- Open Access
Modeling hypertrophic cardiomyopathy with human cardiomyocytes derived from induced pluripotent stem cells
Stem Cell Research & Therapy volume 13, Article number: 232 (2022)
One of the obstacles in studying the pathogenesis of hypertrophic cardiomyopathy (HCM) is the poor availability of myocardial tissue samples at the early stages of disease development. This has been addressed by the advent of induced pluripotent stem cells (iPSCs), which allow us to differentiate patient-derived iPSCs into cardiomyocytes (iPSC-CMs) in vitro. In this review, we summarize different approaches to establishing iPSC models and the application of genome editing techniques in iPSC. Because iPSC-CMs cultured at the present stage are immature in structure and function, researchers have attempted several methods to mature iPSC-CMs, such as prolonged culture duration, and mechanical and electrical stimulation. Currently, many researchers have established iPSC-CM models of HCM and employed diverse methods for performing measurements of cellular morphology, contractility, electrophysiological property, calcium handling, mitochondrial function, and metabolism. Here, we review published results in humans to date within the growing field of iPSC-CM models of HCM. Although there is no unified consensus, preliminary results suggest that this approach to modeling disease would provide important insights into our understanding of HCM pathogenesis and facilitate drug development and safety testing.
Hypertrophic cardiomyopathy (HCM) is a fatal heterogeneous myocardial disease, caused by autosomal dominant sarcomeric gene mutations, which manifests as left ventricular hypertrophy, myocardial hypercontractility, diastolic dysfunction, myofibrillar disarray, and fibrosis [1,2,3]. Epidemiological studies based on echocardiography have shown a prevalence of 1/500 in the population [1, 4], but a higher prevalence (about 1/200) when clinical and genetic diagnoses (including family members) are taken into account [5, 6]. Pare et al.  reported that mutation at the protein level was found in the MYH7 gene, encoding the β-myosin heavy chain (β-MHC). Based on this study, Seidman’s team identified missense mutations in the MYH7 gene that were associated with the first HCM . Subsequently, numerous genetic studies have shown that HCM is an inherited disease of the cardiac sarcomere. Furthermore, the advent of next-generation sequencing and whole-exome sequencing led to the discovery of new HCM mutations in sarcomeric and sarcomere-associated genes [9, 10], reconfirming that HCM is primarily a monogenic sarcomeric disease.
Approximately half of HCM patients harbor mutations in genes that encode sarcomeric proteins and related myofilament elements responsible for regulating cardiomyocyte contraction and cardiac function [11,12,13]. Among the known causal genes, MYH7 and myosin-binding protein C (MYBPC3) are the most common, and they are responsible for about half of familial HCM patients [11, 14, 15]. Mutations of TNNT2, TNNI3, and TPM1 are relatively uncommon, together accounting for less than 10% [16,17,18]. Although less common, cardiac α-actin (ACTC1), myosin light chain 2 (MYL2), myosin light chain 3 (MYL3), and cysteine- and glycine-rich protein in 3 (CSRP3) have also been identified as causes of HCM [19,20,21]. The nine genes mentioned above have the strongest causal role in HCM (Table 1) [9, 22].
For common HCM mutations, such as MYH7 and MYBPC3, their causal role is unambiguous. However, not all HCM mutations cause HCM . Even in the same family, HCM mutations typically show different expressivity (defined as the severity of the phenotype that develops in patients with the pathogenic mutation) and penetrance (defined as the proportion of individuals carrying a pathogenic mutation who display a phenotype) . Because of human genetic diversity, population-specific frequency of variants, and the presence of thousands of coding variants in each exome, it is difficult to distinguish whether causal or incidental variants cause HCM [23, 25,26,27,28].
Although we have identified many mutations that cause HCM, our understanding of the pathogenesis from mutation to phenotype remains incomplete. The reason for this lies not only in the diversity of mutations that lead to similar clinical manifestations but also in the fact that animal models only partially recapitulate human phenotypes. For example, in mouse models, heterozygous mutations in MYH7 or MYBPC3 failed to develop pathognomonic septal hypertrophy seen in patients . Neither has left ventricular obstruction been observed in any mouse models. Either homozygous knock-in or knock-out of the respective gene is lethal (e.g., α-MHC ), or animals develop severe left ventricular dysfunction (e.g., MYBPC3 [31,32,33,34,35,36,37,38]). The mouse reports still did not answer some fundamental questions: (1) the exact physiological roles of the most common sarcomeric proteins in HCM such as β-MHC and cMyBPC; (2) the mechanisms by which these sarcomeric proteins ensure normal cardiac systolic and diastolic function; and (3) the mechanisms of myocardial hypertrophy and disarray in HCM due to mutations in sarcomeric and non-sarcomeric genes with a different function [15, 29, 39, 40].
