Mimicking heart disease in a dish
Kijlstra, Jan David
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Publication date: 2018
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Kijlstra, J. D. (2018). Mimicking heart disease in a dish: Cardiac disease modelling through functional analysis of human stem cell derived cardiomyocytes. Rijksuniversiteit Groningen.
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Benchmarking the maturation
of human pluripotent stem
cell-derived cardiomyocytes:
where is the finish line and
where are we?
J. David Kijlstra1, Peter van der Meer1, and Ibrahim J. Domian2,3,4
1 Department of Experimental Cardiology, University Medical Center Groningen, University of Groningen, Groningen, 9713 GZ, The Netherlands
2 Cardiovascular Research Center, Massachusetts General Hospital, Boston, MA 02114, USA 3 Harvard Medical School, Boston, MA 02115, USA
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Introduction
Human pluripotent stem cell-derived cardiomyocytes (hPSC-CMs) hold great promise for cardiovascular disease modelling, cardiovascular drug screening, and applications in regenerative medicine. Since the development of the first embryonic stem cell lines and the subsequent discovery of induced pluripotent stem cells less than one decade later, advances in the use of hPSC-CMs have partly fulfilled this promise. hPSC-CMs recapitulate the physiology of in vivo cardiomyocytes to a certain extent. hPSC-CMs express most of the cardiac-specific ion channels and currents, have a functional contractile apparatus and respond to cardioactive drugs. As such, hPSC-CMs have been used to model several cardiovascular diseases including hypertrophic cardiomyopathy and long QT syndrome.[1,2] Additionally, in proof-of-principle regenerative medicine studies, hPSC-CMs were used in animal models of myocardial infarction and have been shown to integrate with the native myocardial tissue.[3] However, thus far these regenerative applications have shown little improvement in cardiac function, even if hPSC-CMs were able to integrate in vivo.
The use of hPSC-CMs for clinical and preclinical applications has been limited by the immature phenotype of these cells as previously described in several extensive reviews.[4-15] Compared to adult CMs, hPSC-CMs are immature with respect to their morphology, their sarcomeric structure and ability to generate force, their electrophysiology and calcium handling, and their metabolic phenotype. Many research groups have explored various strategies to mature hPSC-CMs and to some extent have been successful in these attempts. However, critical questions remain about the relationship of partly matured hPSC-CMs to their native counterparts. To what extent does a cell population engineered in vitro resemble the corresponding target cell or tissue in both molecular and functional terms? The systematic analysis of cardiomyocyte maturation has been difficult to quantify and the comparison of maturation approaches across platforms and across manuscripts has been similarly difficult to evaluate. Next to the lack of a standard framework to assess CM maturation, there has also been an absence of tools that allow for the rapid integrated analysis of several functional parameters of hPSC-CMs. Such tools could be crucial to facilitate an improved phenotyping of the maturation status of hPSC-CMs. Herein, we first review the existing literature on the phenotype of cardiac tissue or CMs isolated from human hearts at several developmental stages spanning the early fetal stage to adulthood. In so doing, we quantify several features of cardiac form and function from the fetal to adult stages. Next, we review the strategies that have been implemented to mature hPSC-CMs using human in vivo data as a benchmark. Importantly, we offer a framework for the semi-quantitative comparison of the maturation state of hPSC-CMs. Based on this framework we have developed the Maturation Score, that can be used to compare the maturation of hPSC-CMs to other hPSC-CMs and adult CMs. Finally, we demonstrate the use of this Maturation Score to compare the maturation achieved in hPSC-CMs by various strategies described in this review. This Maturation Score can be developed into an open-source web-based tool for utilization by other researchers within this field of study.
In vivo Cardiac Maturation
Mice are the mainstay of in vivo mammalian developmental biology and in many respects mirror human development remarkably well. This conservation is reflected in the overall homology of the mouse and human genomes. Despite this conservation there exist significant differences between mice and humans in cardiac development and maturation as well as cardiac physiology. These differences can be expected since the two species diverged approximately 75 million years ago and differ in the size and baseline hemodynamics. Advances in human stem cell biology over the past decade make it now possible to study human cardiovascular biology in vitro and opens new opportunities for regenerative cardiovascular medicine. In that context, it is increasingly important to focus on human (as opposed to rodent) myocardial maturation. Accordingly, to establish a baseline benchmark against which human PSC derived CMs can be measured, human data should be primarily used (Supplementary Table 1).
Structure
The heart is the first organ to form during embryonic development and subsequently CMs are amongst the first specialized cell types. Around day 20 after fertilization, the first CMs form in the primitive heart tube. The heart then goes through cardiac looping and septation and by week 6 of gestation the four-chamber heart is formed. Subsequently, the heart grows through CM proliferation until birth while the CMs remain small in size.
Postnatally, there is a major transition from hyperplastic to hypertrophic growth. When this transition occurs, how abruptly it occurs, and to what extent CMs maintain their proliferative capacity during adolescence and adulthood, has been intensely investigated for more than 150 years. Yet, the answers still remain unclear.[16] Nonetheless, two recent studies using different techniques to analyze the same donor hearts reached consensus that new CMs are generated during childhood and that in adult humans this activity is extremely low.[17,18] In the first year after birth, CMs have a volume of about 4,500 μm3. With hypertrophic growth, CMs will subsequently increase in volume by ~10-fold and reach dimensions of 20 by 140 μm in adulthood.[17-19]
The shape of CMs progresses from round in early fetal CMs to the characteristic rod-shape observed in adult CMs with a length-to-width ratio of 7.5.[19,20] The exact moment of this transition in human development is difficult to pinpoint. In rats, it occurs predominantly in the neonatal period.[21] Due to DNA replication without mitosis or cytokinesis, adult CMs often exhibit polyploidy and sometimes binucleation. Although considerable interindividual variance exists, on average 80% of CM nuclei are diploid (2n) until 8 years of age after which 60% of nuclei become tetraploid (4n). [22-24] The fraction of binucleated CMs in adult myocardium is estimated to be 25% at birth and remains steady throughout life.[17,18,25]
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Sarcomeres and Contractility
Sarcomeres, and the molecular interaction between actin and myosin filaments within them, are at the basis of CM contraction. The sarcomeres are small in fetal CMs but grow larger and become highly organized in the adult CM. The sarcomere length expands from 1.8 μm in fetal CMs at 14-19 weeks of gestation [26] to ~2.06 μm in adult CMs.[19,27] Many myofibrillar proteins in the sarcomeres operate in concert to produce a CM contraction, and their partial overlap produces distinct bands on electron microscopy. Z-discs delineate the lateral borders of sarcomeres and act as anchor points for actin filaments, they are formed in the early fetal stage. Next to the Z-disc is the pale I-band where the thin actin filaments do not overlap with thick myosin filaments. In between the I-bands is the darker A-band containing the full length of the thick myosin filaments. A and I-bands are formed in the late fetal stage.[28] In the middle of the A-band is a lighter region called the H-zone, where myosin filaments are not superimposed by actin filaments. At the very center of the H-zone and the sarcomere is the darker M-band, consisting of cytoskeleton proteins that act as anchor points for the myosin filaments. As H-zone and M-bands have not been demonstrated in fetal cardiac tissue, they likely emerge postnatally.
