 µmVan Der Velden et al., 1998; Lundy et al., 2013
T-tubules Absent Present Brette and Orchard, 2003; Yang et al., 2014a
Energy and force Mitochondria Near nuclei, small fraction Throughout cell; 20–40% of cell volume Schaper et al., 1985; Gherghiceanu et al., 2011
Energy Glycolysis ß-oxidation of fatty acid Lopaschuk and Jaswal, 2010; Kim et al., 2013
Contractile force 0.22 ±0.70 to 11.8 ±4.5 mN/mm251 ±8 mN/mm2Van Der Velden et al., 1998; Kita-Matsuo et al.,
2009; Zhang et al., 2013
Proliferation Early hPSC-CM: Yes Late hPSC-CM: No Considered non-proliferative Bergmann et al., 2009; Robertson et al., 2013
Calcium transients Inefﬁcient Efﬁcient Itzhaki et al., 2011
Slow Fast Yang et al., 2014a
AP(action potential) properties Upstroke velocity 15–50 V/s 180–400 V/s Dangman et al., 1982; Drouin et al., 1995; He
et al., 2003; Lundy et al., 2013
Resting membrane potential −20 to −60 mV −90 mV Drouin et al., 1995; Mummery et al., 2003;
Lundy et al., 2013
Conduction velocity 2.1–20 cm/s 41–84 cm/s Nanthakumar et al., 2007; Caspi et al., 2009;
Lee et al., 2012
Capacitance 5–30 pF 150 pF Drouin et al., 1995; Blazeski et al., 2012
Automaticity Spontaneous beating Quiescent Chen et al., 2009; Lundy et al., 2013
mRNA level Cav1.2 Similar to adult cardiomyocyte – Satin et al., 2008
Cavß1 20 fold lower than adult cardiomyocyte – Satin et al., 2008
RyR2 ∼1000 fold lower than adult cardiomyocyte – Satin et al., 2008
Ion channel density (pA/pF) INa −20 to −330 ∼ −50 Valdivia et al., 2005; Fatima et al., 2013;
Ivashchenko et al., 2013
ICaL −2.2 to −11 −2.3 to ∼ −10 Magyar et al., 2000; Er et al., 2009; Fu et al.,
2010; Otsuji et al., 2010
Ito 2.5–13.7 2.3–9.2 Beuckelmann et al., 1993; Wettwer et al.,
1994; Ma et al., 2011; Cordeiro et al., 2013
IKs 0.3–0.7 0.18 Virag et al., 2001; Otsuji et al., 2010; Ma et al.,
2011; Jonsson et al., 2012
IKr 0.4–0.8 0.6 Jost et al., 2009; Fu et al., 2011; Ma et al.,
IK1−0.6 to −3.4 ∼ −12 Schram et al., 2003; Sartiani et al., 2007
INCX 3.6–7.9 (Ca2+inward mode) 2.5–3 Weber et al., 2003; Fu et al., 2010
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Sun and Nunes Bioengineering Pro-maturation Strategies for hPSC-CMs
(Olivetti et al., 1996) with rod-like shape. In addition, hPSC-
CMs possess sparse, disorganized and shorter sarcomeres
(∼1.6 µm), and few or no transverse tubules (T-tubules).
Normal adult cardiomyocytes exhibit well-aligned, longer
sarcomeres (∼2.2 µm) characterized by the presence of Z
discs, and I-, H-, A-, and M-bands.
3) Metabolically, hPSC-CMs are characterized by a relatively low
number of mitochondria and a dependence on glycolysis as
opposed to a predominantly fatty acid metabolism in adult
cardiomyocytes (Yang et al., 2014a).
4) Functionally, hPSC-CMs display a force-generation capacity
(0.22 ±0.70 mN/mm2−11.8 ±4.5 mN/mm2) (Kita-
Matsuo et al., 2009; Zhang et al., 2013) comparable to fetal
cardiomyocytes (2nd trimester) (∼0.4 mN/mm2) (Ribeiro
et al., 2015) and much lower than adult (∼51 mN/mm2) (Van
Der Velden et al., 1998).