In addition, another reason for our limited understanding of the pathophysiology of HCM is that few myocardial tissues and cells have been isolated from HCM patients. And only a very small number of studies have specifically reported on the in vitro phenotype of HCM . The myocardial tissues are commonly obtained from HCM patients undergoing surgical septal myectomy or heart transplantation. However, all these tissue sources represent an advanced stage of HCM, making us wonder to which extent the abnormalities in comparison with the heart tissue without heart failure can reflect primary defects or secondary compensatory outcomes.
The somatic cell reprogramming technique discovered by Yamanaka  paved the way for the generation of cardiomyocytes derived from patient-specific induced pluripotent stem cells (iPSCs), which allowed us to deeply understand the pathogenesis of HCM. The primary cause of HCM is the mutation in sarcomeric genes, leading to changes in Ca2+ handling properties, ion channel remodeling, energy deficiency, and microvascular dysfunction. However, the mechanisms why progressive changes initiated by these primary mutations occur in one individual and not in others are still unknown. Therefore, iPSC models represent a valuable tool to study HCM in vitro. By improving protocols for the generation of iPSC lines and the differentiation of cardiomyocytes from iPSCs, researchers can recapitulate in vitro phenotypes of HCM. iPSC-derived cardiomyocytes (iPSC-CMs) have several advantages: (1) They can be generated from a wide variety of readily available cells, including those in the skin, urine, and blood [43,44,45]; (2) they resemble the early stage of cardiac development and have the ability to show morphological and functional changes that first appear in disease without being hidden by systemic compensatory responses ; and (3) they are patient-specific, enabling genotype–phenotype association and providing an unprecedented platform for the drug screening for individualized therapy.
Up to date, much progress has been made utilizing patient-specific iPSC-CM models to characterize HCM and study the pathogenesis of HCM. In this review, we will provide an in-depth overview of current iPSC-CM models of HCM, including the generation and functional parameters of iPSC-CMs, methods of maturing iPSC-CMs, gene editing in iPSCs, and the HCM phenotype of iPSC-CMs. Finally, we will discuss the future perspectives of iPSC-CM models of HCM.
Generation of iPSC-CM models
As a new approach to mimicking human diseases and associated genetic mutations, iPSC technology became possible since Yamanaka identified four critical reprogramming factors (OCT4, SOX2, KLF4, c-Myc). Depending on the type of donor cells, the combination of reprogramming factors might vary with the fact that specific cell types may endogenously express some factors. For example, c-Myc is not necessary for the reprogramming of fibroblasts . Patient-specific iPSC-CM models require obtaining human tissues to generate iPSC lines. To minimize the invasiveness, a gradual shift from using fibroblasts from skin biopsies to urine or blood cells can be observed [44, 45].
Initially, iPSCs were generated by retroviral transduction [42, 48, 49]. Yamanaka's laboratory used the Moloney-based retroviral vector system, which generated iPSCs with high efficiency, but was restricted to dividing cells. Therefore, lentiviruses were used to improve the transduction efficiency of both dividing and non-dividing cells. However, it was found that the expression of Yamanaka factors was difficult to silence after lentiviral transduction [50, 51], resulting in difficulty in the differentiation of iPSCs . Therefore, inducible systems were used to silence reprogramming factors so that they were not expressed . However, the integration of viruses into host cell DNA increases the risk of insertional mutations . For example, transgenic reactivation of c-Myc showed increased tumorigenicity limiting their application .
To overcome the disadvantages of retroviruses and lentiviruses, non-integrating vectors have been developed, including adenovirus, Sendai virus, virus-free methods like episomal transfection, and synthetic mRNA delivery. In host cells, adenovirus transduction allows the overexpression of reprogramming factors without genomic integration . Sendai virus  is an RNA virus that does not enter the nucleus, thus reducing the risk of genomic insertion. Episomal transfection  is an alternative method to generate virus-free iPSCs. However, the efficiency of this approach is relatively low . In addition, transfection using synthetic mRNA  is a simple, non-integrating strategy with high efficiency, which can overcome innate antiviral responses. Non-integrating vectors generate iPSCs that are more suitable for disease modeling.