The isoforms of multiple myofibrillar proteins switch at various points in development to modulate contractile function. As such they have previously been proposed as markers of maturation. Titin, a large spring-like filament, is the main contributor to passive tension of CMs. Perinatal titin-isoform switching was studied in various mammalian species and found to follow a common pattern: the more compliant N2BA isoform is mostly replaced by the stiffer isoform N2B by 18 days after birth, thereby increasing passive stiffness of the myocardium.[29-31] Human myocardium presumably follows the same pattern, although this has not been shown yet. In the human heart there is significantly more expression of (slow) β-myosin heavy chain (β-MHC) than (fast) α-MHC. Already by 7 weeks gestation, β-MHC comprises ~90% of total MHC protein in the left ventricle and this increases further during maturation.[32] Notably, the β-MHC/α-MHC ratio also increases with heart disease, complicating its utility as a marker of maturation.[32,33] Troponin I, the inhibitory subunit of the troponin complex, also undergoes an isoform switch during maturation.[34] Slow skeletal troponin I (ssTnI) is the dominant isoform in in the fetal heart, while small amounts of cardiac troponin I (cTnI) are also present and increase during gestation. During the first year of life [35,36], ssTnI is replaced by cTnI. In rodents, this switch results in decreased Ca2+sensitivity of the myofilaments.[37-40] Unlike
the limitation posed by MHC, troponin I expression does not change with heart disease. Finally, the expression of myosin light chain (MLC) isoforms, MLC2a and MLC2v, can be used to differentiate between atrial and ventricular CMs. Although data from rodent models is plentiful, there is to our knowledge only one study in human samples on the differential expression of these isoforms. This study demonstrated that MLC2a is expressed in all heart chambers whereas MLC2v is expressed only in the ventricles.[41] The expression of these isoforms did not change in the time range of this study (4-18 weeks of gestation), reducing their utility as potential maturation markers.
The adult myocardium generates a maximum force of ~45 mN/mm2.[42–44] Cardiac force generation increases with maturation. In the fetal stage at 14-19 weeks, CMs contract with a force of just 0.4 mN/mm2 [26,45]. Although this increases to 1.4 mN/mm2 for newborn (3-8 days) and 1.2 mN/mm2for infant (3-7 months) cardiac tissue, it remains one order of magnitude lower than
adult levels.[46] Interestingly, force generation per surface area is consistently reported higher for myofibrils than CMs or cardiac tissue.[26,45,47,48] This discrepancy could be attributed to the lack of extramyofibrillar space in myofibril preparations compared to CMs or cardiac tissue. It should be noted that at 19 weeks of gestation, the maximum force generation of myofibrils already reaches half of the adult level, as opposed to about 1% of adult level for fetal CMs of similar age.
When assaying cardiac force generation, it is critical to note that this readout is heavily impacted by the available calcium concentration, the pacing frequency, and the amount of stretch exerted on cardiac tissue. First, optimal calcium concentrations allow for maximum actin-myosin interaction in the sarcomeres. Additionally, healthy adult myocardium exhibits a positive force-frequency relationship, meaning that within a physiological range, an increase in frequency is accompanied by an increase in force generation, thus allowing for increased cardiac performance during exercise.[49] The force-frequency relationship is flat in newborn ventricular tissue and becomes positive during infancy.[46] However, a reversal back to a flat or negative force-frequency relationship has been observed in cardiac disease.[50,51] Finally, when cardiac tissue is stretched (such as with increased preload on the heart), this leads to a stronger subsequent contraction, which is called the Frank-Starling mechanism. This mechanism is apparent in fetal heart by 10-15 weeks gestation.[52]
Electrophysiology and Calcium Handling
The heart, similar to nerves and skeletal muscle, is activated by depolarization of the CM cell membrane. This initial activation of cardiac tissue results in intracellular calcium release and ultimately contraction, through a process called excitation-contraction coupling. The cardiac adult ventricular action potential has a distinct shape, consisting of five phases, that results from the temporally orchestrated influx and efflux of positive ions through various ion channels on the cell membrane.
In phase 4, when the CM is at rest, the membrane potential is maintained at -90 mV. In phase 0, a rapid influx of Na+ leads to a fast depolarization of the membrane to +50 mV.During this depolarization, voltage-gated L-type calcium channels also open, allowing for an influx of Ca2+. In phase 1, the influx of Na+ stops and there is a brief flow of K+ out of the cell, resulting in a slight repolarization of the membrane. During the plateau of phase 2, the inflow of Ca2+ that started in phase 0 is balanced by the outflow of K+, keeping the membrane potential depolarized for a few hundred milliseconds. Full repolarization occurs during phase 3, as the inflow of Ca2+stops while the outflow of K+ continues.
To date, there is no human data on developmental changes in the current densities that result in a mature cardiac electrophysiological phenotype. Data from numerous animal studies does suggest that significant changes occur between the fetal and adult heart. For example, there is a decrease in the funny current (If ) that coincides with the termination of spontaneous beating in neonatal murine CMs.
Several functional parameters of cardiac electrophysiology were obtained in human fetal tissue that do suggest a similar developmental pattern occurs in humans and animals. The maximum depolarization velocity was reported to increase ~30-fold from the fetal to the adult stage.[20,53,54] Similarly, the action potential amplitude increases between the fetal and adult stages.[20,53,54] The reported action potential durations of adult cardiac tissue vary widely, even at similar pacing
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frequencies and with similar extracellular ion availabilities.[53,54] However, the reported value ofaction potential duration for fetal cardiomyocytes of 370 ms is comparable to that of adult cardiac tissue.[20] The available studies offer conflicting evidence about the resting membrane potential in fetal cardiomyocytes, it has been reported as both ~50% higher (i.e. less negative) [20] and equivalent to adult cardiac tissue.[53–55] It should be noted that this discrepancy could also affect the fetal values of maximum depolarization velocity and action potential amplitude.
Adult CMs are connected at their longitudinal edges by intercalated discs that contain desmosomes, adherens junctions, and gap junctions, which allow rapid transmission of contractile forces and electrical signals through the cardiac tissue. Connexin-43 (Cx43) is the dominant gap junction protein in adult cardiac tissue and its mRNA expression is similar in fetal, neonatal, and adult tissue. [56] However, the spatial distribution of Cx43 does change significantly during development. It is distributed along all sides of the cell membrane in fetal and neonatal CMs but localizes predominantly to the intercalated discs by 7 years of age.[57] The conduction velocity represents a functional measure of the electrical connectivity of cardiac tissue. This measure increases from ~43 cm/s in fetal tissue to ~65 cm/s in adult tissue.[55,58,59]
Transverse tubules (T-tubules) are deep protusions of the cell membrane that occur at intervals of ~2 μm. T-tubules allow for rapid transduction of the action potential throughout the entire cell. They are closely associated with the intracellular calcium stores in the sarcoplasmic reticulum, thus facilitating synchronized calcium release and effective excitation-contraction coupling. Although T-tubules only form postnatally in rodents [60], an electron microscopy study has suggested that in humans T-tubules form in the late fetal stage at 32 weeks gestation.[28] Two recent studies have quantified the density of T-tubules in adult CMs. Upon staining with a membrane dye, T-tubules were found to cover 8-23% of the surface area in slices from the middle of the cells.[61,62]
In adult CMs, action potentials propagate down the transverse tubules where they activate the voltage-gated L-type calcium channels, causing an influx of calcium through the cell membrane. This initial flux of calcium triggers the release of more calcium from the sarcoplasmic reticulum via the ryanodine receptors, a process known as calcium-induced-calcium release. The cytosolic calcium binds to troponin, causing a translocation of tropomyosin on the actin filament [63], which allows for actin-myosin interaction and ultimately contraction of the CM. This entire process is termed excitation-contraction coupling. The elevated cytosolic calcium causes the sarcoplasmic– endoplasmic reticulum Ca2+-ATPase and Na–Ca exchanger to start pumping the calcium back into respectively the SR and extracellular space. This return to the resting calcium level leads to CM relaxation.[49]