5) Electrophysiologically, hPSC-CMs show greater
heterogeneity and immaturity in their electrical properties
than adult cardiomyocytes including: (a) reduced electrical
excitability; (b) decreased excitation–contraction coupling
(ECC); (c) higher resting membrane potential (−20 to −60
mV vs. ∼ −90 mV); (d) low capacitance; (e) smaller upstroke
(15–50 vs. 180–400 V/s) and conduction velocity (2.1–20 vs.
41–84 cm/s); and (f) presence of automaticity (spontaneous
beating), which is found in early fetal cardiomyocytes and
later speciﬁc to pacemaker cells.
These immature features may limit hPSC-CM application and
highlight the need for the development of pro-maturation
strategies to obtain human adult cardiomyocytes in vitro. Given
the complexity of the cardiomyocyte structure and function, the
term “maturation” represents multi-faceted properties used to
evaluate their maturation state. However, the properties reported
in diﬀerent studies have often varied (Figure 1) making it diﬃcult
to draw a direct comparison.
STRATEGIES TO INDUCE HPSC-CM
Cardiomyocytes undergo a series of structural changes and
ultimately reach full maturity in the adult heart, which enables
them to fulﬁll their functional role. This development process is
long (years) and under complex regulation (Ahuja et al., 2007).
hPSC-CMs could mature to adult-like size and morphology
within 3 months post-transplantation into infarcted hearts
of non-human primates (Chong et al., 2014). Long-term
culture in vitro (80–120 days) has been suggested eﬀective in
improving the maturity of hPSC-CMs (Lundy et al., 2013).
However, this is very time-consuming and cost prohibitive.
More strategies to promote the maturity of hPSC-CMs include:
genetic manipulation (e.g., adenovirus-mediated overexpressing
of Kir2.1 Lieu et al., 2013), modulation of microRNAs (e.g.,
lentivirus-mediated overexpression of miR-1 Fu et al., 2011),
delivery of biochemical factors, such as triiodothyronine (Yang
et al., 2014b) and alpha-adrenergic agonist phenylephrine (Foldes
et al., 2011), induction of cell alignment in 3D tissues (Zhu et al.,
2014), and electrical and/or mechanical stimulation.
Of these, mechanical and electrical stimulation are major
biophysical cues that play critical roles in cardiomyocyte growth
and maturation during cardiac development and have been tested
as maturation cues for hPSC-CM. To replicate electromechanical
forces in vitro, hPSC-CMs are cultured in a biomimetic
environment comparable to native cardiac microarchitecture and
subjected to mechanical and/or electrical stimuli. The goal is
to promote the maturity of hPSC-CMs while improving our
understanding of the mechanisms responsible for the adaptive
changes of cardiac tissue under physiological and pathological
Mechanical force plays a critical role during development of
cardiac structure and function (Zimmermann, 2013). It may thus
be important to consider the presence of proper mechanical
signaling or cues when designing a platform for the maturation
of hPSC-CMs, regardless of whether it is in 2D or 3D. Mechanical
stimulation on cells can be implemented by adjusting the
substrate properties (stiﬀness/topography) and/or stretching.
These have been suggested to be eﬀective in improving the
maturation properties of hPSC-CMs.
The eﬀect of substrate rigidity on maturation can be
demonstrated by plating spontaneously contracting hPSC-CMs
on extracellular matrix (ECM) protein-coated tissue culture
surfaces where the matrix composition can be altered to
obtain physiological range of substrate stiﬀness. It’s been shown
that in a range of 4–80 kPa polyacrylamide hydrogels, the
highest diﬀerentiation eﬃciency using hESCs was achieved at
50 kPa (Hazeltine et al., 2014), and that contractile output
of cardiomyocytes increased in response to increased substrate
stiﬀness (4.4–99.7 kPa) (Hazeltine et al., 2012). Two-dimensional
substrates can also be micropatterned to improve hPSC-CM
alignment and sarcoplasmic reticulum (SR) Ca2+cycling (Rao
et al., 2013; Salick et al., 2014), which suggest improved
maturation. However, these two-dimensional structures lack
important features of the natural 3D environment that aﬀect the
Stretch is the major method used to deliver mechanical
stimuli to hPSC-CMs and generally done by applying external
mechanical stress to hPSC-CM constructs in a static (achieved
by increasing the stretch over time or directly to a ﬁxed distance)
or dynamic (mimicking the native cyclic mechanical stimulus on
the cardiac muscle) fashion.