In addition to integrating and non-integrating vectors, there are transgene-free reprogramming methods that can also generate iPSCs by implanting recombinant reprogramming factors into somatic cells . Studies have shown that the reprogramming efficiency was improved by using small molecule compounds, e.g., the histone deacetylase inhibitor valproic acid .
In summary, different integration vectors, non-integration vectors, and non-transgenic reprogramming methods have been developed to improve efficiency and further reduce the risk of genome alteration.
The adult heart is a post-mitotic organ with a very limited regeneration capacity . However, in most cases, it is difficult to isolate primary human cardiomyocytes from surgical specimens and culture them for a long time due to ethical and technical issues. In addition, cardiomyocytes isolated from animals have species differences, e.g., different electrophysiology in comparison with human ones. Therefore, iPSCs are an important source of cardiomyocytes .
The differentiation of cardiomyocytes from human iPSCs can be achieved under in vitro culture conditions by modulating the signaling pathways involved in cardiac development . Currently, there have been three main strategies for the directed differentiation of cardiomyocytes from iPSCs: co-culture with visceral endoderm-like (END-2) cells , embryoid body (EB)-based differentiation , and monolayer cell culture .
During embryonic development, END-2 cells release the factors that lead to cardiac differentiation around the mesoderm . This finding was the basis of the co-culture strategy in which iPSCs can differentiate into cardiomyocytes with the presence of END-2 cells. Although this culture protocol successfully directed the differentiation of iPSCs into cardiomyocytes, the yield was very low (less than 10%) . EB-based differentiation is a serum-mediated three-dimensional (3D) culture approach that relies on the ability of iPSCs to form floating cell aggregates in the low-adherent matrix. These aggregates called EBs can differentiate into cells of all three germ layers. However, due to the presence of serum, this method has low reproducibility and large inter-group differences . Therefore, serum was then replaced by cytokines and growth factors, such as Wnt proteins , bone morphogenetic proteins (BMPs), and activin A [71, 72]. In addition, some small molecules could promote cardiac differentiation, including activators (CHIR99021) and inhibitors (IWR, XAV, IWP2) of the Wnt pathway . However, by using this approach, the number of cells required is large and the differentiation efficiency is low. To overcome these disadvantages, the protocol of monolayer cell culture was developed [74, 75]. This approach can significantly increase the differentiation efficiency of iPSCs with derived cells exhibiting characteristic phenotypes of ventricular, atrial, or junctional cardiomyocytes . Although the details, efficiency, and yield vary widely in various studies, the monolayer cell culture protocol has been used by an increasing number of investigators, probably because of its simplicity and high efficiency [77, 78].
iPSCs and genome editing
iPSC technology has unprecedented advantages. However, using healthy relative- or healthy unrelated donor-derived iPSC-CMs as standard controls in early studies was not adequate . This is because the phenotype differences between patient-derived iPSC-CMs and controls may be the result of different (epi)genetic backgrounds rather than disease-specific variants, or the effect of (epi)genetic backgrounds on phenotype may outweigh that caused by mutations. For example, the action potential duration of different iPSC-CM lines varies largely in healthy controls . Furthermore, it has been reported that the monozygotic twins carrying the same MYH7 mutation differed significantly in the degree of myocardial fibrosis . Therefore, it is sometimes difficult to link direct effects on the function with specific mutations by using healthy controls, as the (epi)genetic background of these cells is largely unknown .
Genome editing techniques like CRISPR-Cas9 , CPF1 , and TALENs  enable the generation of isogenic cell lines, which differ only in the mutation being studied and retain the same (epi)genetic background. Thus, the effects of mutations can be directly compared with their isogenic wild-type controls. Among all genome editing techniques, CRISPR-Cas9 is the first choice due to its low cost, simple structure, and high fidelity. Gene editing techniques can make precise changes to the genome, ranging from insertions or deletions (such as the ~ 65 kb Dip2a gene ) to the point mutation of a single base . By combining iPSCs with genome editing techniques, it is possible to directly compare iPSC-CMs carrying disease-associated mutations with their corresponding isogenic wild-type controls to determine the exact effect of the mutation on disease .