In adult cardiac tissue at 1Hz frequency, the cytosolic calcium concentration increases by about 8-fold during systole.[64-67] The sarcoplasmic reticulum holds a large reserve of calcium, which can be released through exposure to caffeine. After caffeine stimulation, the sarcoplasmic reticulum contributes 76% of the total calcium release. The remainder of calcium influx occurs through the Na-Ca2exchanger, which momentarily reverses ion flux during the action potential upstroke.[64] A regular calcium transient lasts about 450 ms.[61,64,68] There is scarce data available about the calcium handling of cardiac tissue during human development. Nevertheless, it has been shown that the majority of fetal CMs at 16-18 weeks of gestation are caffeine-responsive. Western blot
experiments also indicate an upregulation of sarcoplasmic-endoplasmic reticulum Ca2+-ATPase and a strong downregulation of Na-Ca exchanger during development.[69,70] This is in accordance with the model of calcium handling development that emerges from animal data.[71]
Metabolism
The energy demand of the adult heart is immense compared to most other tissues because of the energy-intensive contractile cycles. The minimal storage of ATP in CMs is only sufficient to sustain the heart beat for a few seconds, highlighting the need for continuous ATP production.[72] As such, mitochondria make up ~25% of the cytoplasmic volume in adult CMs to supply the required ATP for all this sarcomeric activity.[73,74] Although no human data exists on the volumetric density of mitochondria in the developing heart, the expression and activity of various mitochondrial enzymes increases by about 3-fold from 7 weeks of gestation to the neonate, likely due to an increase in mitochondrial number.[75] Likewise, the mitochondrial density in rats increases significantly between the early fetal and neonatal stages.[76]
In adult CMs, the mitochondria are localized between sarcomeres, around the nucleus, and in the subsarcolemma.[74] By contrast, the lower number of mitochondria in fetal CMs are dispersed throughout the cytoplasm and are not clearly spatially distributed in relation to the immature contractile machinery.[28]
The preferred metabolic substrate of the adult myocardium is fatty acid, which along with glucose and lactate is extracted in large amounts from the coronary circulation.[77] About 85% of the extracted fatty acids are used in mitochondrial oxidative phosphorylation compared to only 20-25% for glucose and lactate.[78,79] In various mammalian species the majority of ATP is produced by fatty acid oxidation.[80,81] ATP production in the human heart is likely fueled predominantly by fatty acid oxidation as well, based on the demonstrated extraction and utilization rate of fatty acids. However, the exact contribution of fatty acid oxidation to total ATP production in the adult human heart remains difficult to pin down. Moreover, the human heart remains an omnivore and various metabolic interventions can alter the substrate utilization.[82]
To our knowledge, there is no information about the metabolic substrate use of human fetal or neonatal cardiac tissue. However, there is ample data available from studies on various other mammalian species and human cardiac development likely follows a similar pattern. In the rodent fetal stage, the cardiac workload is relatively low, circulating glucose and lactate levels are high, and fatty acid levels are low. The heart primarily depends on the inefficient processes of anaerobic glycolysis and lactate oxidation.[83,84] After birth (d0-d14), the energy demand of the heart increases, lactate levels fall, glucose levels are maintained, and fatty acid levels rapidly increase.[85] Together, these changes are correlated with a switch from glycolysis to oxidative metabolism with fatty acids as the major contributor.[86-88]
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Strategies to Induce hPSC-CM Maturation
Various strategies have been applied towards the maturation of hPSC-CMs. Below we provide an overview of these strategies and the maturation markers that were assessed. This data will serve as the basis for a framework to compare the results obtained with various strategies. In Supplementary Table 2 we show an overview of this data on hPSC-CM maturation together with the data of adult human CMs.
Long-Term Culture
In vivo, CMs take many years to reach their adult phenotype. hPSC-CMs by comparison currently are differentiated on a much shorter time scale. Beating hPSC-CMs can be observed at 7 days after the start of differentiation [89]and these beating CMs are often experimentally used after about 20 days of differentiation. Several studies have tried to promote the maturation of PSC-CMs by long term culture. After 2 months, hPSC-CMs withdrew from the cell cycle and displayed a number of maturation markers and became more rod-like with a slight increase in aspect ratio.[89] The surface area of hPSC-CMs also increased, although there is a large degree of variability in the surface area reported at baseline and after prolonged culture.[89] The percentage of multinucleated cells increased significantly to approach the adult value after 4 months.[90] Ultrastructural analysis revealed increased sarcomeric organization with the appearance of Z-discs, A- and I bands, and H-zones after 2-4 months of culture.[89,90] Additionally, M-lines were found in a minority of CMs after a 12 month culture period.[91] Surprisingly, the force generation of hPSC-CMs did not increase after 2-3 months in culture [92,93], although an increased fractional shortening was reported after 4 months in culture.[90]
Late-stage hPSC-CMs also showed significantly faster calcium transient kinetics.[91] Additionally, two studies demonstrated electrophysiological maturation with long-term culture.[90,94] The AP upstroke Vmax and APD90 increased while the RMP established a lower baseline, indicative of maturation. The APA was quite similar to the adult value in both studies even before the prolonged culture period. Another study showed that the conduction velocity in hPSC-CM cell sheets increased significantly after 2 months but was still much lower than fetal and adult values.[95] Finally, Lundy et al. also demonstrated that mitochondria were localized in proximity to the myofibrils, a sign of metabolic maturation.
These data demonstrate the partial structural and functional maturation of hPSC-CMs with time, implying either a timekeeping mechanism or a time-dependent signaling pathway. Long-term culture offers limited throughput however. Therefore, many laboratories are attempting other approaches to both accelerate and improve the maturation process.
Electrical Stimulation
Adult CMs are quiescent but in vivo they are continuously and rhythmically exposed to electrical activity to contract. Several studies have recapitulated this electrical stimulation that occurs in
vivo to investigate its effect on hPSC-CM maturation. A caveat to most in vitro pacing protocols is
are activated by an action potential propagating from neighboring cells. Structurally, electrical stimulation somewhat enhances maturation. The aspect ratio of hPSC-CMs increased in response to electrical stimulation both in embryoid bodies [96] where the cells were quite round before stimulation and in engineered heart tissue (EHT) [97] where the cells were already more rod-like at baseline. Another study on hPSC-CMs seeded on a surgical suture to form cardiac biowires did not demonstrate increases in aspect ratio with electrical stimulation however.[98] With regards to CM hypertrophy, the data is likewise conflicting. The same study in cardiac biowires showed unchanged cell surface area after electrical stimulation. Another study using embryoid bodies found a slight increase in cell surface area although the reported before-and-after values were very low.[99] A third study where EHTs grown from early-stage hPSC-CMs were paced with increasing frequency for 2 weeks demonstrated a doubling of surface area.[100] Electrical stimulation furthermore failed to decrease proliferation rates in the biowire system. On ultrastructural analysis, electrical stimulation led to the appearance of sarcomeric H-zones in cardiac biowires and in EHTs it even led to the appearance of all sarcomeric zones including M-lines.[98,100] Sarcomeric maturation was also demonstrated by a slight force increase in cardiac microtissues and EHTs, although the reported values were still respectively fetal-like and infant-like.[100,101] Further maturation of EHTs after pacing was demonstrated by a positive force-frequency relationship.[100] Pacing appears to mature only part of the electrophysiological properties of hPSC-CMs. Most of the studies did not find any changes in action potential parameters after electrical stimulation of cardiac EBs, biowires and EHTs.[102] Conversely, the conduction velocity did increase by 50% in the biowires after pacing for 7 days at 6Hz and it more than doubled in EHTs after pacing for 3 weeks.[98,100] In EHTs, this increase in conduction velocity was also associated with a polarization of connexin 43, a feature of advanced cardiac maturation.[100] Interestingly, hPSC-CMs adapt their spontaneous beating rate to the pacing frequency even after cessation of the electric stimuli.[99] The amplitude of the calcium transient increased both with and without caffeine. Likewise, the release and reuptake rates of calcium were higher. Taken together this indicates improved function of the SR and improved calcium handling after electrical stimulation. Electrical stimulation also increased the presence of mitochondria next to the sarcomeres in cardiac biowires and EHTs. Finally, pacing increased the mitochondrial density in EHTs.