Early studies to test the eﬀect of mechanical stress on
immature cardiomyocytes were performed by seeding cells
in collagen/Matrigel matrix, casting it in circular molds and,
following tissue compaction, the engineered heart tissues (EHT)
(Zimmermann et al., 2002) were subjected to uniaxial cyclic
stretch (2 Hz, 10% elongation). After 1 week, EHTs displayed
important hallmarks of mature myocardium: organized muscle
bundles with aligned sarcomeres and positive force-frequency
relationship (Endoh, 2004). Furthermore, these hEHTs show
a positive inotropic response to extracellular Ca2+and
isoproterenol (Streckfuss-Bomeke et al., 2013).
In another study, hEHTs were generated by mixing single-
cell hESC-CMs in a ﬁbrin/Matrigel gel and casting into a
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Sun and Nunes Bioengineering Pro-maturation Strategies for hPSC-CMs
FIGURE 1 | Schematic diagram illustrating the strategies to promote and the assessment of the maturation of human pluripotent stem cell
(hPSC)-derived cardiomyocytes (hPSC-CM). These approaches may be used individually or in any combination to promote hPSC-CM maturation. The
assessment of the maturation should be physiologically relevant, including readout from morphology (cell alignment, cell shape/size, sarcomeres and T-tubules), gene
expression (sarcomeric, ion channels and their regulators), and function (calcium handling, ECC, electrophysiology, contraction and transplantation).
12 ×3×3 mm agarose mold in which two elastic silicone
posts were inserted from above (Schaaf et al., 2011). Upon
compaction, the cardiac construct strip anchored to the posts
was subjected to static strain and displayed improved cell
alignment and sarcomeric organization compared with age-
matched EBs, and expressed connexin-43 but not in intercalated
disks. Transcription levels of β-MHC increased signiﬁcantly
over time in hEHTs but not in EBs. The hEHTs demonstrated
contractions 5–10 days after casting, reached regular (mean
0.5 Hz) and strong (mean 100 mN) contractions for up to
8 weeks. The constructs exhibited positive chronotropic and
inotropic response to increasing concentrations of extracellular
Ca2+(Schaaf et al., 2011).
Cardiac constructs were also generated by casting collagen-
based hPSC-CMs gels in a 20 mm ×3 mm channel, in which
the ends of the construct were anchored into nylon mesh tabs
attached to a deformable silicon ﬂoor of the well (tissue train,
Flexcell). Upon cell remodeling and gel contraction, the cardiac
constructs were held by the nylon tabs under static tension
or subjected to controlled cyclic stress (1 Hz, 5% elongation)
(Tulloch et al., 2011). After 4 days, there was improvement in cell
alignment and striations within the constructs. Cyclic stretch also
upregulated transcripts of β-MHC, cTnT, ANP, BNP, CACNA1C,
RYR2, and SERCA2 (Tulloch et al., 2011). Functionally, cardiac
constructs subjected to 3 weeks of static strain have increased
their active force in response to increased resting length (Tulloch
et al., 2011), analogous to Frank-Starling curves (an increase
in force with increased preload known as length-dependent
activation) (Glower et al., 1985).