Gene editing technologies are based on endonuclease activity, which aims to insert double-strand breaks (DSBs) into the genome at a precise and desired site. There are two main mechanisms for the repair of DSBs: nonhomologous end joining (NHEJ) and homology-directed repair (HDR) . NHEJ is an efficient but error-prone process that may cause insertion, deletion, or substitution of nucleotides . Insertion/deletion frequently results in frameshift mutations, which lead to premature termination codons (PTCs) . HDR is a more accurate method for the specific repair of DSBs. Based on the use of a homologous template (single-stranded oligonucleotides or double-stranded DNA templates), HDR allows the precise introduction of point mutation , micro-peptide-encoding tags , and even fluorescent proteins at specific positions [92,93,94]. Thus, HDR can be targeted either to introduce specific mutations in healthy iPSCs or to correct preexisting genetic mutations to generate isogenic lines. However, the ratio of HDR is lower compared to NHEJ, limiting the efficiency of gene knock-in or introducing specific mutations . Therefore, different approaches have been developed to inhibit NHEJ or promote HDR [96, 97]. For example, covalent linkage of DNA repair template to CRISPR-Cas9 nuclease can improve HDR efficiency, which may be a promising strategy [98, 99]. In addition, an alternative is using base editors that replace bases in the target DNA without breaking double-stranded DNA and DNA repair templates .
It is indispensable that iPSC-CMs recapitulate the phenotypes of adult cardiomyocytes for disease modeling. However, iPSC-CMs generated using standard protocols differ from adult cardiomyocytes in cellular morphology, sarcomeric protein isoforms, electrophysiology, excitation–contraction coupling, and calcium handling, as well as mitochondrial function and metabolic characteristics. Table 2 summarizes the major phenotypic differences between iPSC-CMs and adult cardiomyocytes.
The iPSC-CMs generated using current protocols are usually round or polygonal with disorganized sarcomeres, whereas adult cardiomyocytes are rod-shaped with an aspect ratio of 5:1–9:1 and organized sarcomeres [101, 102]. iPSC-CMs had lower surface area and volume compared to adult cardiomyocytes . Differences in organelle distribution and morphology between iPSC-CMs and adult cardiomyocytes are also evident. iPSC-CMs in comparison with adult cardiomyocytes lack developed t-tubules networks, which are necessary for efficient contractile function . Furthermore, iPSC-CMs are mostly mononucleated, whereas adult cardiomyocytes are usually multinucleated .
Sarcomeric protein isoforms
During cardiac development, many sarcomeric proteins undergo isoform switching, which can be used as useful markers to determine the maturity level of cardiomyocytes. For example, a switch of the cardiac troponin I (cTnI) isoforms occurs during the maturation of cardiomyocytes. In the fetal heart, expression levels of slow skeletal troponin I (ssTnI) are higher than that of cTnI, whereas cTnI expression levels in adult cardiomyocytes are elevated and ssTnI levels are decreased . Bedada et al.  reported that iPSC-CMs mainly expressed ssTnI, indicating their relative immaturity. In addition, myosin heavy chain (MHC) also undergoes isoforms switching during maturation. In human hearts, β-MHC (encoded by the MYH7 gene) dominates throughout the development of cardiomyocytes, and levels of which increase with age . Compared with adult cardiomyocytes, the content of α-MHC (encoded by the MYH6 gene) in iPSC-CMs is higher. However, α-MHC slowly converts to β-MHC as iPSC-CMs mature. Finally, fetal cardiomyocytes predominantly express the N2BA isoform of titin, whereas adult cardiomyocytes mainly express the N2B isoform . It was reported that, similar to fetal cardiomyocytes, iPSC-CMs primarily express the N2BA isoform . Therefore, we can judge whether iPSC-CMs are mature based on the ratios of cTnI/ssTnI, α-MHC/β-MHC, and N2B/N2BA.
Directly comparing differences in electrophysiological properties between iPSC-CMs and adult cardiomyocytes is challenging due to experimental differences, tissue heterogeneity, and disease states. However, the detailed electrophysiological characterization of iPSC-CMs has been reported [111, 112]. Prior studies have documented that iPSC-CMs in comparison with adult cardiomyocytes displayed action potential (AP) phenotypes characterized by smaller maximum diastolic potential and slower upstroke velocity [76, 111]. In addition, it has been reported that the ventricular-like iPSC-CMs shared electrophysiological properties analogous to those of adult cardiomyocytes, including the distinct plateau phase (phase 2) followed by accelerated repolarization (phase 3), the AP duration in the normal range of QT interval, and the maximum diastolic potential which was close to that of adult cardiomyocytes .