Biochemical Cues
Hormonal Stimulation
Several hormones play crucial roles in cardiac development and correspondingly they have been identified as inducing maturation of hPSC-CMs. T3 exposure increased cell size and sarcomere length, reduced proliferation, and improved calcium handling, mitochondrial function and consequently contractility of hPSC-CMs.[103] Another study investigated various hormonal factors and again found hPSC-CM maturation after T3 exposure.[104] The effects of T3 were further improved by the addition of IGF-1 and dexamethasone. This cocktail also made the hPSC-CMs amenable to further maturation with adrenergic stimulation. Commercially available T3 based maturation medium (Pluriomics BV) also increased force generation, improved sarcomeric structure, and matured the action potential characteristics.[26] T3 exposure did not suffice to switch the isotype of troponin I, another hallmark of sarcomeric maturation.[34]
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Metabolic SubstrateSince fatty acid oxidation is the preferred mode of ATP production in the adult heart, two recent studies have examined the effect of changing the metabolic substrate on the maturation of hPSC-CMs. Standard hPSC-CM culture media contains high levels of glucose without fatty acids. When hPSC-CMs in 2D culture were fed with glucose free media that enriched with fatty acids, a glycolytic-to-oxidative metabolic shift occurred. Subsequently, this metabolic shift was associated with improved structural and functional maturation of the cells. After 20 days of culture in the glucose-free media, hPSC-CMs became more rod-like, showed increased sarcomeric organization, and were more often binucleated. Functionally, hPSC-CMs demonstrated faster calcium transient kinetics, improved electrophysiology, higher mitochondrial respiration, and stronger contractions.[105] Mills et al. observed a similar metabolic shift upon exposure of hPSC-CMs embedded in cardiac organoids to lipid-enriched media. Additionally, they found a decrease in proliferation, reduced contraction and relaxation times, and an inotropic response to isoprenaline. However, no changes were observed in CM structure, calcium transient kinetics or electrophysiology.[106] This discrepancy could be due to the culture systems used. Mills et al. reported significant increases in maturation in the cardiac organoids compared to cells cultured in 2D, therefore it could be that the metabolic shift induced by media change was unable to cause further maturation of several markers.
Genetic Manipulation
Recent investigations have shed light on the transcriptome during normal cardiac development and informed efforts to force the maturation of hPSC-CMs by mimicking the natural gene expression program and its epigenetic regulation.[107] Kuppusamy et al. performed miRNA sequencing experiments on hPSC-CMs that were cultured for 13.5 months. They found an upregulation of the let7 family of miRNAs. Subsequent overexpression of let7 miRNAs in young hPSC-CMs induced a metabolic shift towards FA oxidation, a matured morphology and increased force generation. Similarly, co-culture of CMs with endothelial cells (EC) was found to induce maturation and the upregulation of several miRNAs. These miRNAs also prompted a matured morphology and electrophysiology of hPSC-CMs independently of co-culture with ECs.[108] In another example, miRNA-1 was found to be one of the most differentially expressed miRNAs when comparing adult heart tissue to hESCs. Addition of miRNA to hPSC-CMs slightly furthered their electrophysiological maturation.[109]
Another approach is the direct lentiviral-induced overexpression of genes that are insufficiently upregulated in immature hPSC-CMs. Indeed, overexpression of Kir2.1 channels hyperpolarized the RMP and prevented the automaticity that is characteristic of immature hPSC-CMs. Sarcomeric gene expression showed a reverse pattern towards immaturity however, likely due to the lack of contractile activity.[110] As such, the forced expression of genetic factors should be thoughtfully combined with other maturation cues to reach their full potential.
Co-Culture With Other Cell Types
Although CMs compose 80% of the adult myocardial volume, they account for only 30% of the total cell number.[111] The most common non-myocytes in the heart are fibroblasts and endothelial cells
which provide respectively structural support and vascularization of the myocardium. As such, it has been hypothesized that these cells are essential for proper function of engineered heart tissue. Indeed, multiple studies found that EHTs composed of a pure hPSC-CM population are unable to compact the extracellular matrix and as a result fail to form stable and integrated cardiac tissue. Conversely, the addition non-myocytes results in compact tissues with a higher cell density and improved alignment of all cells.[112-114] Furthermore, the CMs in these co-cultured EHTs display signs of maturation as indicated by a decrease in proliferation [114], increased sarcomeric organization and sarcomere length [115], and improved electrophysiology.[112,116] These alterations in CM phenotype do not consistently result in increased force generation however.[113-115]
Mechanical Stimulation
Cells within the native heart are subjected to several forms of mechanical stress. The inflow of blood during diastole creates cyclic stretch, the stiffness of the ECM and surrounding cells creates static stretch, and the laminar blood flow creates shear stress, mostly on the endothelial cells. These forces have been reproduced in vitro through the static or cyclic stretching of hPSC-CMs [117-120], culturing hPSC-CMs on substrates of varying stiffness [93,121]or the laminar flow of media over hPSC-CMs.[122] Static stretch has been shown to induce hypertrophy of hPSC-CMS.[118] Both static and cyclic stretch resulted in alignment of the hPSC-CMs in EHTs and concurrently maturation of contractility with increases in force generation and an improved force-frequency relationship. [117,118] This increased contractility was most pronounced in the cyclic stretch group, possibly due to improvements in the calcium handling. Cyclic stretch was found to produce higher peak fluorescence of the calcium signal indicating an enhanced function of the SR, whereas static stretch did not.[117] Shen et al. combined the application of cyclic stretch and shear stress by laminar flow of media in a bioreactor which caused a significant increase in sarcomere length.[122]
The standard polystyrene cell-culturing surface for hPSC-CMs in 2D is far more rigid (~3000 MPa) than the stiffness that native CMs are exposed to during development (5 kPa) and in adulthood (15 kPa). Soft substrates allow for shortening of hPSC-CMs in culture and measurements of force generation based on this shortening against a defined stiffness. The effect of exposing hPSC-CMs to substrates of varying stiffness on their force generation is somewhat unclear. One study reported that between the range of 4.4 kPa to 99.7 kPa, hPSC-CMs produce more force when exposed to higher stiffness, up to levels comparable with adult heart tissue when exposed to 99.7 kPa.[93] In contrast, another study demonstrated that the sarcomeric structure of hPSC-CMs on substrates of 35 kPa was disrupted and as a result the force generation was reduced by 90% compared to hPSC-CMs on softer substrates.[121] Similarly to what was found in EHTs, exposing hPSC-CMs to mechanical stress in 2D culture increased their surface area.[93]
Extracellular Matrix Composition and Topographical Cues
Extracellular Matrix Composition
The adult cardiac extracellular matrix is composed of a mixture of collagens, laminins, fibronectins, and a range of various other glycans and proteoglycans.[123] Extracellular matrix resembling this mixture may be well suited for hPSC-CM culture. Matrigel is a complex mixture containing many
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extracellular matrix proteins and has been shown to have a larger effect on the electrophysiologicalmaturation of hPSC-CMs than laminin, fibronectin or collagen alone.[124] Interestingly, this effect was only present when the cells were cultured on PDMS as opposed to glass, indicating a synergistic role between substrate stiffness and extracellular matrix composition. Likewise, cell surface area only increased with Matrigel when the cells were cultured on PDMS. Three other studies showed significant improvements in maturation of hPSC-CMs by comparing cells cultured on glass or rigid plastic to respectively synthetic co-polymers and a thick layer of Matrigel.[125,126] Because both are soft substrates of undefined stiffness, this makes it challenging to distinguish between the effects of the surface chemistry and substrate stiffness.