Mihic et al. (2014) used cyclic mechanical stretch to
enhance the viability and functional maturation of hPSC-
CM tissue constructs prior to implantation into the damaged
myocardium. The constructs were generated by seeding hESC-
CMs in a 30 ×10 ×7 mm gelatin sponges. After 2 days of
compaction, the cardiac constructs were subjected to 3 days
of uniaxial cyclic stretch (1.25 Hz, 12% elongation). Compared
to unstretched controls, cyclically stretched cardiac constructs
exhibited increased number of cells, cell size and elongation,
increased expression of connexin-43, and upregulated mRNA
expression of MYH7, CACNA1C, HCN4, KCNH2, SCN5A, and
KCNJ2. Functionally, the cyclically stretched cardiac constructs
were demonstrated faster contraction rates with shorter calcium
Zhang et al. (2013) used a platform to promote hESC-
CMs alignment within cardiac patch via locally controlling the
direction of passive tension. hESC-CMs (48–90% purity) were
cultured for 2 weeks in a mixture of ﬁbrin and Matrigel in 7
×7 mm2polydimethylsiloxane (PDMS) molds with staggered
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Sun and Nunes Bioengineering Pro-maturation Strategies for hPSC-CMs
hexagonal posts (1.2 mm long) to generate a cardiac patches with
elliptical pores formed around the posts upon tissue compaction.
The resultant hESC-CMs in the 3D patches exhibited a maximal
conduction velocity of 25.1 cm/s, and longer sarcomeres (2.09
±0.02 vs. 1.77 ±0.01 µm), and enhanced expression of
genes involved in cardiac contractile function, including cTnT,
αMHC, CASQ2 and SERCA2 when compared to age and purity
matched hESC-CMs cultured in monolayers (Zhang et al., 2013).
Moreover, maximum contractile forces and active stresses of
cardiac patches were 3.0 ±1.1 mN and 11.8 ±4.5 mN/mm2,
respectively, and the patches were shown to generate Frank-
Starling curves with respect to both active and passive force
as well as positive inotropic response to isoproterenol (Zhang
et al., 2013). These author’s ﬁndings highlight the superiority of
3D vs. 2D culture models. However, no improvements in the
electrophysiological properties were reported.
These studies have established the signiﬁcance of mechanical
stimuli as a maturation cue for hPSC-CMs. However, it
should be noted that the contractile forces measured from
the aforementioned EHTs were related to the biomaterial
composition (e.g., collagen vs. ﬁbrin). Such material variability
may aﬀect the hPSC-CM phenotype, which consequently cause
the variation of functional readout including contractile force.
Furthermore, other variables in these mechanical stimulation
regimes, such as the cell culture condition and duration
of stimulation, makes it diﬃcult to determine an optimal
mechanical stress protocol for generating mature cardiac tissues.
Cardiomyocytes are rhythmically and synchronously contracting
in response to electrical signals. This process of converting
electrical signals into contraction (commonly known
as excitation-contraction coupling or ECC) requires the
coordinated activity of several ion channels (Liu et al., 2016).
The developmental changes in these ion channels are under
complex regulation and accompany changes in electrical
properties of cardiomyocytes across the fetal and postnatal
stages, with a speciﬁc electrophysiological “signature” in mature
adult cardiomyocytes. hPSC-CMs have been shown to be
electrophysiologically immature. Studies recapitulating in vitro
the electrical activity cardiomyocytes are exposed to in vivo have
demonstrated that electrical stimulation promotes aspects of
We have devised a platform called “biowire,” to mature hPSC-
CMs by combining 3D culture and electrical stimulation (Nunes
et al., 2013; Sun and Nunes, 2016). Biowires were generated by
culturing hPSC-CMs in collagen hydrogels around a surgical
suture to form cardiac tissues of ∼600 µm in diameter (Nunes
et al., 2013). Biowires were subjected to 7 days of electrical ﬁeld
stimulation (3 V/cm, 1 ms pulse, starting at 1 Hz with step-wise
increases to 3 or 6 Hz). At the endpoint, hPSC-CMs exhibited
properties compatible with cardiomyocyte maturation, such as
improved cell and myoﬁbril alignment, improved sarcomeric
banding, larger cardiomyocyte area and lower proliferation rates,
compared with age-matched EBs. Automaticity was signiﬁcantly
higher in EB-derived cardiomyocytes compared to control
biowires, which was comparable to that in biowires subjected
to the 6-Hz regimen. Electrical stimulation also signiﬁcantly
increased the conduction velocity of biowires from ∼11.5 to
18.5 cm/s. Biowires exposed to electrical stimulation also showed
increased Ca2+transient amplitudes vs. unstimulated controls.