Different ionic currents have been characterized in iPSC-CMs, including sodium (INa), calcium (ICa), hyperpolarization-activated pacemaker (If), transient outward potassium (Ito), inward rectifier potassium (IK1), and the rapid and slow activating components of the delayed rectifier potassium currents (IKr and IKs, respectively). iPSC-CMs have prominent INa and ICa with activation and inactivation gating properties that are similar to those of human ventricular cardiomyocytes [111, 113, 114]. Ma et al.  reported the presence of If in ventricular-like iPSC-CMs, which promoted auto-depolarization in phase 4. Three K+ currents (Ito, IKr, and IKs) were observed in iPSC-CMs with their maximum current densities and activation characteristics comparable with those of adults CMs [55, 111]. However, IK1 in iPSC-CMs was either undetectable or significantly smaller than that in adult cardiomyocytes.
In summary, multiple ion channels are present in both iPSC-CMs and adult cardiomyocytes, which lead to characteristic AP. But significant differences do exist, such as reduced IK1 and the existence of If. Consequently, the iPSC-CMs exhibit spontaneous automaticity, which is not recorded in adult cardiomyocytes.
Excitation contraction coupling and calcium handling
Cardiac contraction and relaxation are accomplished through excitation–contraction coupling, which is the orchestrated cycling of calcium between cytoplasm, sarcoplasmic reticulum (SR), and troponin . The observed whole-cell intracellular Ca2+ concentration ([Ca2+]i) transients in iPSC-CMs demonstrated the presence of excitation–contraction coupling resembling native cardiomyocytes . It has been reported that Ca2+ influx into cells through depolarization-activated L-type Ca2+ channels leads to the release of SR Ca2+ stores via Ca2+-sensitive ryanodine receptors, recapitulating a process known as Ca2+-induced Ca2+ release in iPSC-CMs . It was noted that the Ca2+ cycling properties of iPSC-CMs cultured on micro-grooved substrates were significantly improved with a shorter time to peak and more organized SR Ca2+ release in response to caffeine . Besides, iPSC-CM-based HCM modeling indicates the existence of functional SR and Ca2+ transients . However, Ca2+ transients in iPSC-CMs are small with a relatively slow rise characterized by a U-shape waveform, suggesting that the calcium handling properties of iPSC-CMs are relatively immature . Several studies have shown that the Ca2+ handling function in iPSC-CMs may be affected by poorly developed SR and T-tubules deficiency [104, 120].
Mitochondrial function and metabolism
The heart contains a large number of mitochondria, comprising up to 23% in human, 22% in dog, 28% in rat, and 32% in mouse myocardium . The mitochondria of adult cardiomyocytes are mostly rod-shaped and evenly arranged along the sarcomere . Cellular factors controlling mitochondria are highly expressed in adult cardiomyocytes, including fission factor Drp1 and fusion factors Mfn1, Mfn2, and Opa1 . Compared with adult cardiomyocytes, iPSC-CMs have fewer and thinner mitochondria, which usually aggregate near the nucleus with decreased expression of mitochondrial dynamics proteins [120, 124].
During the early stages of cardiac development, cardiomyocytes primarily rely on glycolysis as an energy source (80%). As cardiomyocytes mature and terminally differentiate, mitochondrial oxidative phosphorylation, mainly in the form of glucose oxidation and fatty acid β-oxidation, becomes the major energy source (80%). Interestingly, cardiac metabolism in patients with HCM switches back to a more fetal phenotype with an increase in glycolysis and a decrease in fatty acid β-oxidation . Similar to fetal cardiomyocytes, iPSC-CMs mainly rely on glycolysis as an energy source .
Approaches to enhance iPSC-CMs maturation
The relatively immature iPSC-CMs make HCM modeling challenging, as it is uncertain whether iPSC-CMs can recapitulate human HCM phenotypes. Furthermore, the understanding of early pathological events would be affected by the maturation levels of iPSC-CMs. Several culture methods and techniques have been proposed to generate mature and homogeneous cardiomyocytes, including prolonged culture time , triiodothyronine (T3) hormone treatment , microRNA (miRNA) overexpression , metabolic manipulation , increased substrate stiffness , three-dimensional (3D) tissue engineering , mechanical stress , and electrical stimulation . The general principle of these methods is to simulate in vivo environment and subject iPSC-CMs to relatively stable physical and/or humoral stimuli to promote their structural and functional maturation.