Extracellular Matrix Topographical Cues
The adult myocardium is highly anisotropic to allow for effective electrical conduction and force generation. hPSC-CMs can be forced to align by using topographical cues such as micro-grooves [127-130], microcontact printed patterns such as rectangles and lines [121,131], and nanofibers. [132,133] This in turn increases the aspect ratio of the cells to produce more rod-like CMs and impacts other factors of the CM phenotype towards a more mature phenotype. Microgrooves increased the surface area and sarcomere length of hPSC-CMs.[127] Furthermore, the calcium transient kinetics were faster in one study but not in two other studies.[129,133] Besides a decrease in conduction velocity on microgrooved substrates in one report [130], the electrophysiology was consistently unchanged.[128,129,132] Microcontact printing of ECM into 2000 μm2rectangles with a physiologic
7:1 aspect ratio allowed single hPSC-CMs to assume a similar size and shape upon attachment to the substrate. This improved the alignment of sarcomeres within the cells and subsequently the force generation.[121] hPSC-CMs that were induced to form lines using a similar method failed to show polarized Cx43 localization, indicating that this alignment was not sufficient to induce electrophysiological maturation.[131] Aligned nanofiber scaffolds induced an increased in hPSC-CM surface area. Alignment of cells using this approach had no effect on their electrophysiology and sarcomere organization. The data about calcium handling and sarcomere length remains inconclusive.[132,133]
3-Dimensional Constructs
The heart consists of a 3D arrangement of aligned cylindrical CMs, which form myofibers that are adjacent to interstitial fibroblasts, microvasculature, and the ECM. hPSC-CMs in a 2D culture setting cannot fully recapitulate this complex architecture, therefore numerous groups have reported 3D tissue engineering of hPSC-CMs. Where direct comparisons with 2D culture were made, indeed it has become clear that tissue engineering has profound effects on the maturation of hPSC-CMs. Although an early study with an EHT has shown that an unstressed 3D culture does not align hPSC-CMs [119], most tissue engineering approaches result in EHTs with a mature-like degree of anisotropy due to static stress from pillars embedded in the tissue on both ends. Over time, the tissue compacts and aligns itself along the direction between these pillars. When such an EHT was compared with a monolayer of hPSC-CMs, the EHT demonstrated metabolic maturation as indicated by increased use of fatty acids to produce ATP and a mitochondrial proteome similar to the adult heart.[134] Another
study reported on the effects of seeding hPSC-CMs around a surgical suture to form a biowire. Compared to EB culture, the resulting hPSC-CMs were larger and more rod-shaped, proliferated less, displayed less automaticity and had greatly improved electrophysiology overall.[98] When hPSC-CMs were engineered into fibrin-based cardiac tissue patches, their sarcomeres lengthened to be on par with adult heart tissue.[135] Furthermore, this approach doubled the conduction velocity compared to 2D monolayer culture. Fong et al., demonstrated that when hPSC-CMs were cultured in 3D hydrogel made of bovine native ECM, this accelerated the calcium transient kinetics compared to 2D culture on the same material.[136]
Combined Strategies
In vivo cardiac maturation is the result of a complex interplay of many factors. The individual strategies
described above have been very informative for our understanding of the various processes involved in CM maturation. However, to closely simulate the heart’s natural development, various groups have adopted approaches where more than one stimulus for maturation is present. The combination of several stimuli may be necessary to further the maximum maturation status that can be achieved with hPSC-CMs. One example of such a combinatorial approach is the impressively comprehensive effort by Tiburcy et al., who have defined the optimal culture conditions for the generation of relatively mature EHTs. They demonstrated that these optimized culture conditions were applicable to multiple hESC- and hiPSC-lines. The combination of tissue engineering with optimized ECM proteins and optimized media components yielded EHTs with a sarcomeric ultrastructure containing M-bands, thus resembling the adult heart. Importantly, these EHTs demonstrated a positive force-frequency relationship and significant improvements were observed in cell size and sarcomere length.[137] Another study combined two weeks of static stretch of EHTs with one week of electrical stimulation. Compared to static stretch only, the combination with electrical stimulation induced a further strengthening of the tissue with increased force generation, indicating a clear synergistic effect between these two stimuli.[118]
hPSC-CM Maturation In Vivo
In the context of studies into regenerative therapy, a select number of research groups have indeed demonstrated maturation of hPSC-CMs after implantation of these cells in infarcted rodent and primate hearts. The maturation status of transplanted hPSC-CMs has not yet been fully characterized however, owing to the obvious difficulty of retrieving these cells from the animal hearts for analysis. hPSC-CMs that were injected into mouse MI models initially proliferated and then switched to hypertrophic growth after 3 months. After 6 months, the cells exhibited sarcomeric maturity evident by the appearance of M bands. Interestingly, the implanted hPSC-CMs proliferated more than in vitro controls, possibly due to the regenerative state of the mouse heart after myocardial infarction. [138] Similar hypertrophic growth and sarcomeric maturation was observed in a rat MI model.[139] hPSC-CMs also showed gradual maturation after injection in a non-human primate MI model.[3]
6
Framework for the Comparison of hPSC-CMs to Adult
CMs
As reviewed above, many of the papers describing approaches to mature hPSC-CMs have demonstrated significant progress in the maturation state of hPSC-CMs. Using various strategies, researchers have been able to induce improvements in almost all of the known markers of cardiac maturation. Indeed, when many reports on hPSC-CM maturation are combined, a picture emerges displaying that hPSC-CMs can in fact attain adult values for most of the maturation markers (Supplementary Table 3).
However, there is still much ground to be gained. Individual reports in most cases achieve maturation of some maturation markers, but not others. And while significant improvements are often achieved, often the achieved maturation of hPSC-CMs stops short of reaching beyond the fetal or neonatal stage.
This review has the aim to answer two questions related to hPSC-CM maturation: i) “where is the finish line?” and ii) “where are we?”. In this regard we offer here a framework for the semi-quantitative comparison of maturation of hPSC-CMs between studies that allows for the calculation of a multi-factorial Maturation Score.
Figure 1A illustrates the calculation of a Group Maturation Score (GMS), with groups arranged according to the first part of this manuscript: structure, sarcomeres and contractility, electrophysiology and calcium handling, and metabolism. First, based on the reported value for a maturation marker, and the known immature and adult values for this maturation marker in vivo, the Maturation Value (MV) is calculated. This results in a MV between 0 and 1 with 0 being equal to the immature value (fetal or neonatal) in vivo and 1 being equal to the adult value in vivo. Second, if the MV is larger than 1 it is corrected back to a fraction (adjusted MV; MVa) by dividing MV by 1. This is to ensure that values that decrease with maturation, such as time to peak of the calcium transient, are scored correctly. Additionally, it ensures that reported values that have overshot the adult value are also scored correctly. Third, the MVa is converted to a Single Maturation Score (SMS) based on the quintile of the MV
a between 0 and 1. A MVa from 0-0.20 will be termed a poor SMS, a MVa from 0.21-0.40 will be termed a moderate SMS, a MVa from 0.41-0.60 will be termed a fair SMS, a MVa from 0.61-0.80 will be termed a good SMS, and a MV
a from 0.81-1.00 will be termed a very good SMS. Next, the GMS is determined by the average of SMSs for that group. A minimum of two SMSs have to be available to calculate the GMS, except for the Metabolism GMS when the percentage of fatty acids used as metabolic substrate is assessed.
To illustrate the use of this Maturation Score, we have examined the papers published by Tiburcy et al. and Ronaldson-Bouchard et al.[100,137] A visual depiction of the resulting Maturation Scores is presented in Figure 1B. Both articles demonstrate advanced maturation of hPSC-CMs. Structurally, Tiburcy et al. report an aspect ratio of 7.6 and a volume of 12101 μm3, resulting in a
good Group Maturation Score for Structure. Within the group of structural maturation markers as listed in Supplementary Table 1, Ronaldson-Bouchard et al. have only measured the surface area. Therefore, this article did not meet the requirement for having measured at least 2 maturation markers per group to calculate the Group Maturation Score. Within the sarcomeres and contractility
maturation markers, Tiburcy et al. report a sarcomere length of 1.93 μm, a sarcomeric ultrastructure displaying Z-discs, A- and I-bands, H-zones and M-lines, and a maximum force generation of 6.2 mN/ mm2, resulting in a fair Group Maturation Score. Ronaldson-Bouchard et al. report a more mature
sarcomere length of 2.2 μm, a similar sarcomeric ultrastructure with all 5 bands visible on electron microscopy, and a slightly lower maximum force generation of 3.3 mN/mm2, also resulting in a fair
Group Maturation Score. Regarding the Electrophysiology and Calcium Handling group, Tiburcy et al. reported several electrophysiological characteristics of their tissues with varying degrees of maturity. They reported a very mature action potential amplitude but relatively immature maximum upstroke velocity, action potential duration and resting membrane potential, resulting in a moderate Group Maturation Score. Ronaldson-Bouchard et al. reported immature values for the action potential duration and conduction velocity but relatively mature values for the resting membrane potential and gap junction distribution, resulting in a fair Group Maturation Score for Electrophysiology and Calcium Handling. Tiburcy et al. did not assess the metabolic maturity of their engineered heart tissues. Ronaldson-Bouchard et al. reported highly mature metabolic phenotypes for their engineered heart tissues with an adult-like mitochondrial density and adult-like use of fatty acids, resulting in a very good Group Maturation Score for Metabolism.