hPSC-CMs in biowires exhibited improved hERG current and
inward rectiﬁer current (Ik1) densities, which were further
enhanced by electrical stimulation. This study revealed for the
ﬁrst time that these changes were dependent on the electrical
stimulation rate as evidenced by greater extent of maturation
obtained in the biowires exposed to the 6 Hz stimulation
ramp-up regimen (vs 3 Hz) (Nunes et al., 2013). However,
given the presence of the silk suture the force of contraction
generated by the hPSC-CMs could not be measured. The use of a
biodegradable suture may make this possible in the future.
Others have shown that hESC-CMs subjected to 2-week-long
electrical conditioning (2.5 V/cm, 1 Hz, 5 ms pulse) exhibited
lower spontaneous activity, hyperpolarized resting potential,
increased intracellular Ca2+transients, structured organization
of myoﬁlaments, and an upregulation of Kir2.1, CSQ2, junctin,
triadin, SERCA, Cav3, Amp2, MHC, and MLC genes (Lieu
et al., 2013). In another study, beating EBs seeded on gelatin-
coated plates and subjected to 4-day-long electrical stimulation
(6.6 V/cm, 1 Hz, 2 ms pulse) exhibited cell elongation,
increased action potential duration, increased Ca2+transients
and increased expression of cardiac-speciﬁc gene including
HCN1, MLC2V, SCN5A, SERCA, Kv4.3, and GATA4 (Chan et al.,
In a recent study, EBs diﬀerentiated from hPSCs were
subjected to electrical conditioning (5 V/cm, 0.5, 1 and 2
Hz, 2 ms pulse) continuously for 7 days (Eng et al., 2016).
Such electrical stimulation enhanced connexin expression and
sarcomeric structure. Cardiomyocytes adapted their autonomous
beating rate to the frequency at which they were stimulated, an
eﬀect mediated by the emergence of a rapidly depolarizing cell
type, and the expression of hERG. The resultant cardiomyocytes
were robust and could maintain the adapted beating rates for up
to 2 weeks after the cessation of electrical stimulation (Eng et al.,
While electrical stimulation has consistently improved the
maturation of hPSC-CMs, one possible drawback of utilizing
electrical stimulation is the limited scalability. This may not be
of concern for its utilization in drug screening platforms but
may hinder its application in cell maturation for regenerative
Combined Mechanical and Electrical
Eﬀorts have also been made to examine the eﬀect of combining
mechanical and electrical stimulation, sequentially or
concurrently, to hPSC-CM constructs. Hirt et al. (2014)
generated spontaneously beating ﬁbrin/Matrigel-based hPSC-
CM constructs with static stretch and subjected them to electrical
ﬁeld stimulation (2 V/cm, 4 ms pulse, 2 Hz for 1 week and
1.5 Hz thereafter) for at least 10 days. This increased cell
alignment, sarcomere organization, Ca2+-response curves,
force generation and inotropic response to β-adrenergic
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Sun and Nunes Bioengineering Pro-maturation Strategies for hPSC-CMs
stimulation while decreasing automaticity. (Hirt et al.,
In another study, hPSC-CMs were embedded into a collagen-
based scaﬀold and then subjected to static stress for 2 or 1 week
of static stress and 1 week of combined static stress and electrical
pacing (5 V/cm, 2 Hz, 5 ms pulse) (Ruan et al., 2016). Compared
to no stress/no pacing controls, 2-week static stress conditioning
promoted cell alignment, passive stiﬀness, cardiac hypertrophy,
and increased contractility of hPSC-CM constructs (0.63 ±0.10
mN/mm2vs. 0.055 ±0.009 mN/mm2). The contractility of the
constructs could be further increased by combining stretch with
1-week electrical stimulation (1.34 ±0.19 mN/mm2). Combined
static stress and electrical stimulation enhanced expression of
SR-related proteins (RYR2 and SERCA2) (Ruan et al., 2016).