Initial studies tried to promote the maturation of iPSC-CMs by prolonging culture time. For example, late-stage iPSC-CMs (80–120 days of in vitro culture) exhibited increased cell volume, greater density and alignment of myofibril, and a significant increase in the proportion of multinucleated cardiomyocytes . Furthermore, long-term cultured iPSC-CMs showed elevated levels of mitochondrial oxidative phosphorylation, enhanced contractility, and responsiveness to isoproterenol . It has been reported that protein kinase A/proteasome-dependent signaling pathway modulated mitochondrial respiratory chain proteins and enhanced metabolic output of iPSC-CMs during long-term culture, resulting in increased cell contractility . However, the arrangement of sarcomeres in iPSC-CMs remained disordered compared to adult cardiomyocytes, suggesting that additional approaches are required to achieve full maturation.
The use of a T3-containing medium can promote the molecular, morphological, and functional maturation of iPSC-CMs, including increased expression of genes encoding sarcomeric proteins, improved sarcomeric organization, and increased action potential amplitudes and contraction force . Transcriptomic analysis revealed that iPSC-CMs treated with T3 were more mature than those without T3 . In addition, the combination of T3 and glucocorticoids promotes the formation of T-tubules and enhances excitation–contraction coupling . These studies indicate that T3 plays an important role in the maturation of iPSC-CMs. However, Bedada et al.  reported that T3 did not affect the cTnI/ssTnI ratio in iPSC-CMs, which could be used as a genetic marker for cardiomyocyte maturity.
Genomics revealed that miRNAs are key regulators during cardiac development . Delivering miRNAs to human embryonic stem cell-derived cardiomyocytes resulted in increased cell size and proportion of binucleated cells, improved sarcomere alignment and calcium handling . Furthermore, overexpression of miRNA in the let-7 family increased cell size and sarcomere length with enhanced contractility and mitochondrial oxidative phosphorylation, promoting the maturation of iPSC-CMs [128, 138].
During cardiac development, the main energy source of cardiomyocytes undergoes a shift from glycolysis to oxidative phosphorylation of fatty acids and glucose . This metabolic change can be mimicked by adjusting medium composition to promote maturation of iPSC-CMs, such as by replacing high-carbohydrate, glucose-based medium with a low-carbohydrate, fatty acid-based medium [129, 139]. However, a subsequent study indicated that culturing iPSC-CMs in a medium with rich fatty acids induced lipotoxicity, causing cell death . To overcome the disadvantage, the medium containing galactose and fatty acids was used to culture iPSC-CMs, which displayed elongated cell morphology, improved sarcomeric organization, increased myofibril force generation, and elevated levels of oxidative metabolism .
The dynamic cellular environment of heart tissues cannot be fully recapitulated in monolayer cultured cells. Culture substrates with increased stiffness were thus used to resemble those of the native tissue. It was reported that culturing mouse embryonic stem cells on polymer mattress can promote their differentiation into cardiomyocytes . By screening the library of polymers comprised of polyethylene glycol (PEG), hydrophobic poly-ε-caprolactone (PCL), and carboxylated PCL (CPCL), Chun et al.  found that culturing iPSC-CMs on a 4% PEG-96% PCL matrix enhanced cell contractility and mitochondrial function, as well as isoform switching from ssTnI to cTnI. Afterward, Feaster et al.  cultured iPSC-CMs on a 0.4- to 0.8-mm-thick mattress of undiluted Matrigel (mattress iPSC-CMs) and on a control substrate of 0.1-mm-thick, diluted Matrigel (control iPSC-CMs). Compared with control iPSC-CMs, mattress iPSC-CMs exhibited rod-shaped morphology, increased sarcomere length, and elevated expression levels of cTnI.
Monolayer cell culture models have been widely used in cardiovascular disease research. However, they can neither fully mimic the cellular environment in the heart nor recapitulate the architecture of myocardial tissue. In contrast, 3D cardiac tissues can better simulate the structure and microenvironment of the human heart, which are important for disease modeling . iPSC-CMs can be mixed with scaffolds to form engineered heart tissues (EHTs) . Consisting of collagen, gelatin, hyaluronic acid, and natural extracellular matrix (ECM) extracts, hydrogel scaffolds are frequently used to improve the maturation of iPSC-CMs and serve as a model for measuring myocardial tissues contractility [147, 148].