Figure 1B demonstrates that most commonly maturation is only achieved in part of the Group Maturation Scores. This illustrates that isolated strategies towards the maturation of hPSC-CMs thus far have not been able to achieve advanced maturation of all aspects of hPSC-CM function at the same time. To fully unlock the maturation potential of hPSC-CMs within one system, multiple interventions that induce maturation will likely have to be combined.
Any attempt at comparing a plethora of approaches towards maturation, measured by an equally variable number of assays, is met by some inherent limitations. Measurements on all the maturation markers can be influenced to varying degrees by the experimental setup. For example, differences in cardiac differentiation protocols, the frequency of stimulation, the specific antibody used to score proliferation, the temperature and CO2 levels, can all affect the data obtained from hPSC-CMs and cardiac tissue or CMs isolated from the in vivo setting. Nevertheless, the importance of having a clear definition of cardiac maturation and being able to know how close we are to achieving that goal cannot be understated.
Currently, we are working on the development of a web-based tool that will facilitate the use of the Maturation Score for other researchers.
6
Figure 1 | Setup of the Maturation Score. The calculation of a Maturation Score is shown (A) An example of
the calculation of the Maturation Score and their visual representation is shown for two articles (B).
Overview of Maturation Achieved with hPSC-CMs
Through assessment of the maturation data from various papers describing strategies to mature hPSC-CMs, it becomes clear that hPSC-CMs indeed possess a great potential to mimic adult CMs. Maturation studies on hPSC-CMs have demonstrated that for the majority of maturation markers outlined in Supplementary Table 1, values very close to those of adult CMs can be achieved in hPSC-CMs (Supplementary Table 3).
Integrated Functional Analysis Crucial to Determine
Maturation
A major impediment to assessing the maturation of hPSC-CMs is that commonly used methods for the functional analysis of hPSC-CMs do not allow for a straightforward integrated analysis of all the various markers of maturation.
It is evident that methods that allow for the integrated analysis of hPSC-CMs are crucial to fully analyze the developmental maturity of these cells. Ideally, such methods can analyze multiple parameters of hPSC-CM function simultaneously in a standardized and medium- or high-throughput fashion. Our BASiC method as described in Chapter 2, coupled with microcontacprinted cardiac microconstructs as described in Chapters 3 and 5, offers such a standardized medium-throughput platform allowing for the integrated analysis of hPSC-CMs. The BASiC method can simultaneously quantify contractile kinetics, force generation, electrophysiology and calcium handling of hPSC-CMs. Subsequent immunocytochemistry analysis of the cardiac microconstructs allows for the quantification of several structural parameters such as aspect ratio, volume, binucleation, and sarcomere length. Finally, addition of the SeaHorse assay to this platform as described in Chapter 4 can be used to analyze the metabolic maturation status of cardiac microconstructs. Figure 2 shows a schematic overview of the maturation markers that can be assayed using the BASiC method.
Figure 2 | Schematic Overview of Integrated Analysis of hPSC-CM Function and Maturation Status Using
6
Future Perspectives
Transcriptomic analysis can provide great insight into the extent to which engineered hPSC-CMs resemble their in vivo counterparts on the gene expression level. To this end, some studies into maturation of hPSC-CMs now include transcriptome experiments such as RNA sequencing.[140] The
in vivo expression of genes during cardiac development has also been assayed. Based on this in vivo
data, some excellent tools have been established that allow for comparison with RNA sequencing data from engineered cells.[141] These tools also offer a score to quantify the resemblance between gene expression of the engineered cells and in vivo adult CMs. Herein, we have endeavored to provide a similar framework to assess hPSC-CM maturation based on the structural and functional phenotype rather than the transcriptome. These two methods of assessing hPSC-CM maturation are complementary and a further development of our framework could be the incorporation of the transcriptome maturation score based on the excellent state-of-the-art tools available. The incorporation of genetic expression data will be even more important as transcriptomic analysis becomes more widespread.
References
1. Moretti A, Bellin M, Welling A, Jung CB, Lam JT, Bott-Flügel L, et al. Patient-Specific Induced Pluripotent Stem-Cell Models for Long-QT Syndrome. N Engl J Med. 2010;363: 1397–1409.
2. Lan F, Lee AS, Liang P, Sanchez-Freire V, Nguyen PK, Wang L, et al. Abnormal calcium handling properties underlie familial hypertrophic cardiomyopathy pathology in patient-specific induced pluripotent stem cells. Cell Stem Cell. Elsevier; 2013;12: 101–113.
3. Chong JJH, Yang X, Don CW, Minami E, Liu Y-W, Weyers JJ, et al. Human embryonic-stem-cell-derived cardiomyocytes regenerate non-human primate hearts. Nature. 2014;510: 273–277.
4. Robertson C, Tran DD, George SC. Concise review: maturation phases of human pluripotent stem cell-derived cardiomyocytes. Stem Cells. Wiley Online Library; 2013;31: 829–837.
5. Yang X, Pabon L, Murry CE. Engineering adolescence: maturation of human pluripotent stem cell-derived cardiomyocytes. Circ Res. Am Heart Assoc; 2014;114: 511–523.
6. Aigha I, Raynaud C. Maturation of pluripotent stem cell derived cardiomyocytes: The new challenge. Glob Cardiol Sci Pract. ncbi.nlm.nih.gov; 2016;2016: e201606.
7. Tan SH, Ye L. Maturation of Pluripotent Stem Cell-Derived Cardiomyocytes: a Critical Step for Drug Development and Cell Therapy. J Cardiovasc Transl Res. Springer; 2018; doi:10.1007/s12265-018-9801-5 8. Feric NT, Radisic M. Maturing human pluripotent stem cell-derived cardiomyocytes in human engineered
cardiac tissues. Adv Drug Deliv Rev. Elsevier; 2016;96: 110–134.
9. Scuderi GJ, Butcher J. Naturally Engineered Maturation of Cardiomyocytes. Front Cell Dev Biol. frontiersin. org; 2017;5: 50.
10. Sun X, Nunes SS. Bioengineering Approaches to Mature Human Pluripotent Stem Cell-Derived Cardiomyocytes. Front Cell Dev Biol. ncbi.nlm.nih.gov; 2017;5: 19.
11. Kolanowski TJ, Antos CL, Guan K. Making human cardiomyocytes up to date: Derivation, maturation state and perspectives. Int J Cardiol. internationaljournalofcardiology.com; 2017;241: 379–386.
12. Bedada FB, Wheelwright M, Metzger JM. Maturation status of sarcomere structure and function in human iPSC-derived cardiac myocytes. Biochim Biophys Acta. Elsevier; 2016;1863: 1829–1838.
13. Besser RR, Ishahak M, Mayo V, Carbonero D, Claure I, Agarwal A. Engineered Microenvironments for Maturation of Stem Cell Derived Cardiac Myocytes. Theranostics. ncbi.nlm.nih.gov; 2018;8: 124–140.
14. Keung W, Boheler KR, Li RA. Developmental cues for the maturation of metabolic, electrophysiological and calcium handling properties of human pluripotent stem cell-derived cardiomyocytes. Stem Cell Res Ther. stemcellres.biomedcentral.com; 2014;5: 17.
15. Atmanli A, Domian IJ. Recreating the Cardiac Microenvironment in Pluripotent Stem Cell Models of Human Physiology and Disease. Trends Cell Biol. 2017;27: 352–364.
16. Carvalho AB, de Carvalho ACC. Heart regeneration: Past, present and future. World J Cardiol. 2010;2: 107–111. 17. Mollova M, Bersell K, Walsh S, Savla J, Das LT, Park S-Y, et al. Cardiomyocyte proliferation contributes to heart
growth in young humans. Proc Natl Acad Sci U S A. 2013;110: 1446–1451.
18. Bergmann O, Zdunek S, Felker A, Salehpour M, Alkass K, Bernard S, et al. Dynamics of Cell Generation and Turnover in the Human Heart. Cell. 2015;161: 1566–1575.