CONCLUSIONS AND FUTURE
The eﬀorts to mimic native biophysical stimulation to mature
hPSC-CMs have led to a number of eﬀective strategies to
mature hPSC-CMs and advance our understanding of how
these cues aﬀect cardiomyocyte structure and function. However,
the properties assessed often varied between studies making
it diﬃcult to draw a direct comparison between the diﬀerent
strategies. This is accentuated by the lack of uniformity in
cardiomyocyte maturation in artiﬁcial, in vitro settings where
electrical stimulation seems to have a stronger impact on
electrical properties while mechanical stimulation improves
structural components and force generation with smaller impact
on electrical properties. This argues for a homogeneity in the
parameters utilized as functional readouts (electrophysiology,
calcium dynamics, force of contraction and ultrastructure).
Although progress has been made, an adult-like phenotype in
vitro has yet to be reported. This can have multiple limitations
regarding application. First, the maturation status of hPSC-CMs
should be staged and documented depending on the potential
application sceneries. For example, for myocardial infarction
(MI) therapy, less mature cardiomyocytes might adapt better for
transplantation into the infarcted myocardium (Reinecke et al.,
1999). However, the best-deﬁned maturation stage of hPSC-CMs
for transplantation into MI remains to be determined.
Second, the hPSC-CMs obtained from existing cardiac
diﬀerentiation protocols are a mixed population of ventricular-,
atrial-, and nodal-like cells. Such heterogeneity represents
a limitation for certain applications, e.g., transplantation of
high purity of ventricular cardiomyocytes to potentially avoid
tachyarrhythmias caused by spontaneously ﬁring (nodal-like)
cells; and high throughput (HTS) drug testing platforms for
cardiac drug responses.
Third, the signiﬁcance of the in vivo environment for the
maturation of cardiomyocytes should be noted. Immature hPSC-
CMs diﬀerentiated in vitro could mature to adult size and
morphology after transplantation into the infarcted hearts of
non-human primates (Chong et al., 2014).
Proper cardiac development and function requires other
cell types, such as ﬁbroblasts, endothelial, and smooth muscle
cells that may have an impact in cardiomyocyte maturation.
While there is still controversy regarding whether non-
cardiomyocytes may promote hPSC-CM maturation via
secretion of undeﬁned factors (Kim et al., 2010; Lundy et al.,
2013), a full understanding of these interactions may help
to uncover unknown cues, which could then be used to
promote hPSC-CM maturation in the absence of a speciﬁc
hPSC-CMs have shown great promises in various applications
including cardiac development, regenerative medicine, disease
modeling, and drug testing/screening/discovery. The generation
of a large number of mature hPSC-CMs is essential to achieve
these goals. Importantly, these approaches are not mutually
exclusive (Figure 1) and there’s been a trend to combine
the existing strategies to obtain more eﬀective maturation.
The combination of mechanical and electrical stimulation has
shown possible synergistic eﬀects with a 2-fold increase in
contractility (Ruan et al., 2016). This trend should lead to exciting
discoveries regarding hPSC-CM maturation and possibly the
achievement of adult-like cardiomyocytes in vitro for the ﬁrst
XS and SN conceived and wrote the manuscript.
This work was supported by a grant-in-aid from the Heart and
Stroke Foundation of Canada (G-14-0006265), operating grants
from the Canadian Institutes of Health Research (137352 and
143066) and a J.P. Bickell foundation grant (1013821) to SN.
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Conﬂict of Interest Statement: The authors declare that the research was
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