During human growth and development, subjecting cardiomyocytes to increasing workloads can promote cardiac maturation . Two recent studies demonstrated the feasibility of this approach by modulating tissue stress and electrical stimulation frequency, respectively. Abilez et al.  found that increasing the tension of EHTs could promote their maturation, including improved cell alignment and calcium dynamics, and increased expression of mature cardiomyocyte genes. In addition to mechanical stress, electrical stimulation can also improve the maturity of iPSC-CMs. It was reported that iPSC-CMs under electrical stimulation exhibited advanced levels of structural and functional maturity, e.g., adultlike gene expression profiles, remarkably organized sarcomere, the presence of T-tubules, positive force–frequency relationship, and improved calcium handling . Furthermore, the combination of these two approaches was also used to promote the maturation of iPSC-CMs .
HCM phenotypes of iPSC-CMs
With the advancement of culture and differentiation protocols, iPSCs can be efficiently generated and directed to differentiate into cardiomyocytes. Therefore, more and more laboratories use iPSC-CMs to establish in vitro HCM models [89, 152]. However, how do HCM phenotypes of iPSC-CMs compare with those of “real” human HCM? To solve this question, our study statistically analyzed 28 studies reporting phenotypes of human iPSC-CMs either derived from iPSC lines of patients with HCM or from human iPSC lines in which the HCM mutation had been genetically introduced (Table 3).
Histologically, HCM features pathological hypertrophy of cardiomyocytes, sarcomere disorder, and myocardial fibrosis. iPSC-CMs harboring HCM mutations exhibited hypertrophied cardiomyocytes and disorganized sarcomere [79, 153, 155]. In addition, previous studies have reported that the proportion of multinucleated cells was significantly increased in iPSC-CMs carrying HCM mutations [157, 166, 178]. However, studies have also shown that iPSC-CMs with HCM mutations varied widely in cell size with surface areas ranging from 800  to > 6000 μm2  and volumes ranging from 5.8  to 120 μm3 . iPSC-CMs appear extremely low compared to the 15,000–40,000 μm3 in adult cardiomyocytes . In addition to culture protocols (e.g., culturing time, medium composition, etc.), differences in imaging techniques and methods of measurement may explain the scatter.
Hypercontractility has been reported in several studies of mutations in MYH7, MYBPC3, ACTN2, and ACTC1. The analysis of single-cell video recordings by pixel quantification software confirmed hypercontractility of patient-derived iPSC-CMs with MYH7 R663H mutation . In addition, Cohn et al.  measured cardiac microtissues (CMTs) generated from iPSC-CMs with MYBPC3 R502W mutation by traction force microscopy, finding that the diseased CMTs generated increased twitch force and maximum contraction velocity but without changes in contraction time. HCM CMTs exhibited prolonged relaxation half-time, consistent with previously reported that impaired relaxation was the consequence of HCM mutations [15, 180]. Likewise, iPSC-CMs with ACTN2 T247M mutation exhibited increased peak force and prolonged relaxation time, but no change in time to peak contraction . Compared with healthy isogenic controls in which CRISPR/Cas9 was used to correct ACTC1 G301A mutation in patient lines, diseased iPSC-CMs displayed increased cell shortening, prolonged contraction, and relaxation times .
However, it has also been reported that iPSC-CMs with MYH7 and MYBPC3 mutations displayed a hypocontractile phenotype. Mosqueira et al.  generated a series of isogenic iPSC-CM lines with MYH7 R453C mutation and then formed EHTs to measure contractile properties. Surprisingly, they found that EHTs carrying the HCM mutation exhibited decreased contractile force and prolonged contraction time but little change in relaxation time. Moreover, Seeger et al.  used traction force microscopy in MYBPC3 R943x iPSC-CMs treated with dexamethasone, T3, and insulin-like growth factor 1 (IGF1) to characterize their contractility. The diseased iPSC-CMs showed significantly decreased contractile force generation, prolonged contraction, and relaxation kinetics.
The above studies have shown that iPSC-CMs with HCM mutations did not always exhibit hypercontractility, and even mutations in the same genes (such as MYH7 and MYBPC3) might display opposite phenotypes [156, 158, 171, 176]. Furthermore, different mutations may have different pathogenesis at the sarcomere level. For example, haploinsufficiency  may be the main pathogenesis of HCM phenotypes in MYBPC3 c.2373dupG iPSC-CMs, whereas activation of the nonsense-mediated mRNA decay (NMD) pathway  may be the main cause of HCM phenotypes in iPSC-CMs harboring MYBPC3 R943x mutation.