19. Gerdes AM, Kellerman SE, Moore JA, Muffly KE, Clark LC, Reaves PY, et al. Structural remodelling of cardiac myocytes in patients with ischemic cardiomyopathy. Circulation. 1992;86: 426–430.
20. Mummery C, Ward-van Oostwaard D, Doevendans P, Spijker R, van den Brink S, Hassink R, et al. Differentiation of human embryonic stem cells to cardiomyocytes: role of coculture with visceral endoderm-like cells. Circulation. 2003;107: 2733–2740.
21. Anversa P, Olivetti G, Loud AV. Morphometric study of early postnatal development in the left and right ventricular myocardium of the rat. I. Hypertrophy, hyperplasia, and binucleation of myocytes. Circ Res. 1980;46: 495–502.
22. Adler CP. [DNA in growing hearts of children. Biochemical and cytophotometric investigations (author’s transl)]. Beitr Pathol. 1976;158: 173–202.
6
23. Brodsky VY, Ya. Brodsky V, Chernyaev AL, Vasilyeva IA. Variability of the cardiomyocyte ploidy in normal human hearts. Virchows Arch B Cell Pathol Incl Mol Pathol. 1992;61: 289–294.
24. Adler CP, Costabel U. Cell number in human heart in atrophy, hypertrophy, and under the influence of cytostatics. Recent Adv Stud Cardiac Struct Metab. 1975;6: 343–355.
25. Olivetti G, Cigola E, Maestri R, Corradi D, Lagrasta C, Gambert SR, et al. Aging, cardiac hypertrophy and ischemic cardiomyopathy do not affect the proportion of mononucleated and multinucleated myocytes in the human heart. J Mol Cell Cardiol. 1996;28: 1463–1477.
26. Ribeiro MC, Tertoolen LG, Guadix JA, Bellin M, Kosmidis G, D’Aniello C, et al. Functional maturation of human pluripotent stem cell derived cardiomyocytes in vitro – Correlation between contraction force and electrophysiology. Biomaterials. 2015;51: 138–150.
27. Rubin R, Strayer DS, Rubin E. Rubin’s Pathology: Clinicopathologic Foundations of Medicine. Lippincott Williams & Wilkins; 2011.
28. Kim HD, Kim DJ, Lee IJ, Rah BJ, Sawa Y, Schaper J. Human fetal heart development after mid-term: morphometry and ultrastructural study. J Mol Cell Cardiol. 1992;24: 949–965.
29. Lahmers S, Wu Y, Call DR, Labeit S, Granzier H. Developmental control of titin isoform expression and passive stiffness in fetal and neonatal myocardium. Circ Res. 2004;94: 505–513.
30. Opitz CA, Leake MC, Makarenko I, Benes V, Linke WA. Developmentally regulated switching of titin size alters myofibrillar stiffness in the perinatal heart. Circ Res. 2004;94: 967–975.
31. Warren CM, Krzesinski PR, Campbell KS, Moss RL, Greaser ML. Titin isoform changes in rat myocardium during development. Mech Dev. 2004;121: 1301–1312.
32. Reiser PJ, Portman MA, Ning XH, Schomisch Moravec C. Human cardiac myosin heavy chain isoforms in fetal and failing adult atria and ventricles. Am J Physiol Heart Circ Physiol. 2001;280: H1814–20.
33. Miyata S, Minobe W, Bristow MR, Leinwand LA. Myosin heavy chain isoform expression in the failing and nonfailing human heart. Circ Res. 2000;86: 386–390.
34. Bedada FB, Chan SS-K, Metzger SK, Zhang L, Zhang J, Garry DJ, et al. Acquisition of a quantitative, stoichiometrically conserved ratiometric marker of maturation status in stem cell-derived cardiac myocytes. Stem Cell Reports. 2014;3: 594–605.
35. Sasse S, Brand NJ, Kyprianou P, Dhoot GK, Wade R, Arai M, et al. Troponin I gene expression during human cardiac development and in end- stage heart failure. Circ Res. 1993;72: 932–938.
36. Hunkeler NM, Kullman J, Murphy AM. Troponin I isoform expression in human heart. Circ Res. 1991;69: 1409–1414.
37. Metzger JM, Michele DE, Rust EM, Borton AR, Westfall MV. Sarcomere thin filament regulatory isoforms. Evidence of a dominant effect of slow skeletal troponin I on cardiac contraction. J Biol Chem. 2003;278: 13118–13123.
38. Siedner S, Krüger M, Schroeter M, Metzler D, Roell W, Fleischmann BK, et al. Developmental changes in contractility and sarcomeric proteins from the early embryonic to the adult stage in the mouse heart. J Physiol. 2003;548: 493–505.
39. Thompson BR, Houang EM, Sham YY, Metzger JM. Molecular determinants of cardiac myocyte performance as conferred by isoform-specific TnI residues. Biophys J. 2014;106: 2105–2114.
40. Westfall MV, Rust EM, Metzger JM. Slow skeletal troponin I gene transfer, expression, and myofilament incorporation enhances adult cardiac myocyte contractile function. Proc Natl Acad Sci U S A. 1997;94: 5444–5449.
41. Chuva de Sousa Lopes SM, Hassink RJ, Feijen A, van Rooijen MA, Doevendans PA, Tertoolen L, et al. Patterning the heart, a template for human cardiomyocyte development. Dev Dyn. 2006;235: 1994–2002.
42. Holubarsch C, Lüdemann J, Wiessner S, Ruf T, Schulte-Baukloh H, Schmidt-Schweda S, et al. Shortening versus isometric contractions in isolated human failing and non-failing left ventricular myocardium: dependency of external work and force on muscle length, heart rate and inotropic stimulation. Cardiovasc Res. 1998;37: 46–57.
43. Mulieri LA, Hasenfuss G, Leavitt B, Allen PD, Alpert NR. Altered myocardial force-frequency relation in human heart failure. Circulation. 1992;85: 1743–1750.
44. Hasenfuss G, Mulieri LA, Blanchard EM, Holubarsch C, Leavitt BJ, Ittleman F, et al. Energetics of isometric force development in control and volume-overload human myocardium. Comparison with animal species. Circ Res. 1991;68: 836–846.
45. Racca AW, Klaiman JM, Pioner JM, Cheng Y, Beck AE, Moussavi-Harami F, et al. Contractile properties of developing human fetal cardiac muscle. J Physiol. 2016;594: 437–452.
46. Wiegerinck RF, Cojoc A, Zeidenweber CM, Ding G, Shen M, Joyner RW, et al. Force frequency relationship of the human ventricle increases during early postnatal development. Pediatr Res. 2009;65: 414–419.
47. Belus A, Piroddi N, Scellini B, Tesi C, D’Amati G, Girolami F, et al. The familial hypertrophic cardiomyopathy-associated myosin mutation R403Q accelerates tension generation and relaxation of human cardiac myofibrils. J Physiol. 2008;586: 3639–3644.
48. Piroddi N, Belus A, Scellini B, Tesi C, Giunti G, Cerbai E, et al. Tension generation and relaxation in single myofibrils from human atrial and ventricular myocardium. Pflugers Arch. 2007;454: 63–73.
49. Bers DM. Cardiac excitation–contraction coupling. Nature. 2002;415: 198–205.
50. Pieske B, Trost S, Schütt K, Minami K, Just H, Hasenfuss G. Influence of Forskolin on the force-frequency behavior in nonfailing and end-stage failing human myocardium. Basic Res Cardiol. 1998;93: s066–s075. 51. Hasenfuss G, Schillinger W, Lehnart SE, Preuss M, Pieske B, Maier LS, et al. Relationship Between Na -Ca2
Exchanger Protein Levels and Diastolic Function of Failing Human Myocardium. Circulation. 1999;99: 641–648. 52. Ursem NT, Struijk PC, Hop WC, Clark EB, Keller BB, Wladimiroff JW. Heart rate and flow velocity variability as
determined from umbilical Doppler velocimetry at 10-20 weeks of gestation. Clin Sci . 1998;95: 539–545. 53. Koncz I, Szél T, Bitay M, Cerbai E, Jaeger K, Fülöp F, et al. Electrophysiological effects of ivabradine in dog and
human cardiac preparations: potential antiarrhythmic actions. Eur J Pharmacol. 2011;668: 419–426.
54. Drouin E, Charpentier F, Gauthier C, Laurent K, Le Marec H. Electrophysiologic characteristics of cells spanning the left ventricular wall of human heart: evidence for presence of M cells. J Am Coll Cardiol. 1995;26: 185–192. 55. Gennser G, Nilsson E. Excitation and impulse conduction in the human fetal heart. Acta Physiol Scand.
1970;79: 305–320.
56. Chen SC, Davis LM, Westphale EM, Beyer EC, Saffitz JE. Expression of multiple gap junction proteins in human fetal and infant hearts. Pediatr Res. 1994;36: 561–566.
57. Vreeker A, van Stuijvenberg L, Hund TJ, Mohler PJ, Nikkels PGJ, van Veen TAB. Assembly of the Cardiac Intercalated Disk during Pre- and Postnatal Development of the Human Heart. PLoS One. 2014;9: e94722. 58. Nanthakumar K, Jalife J, Massé S, Downar E, Pop M, Asta J, 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. 2007;293: H875–80.
59. Taggart P, Sutton PM, Opthof T, Coronel R, Trimlett R, Pugsley W, et al. Inhomogeneous transmural conduction during early ischaemia in patients with coronary artery disease. J Mol Cell Cardiol. 2000;32: 621–630. 60. Ziman AP, Gómez-Viquez NL, Bloch RJ, Lederer WJ. Excitation-contraction coupling changes during postnatal
cardiac development. J Mol Cell Cardiol. 2010;48: 379–386.
61. Høydal MA, Kirkeby-Garstad I, Karevold A, Wiseth R, Haaverstad R, Wahba A, et al. Human cardiomyocyte calcium handling and transverse tubules in mid-stage of post-myocardial-infarction heart failure. ESC Heart Fail. 2018; doi:10.1002/ehf2.12271
62. Parikh SS, Blackwell DJ, Gomez-Hurtado N, Frisk M, Wang L, Kim K, et al. Thyroid and Glucocorticoid Hormones Promote Functional T-Tubule Development in Human-Induced Pluripotent Stem Cell-Derived Cardiomyocytes. Circ Res. 2017;121: 1323–1330.
63. Risi C, Eisner J, Belknap B, Heeley DH, White HD, Schröder GF, et al. Ca-induced movement of tropomyosin on native cardiac thin filaments revealed by cryoelectron microscopy. Proc Natl Acad Sci U S A. 2017;114: 6782–6787.
64. Piacentino V 3rd, Weber CR, Chen X, Weisser-Thomas J, Margulies KB, Bers DM, et al. Cellular basis of abnormal calcium transients of failing human ventricular myocytes. Circ Res. 2003;92: 651–658.
65. Vahl CF, Bonz A, Timek T, Hagl S. Intracellular calcium transient of working human myocardium of seven patients transplanted for congestive heart failure. Circ Res. 1994;74: 952–958.
66. Beuckelmann DJ, Näbauer M, Erdmann E. Intracellular calcium handling in isolated ventricular myocytes from patients with terminal heart failure. Circulation. 1992;85: 1046–1055.
6
67. Gwathmey JK, Slawsky MT, Hajjar RJ, Briggs GM, Morgan JP. Role of intracellular calcium handling in force-interval relationships of human ventricular myocardium. J Clin Invest. 1990;85: 1599–1613.
68. Pieske B, Kretschmann B, Meyer M, Holubarsch C, Weirich J, Posival H, et al. Alterations in intracellular calcium handling associated with the inverse force-frequency relation in human dilated cardiomyopathy. Circulation. 1995;92: 1169–1178.
69. Liu J, Fu JD, Siu CW, Li RA. Functional sarcoplasmic reticulum for calcium handling of human embryonic stem cell-derived cardiomyocytes: insights for driven maturation. Stem Cells. 2007;25: 3038–3044.
70. Fu J-D, Jiang P, Rushing S, Liu J, Chiamvimonvat N, Li RA. Na+/Ca2+ exchanger is a determinant of excitation-contraction coupling in human embryonic stem cell-derived ventricular cardiomyocytes. Stem Cells Dev. 2010;19: 773–782.
71. Louch WE, Koivumäki JT, Tavi P. Calcium signalling in developing cardiomyocytes: implications for model systems and disease. J Physiol. 2015;593: 1047–1063.
72. Kolwicz SC Jr, Purohit S, Tian R. Cardiac metabolism and its interactions with contraction, growth, and survival of cardiomyocytes. Circ Res. 2013;113: 603–616.
73. Barth E, Stämmler G, Speiser B, Schaper J. Ultrastructural quantitation of mitochondria and myofilaments in cardiac muscle from 10 different animal species including man. J Mol Cell Cardiol. 1992;24: 669–681.
74. Schaper J, Meiser E, Stämmler G. Ultrastructural morphometric analysis of myocardium from dogs, rats, hamsters, mice, and from human hearts. Circ Res. 1985;56: 377–391.
75. Marin-Garcia J, Ananthakrishnan R, Goldenthal MJ. Heart mitochondrial DNA and enzyme changes during early human development. Mol Cell Biochem. 2000;210: 47–52.
76. Hattori F, Chen H, Yamashita H, Tohyama S, Satoh Y-S, Yuasa S, et al. Nongenetic method for purifying stem cell-derived cardiomyocytes. Nat Methods. 2010;7: 61–66.
77. Bing RJ, Siegel A, Ungar I, Gilbert M. Metabolism of the human heart. II. Studies on fat, ketone and amino acid metabolism. Am J Med. 1954;16: 504–515.
78. Wisneski JA, Gertz EW, Neese RA, Mayr M. Myocardial metabolism of free fatty acids. Studies with 14C-labeled substrates in humans. J Clin Invest. 1987;79: 359–366.
79. Wisneski JA, Gertz EW, Neese RA, Gruenke LD, Morris DL, Craig JC. Metabolic fate of extracted glucose in normal human myocardium. J Clin Invest. 1985;76: 1819–1827.
80. Miller HI, Yum KY, Durham BC. Myocardial free fatty acid in unanesthetized dogs at rest and during exercise. Am J Physiol. 1971;220: 589–596.
81. Shipp JC, Opie LH, Challoner D. Fatty Acid and Glucose Metabolism in the Perfused Heart. Nature. 1961;189: 1018–1019.
82. Kaijser L. Effect of metabolic intervention on substrate metabolism in the human heart. Adv Myocardiol. 1980;2: 51–59.
83. Cox SJ, Gunberg DL. Energy metabolism in isolated rat embryo hearts: effect of metabolic inhibitors. J Embryol Exp Morphol. 1972;28: 591–599.
84. Werner JC, Sicard RE. Lactate metabolism of isolated, perfused fetal, and newborn pig hearts. Pediatr Res. 1987;22: 552–556.
85. Medina JM. The role of lactate as an energy substrate for the brain during the early neonatal period. Biol Neonate. 1985;48: 237–244.
86. Itoi T, Lopaschuk GD. The contribution of glycolysis, glucose oxidation, lactate oxidation, and fatty acid oxidation to ATP production in isolated biventricular working hearts from 2-week-old rabbits. Pediatr Res. 1993;34: 735–741.
87. Lopaschuk GD, Spafford MA, Marsh DR. Glycolysis is predominant source of myocardial ATP production immediately after birth. Am J Physiol. 1991;261: H1698–705.
88. Lopaschuk GD, Spafford MA. Energy substrate utilization by isolated working hearts from newborn rabbits. Am J Physiol. 1990;258: H1274–80.
89. Snir M, Kehat I, Gepstein A, Coleman R, Itskovitz-Eldor J, Livne E, et al. Assessment of the ultrastructural and proliferative properties of human embryonic stem cell-derived cardiomyocytes. Am J Physiol Heart Circ Physiol. 2003;285: H2355–63.