Abnormal calcium handling
The myocyte excitation–contraction coupling can be regulated, in part, through modulation of calcium release and uptake . It was reported that L-type calcium channels were excessively activated in HCM iPSC-CMs, resulting in a pronounced increase in Ca2+ currents and elevated [Ca2+]i . In addition, Lan et al.  reported that iPSC-CMs with MYH7 R663H mutation exhibited significantly smaller SR Ca2+ release as compared to controls, leading to increased [Ca2+]i. The increased [Ca2+]i can drive the electrogenic Na+–Ca2+ exchanger (NCX), resulting in further inward flow of Na + and inducing delayed after depolarization (DAD) .
The relationship between [Ca2+]i and contractile force is a highly regulated biological constant with half-maximal twitch force at [Ca2+]i of ~ 0.65 μM . Based on this, Wu et al.  defined the ratio of sarcomere contraction rate to Ca2+ transient amplitude (dF/Δ [Ca2+]i) as an indicator of myofilament calcium sensitivity, which was significantly increased in HCM iPSC-CMs. In addition, several studies have reported increased Ca2+ sensitivity in iPSC-CMs harboring HCM mutations [88, 158, 161, 173]. Clearly, increased Ca2+ sensitivity in HCM iPSC-CMs would lead to hypercontractility at lower [Ca2+]i and prolonged Ca2+ decay time, which is consistent with the clinical phenotypes of preserved systolic function but diastolic dysfunction in HCM patients .
Decreased energetic efficiency
HCM mutations may result in decreased energetic efficiency of the cross-bridge cycle, i.e., inefficient usage of ATP for contraction, and increased oxygen consumption [183, 184]. Likewise, iPSC-CMs with MYH7 c.C9123T mutation showed increased basal and maximal respiration, and elevated ATP production, but decreased contractility . In addition, HCM mutations frequently lead to increased energy depletion in cardiomyocytes. For instance, Toepfer et al.  in their study reported that MYH7 mutations increased proportions of myosins in the disordered relaxed state (DRX) conformation and decreased in the super relaxed state (SRX) conformations, which contributed to the significantly decreased phosphocreatine/ATP ratio, and energetic and metabolic stress.
Prospects of iPSCs in modeling HCM
Detecting the pathological significance of gene mutations and determining their pathogenicity
A major challenge faced by doctors treating patients with suspected HCM is how to determine the pathogenicity of specific genetic mutations, especially with incomplete gene penetrance and asymptomatic patients. With the development of iPSC technology, the HCM phenotype can be recapitulated in vitro by generating patient-derived iPSC-CMs . Furthermore, HCM mutations can be corrected by gene editing techniques such as the CRISPR-Cas9 system, eliminating differences in (epi)genetic background, and directly comparing the diseased iPSC-CMs with healthy isogenic iPSC-CMs to examine the causality of HCM mutations .
Discovering new disease mechanisms
The iPSC-CMs models can help us discover new cellular mechanisms caused by HCM mutations. For example, Wu et al.  proposed that elevated [Ca2+]i was a potential mechanism leading to the arrhythmic phenotype in HCM iPSC-CMs, which was validated in a subsequent study. The Jamie lab  reported that a combination treatment of dantrolene and ranolazine was performed on the iPSC-CM lines with MYH7 R403Q mutation to decrease [Ca2+]i. This strategy proved effective in significantly reducing the frequency of arrhythmic events in HCM iPSC-CMs to levels comparable to isogenic healthy controls. It remains to be seen whether this conclusion applies to other HCM mutations.
Verifying the disease mechanism caused by HCM mutations
iPSC technology allowed us to test the hypothesis generated from human studies. For example, the study of MYBPC3 R943x mutation by Seeger et al.  demonstrated that activation of the NMD pathway is the main pathogenesis of HCM. In addition, Prondzynski et al.  reported that iPSC-CMs harboring MYBPC3 c.1358-1359insC mutation displayed hypertrophic phenotype and had decreased MYBPC3 mRNA and cMyBPC levels, validating the hypothesis of haploinsufficiency.
Aiding risk stratification and prognosis
If the cellular phenotype of mutant-bearing iPSC-CMs is stable and reliable, and clinical relevance can be established, the cellular phenotype can serve as an indicator for clinical risk stratification and prognosis. To achieve this goal, in future studies, our efforts should focus on standardizing culture time, optimizing culture conditions, and developing novel maturation methods to generate normalized, mature iPSC-CMs. In addition, long-term, prospective studies of iPSC-CM phenotype/clinical phenotype correlation must be performed in a large number of mutation carriers to determine the natural history and prognosis of HCM.
Use for gene therapy
Gene editing technologies can specifically repair gene mutations and have broad prospects in the treatment of HCM. Prondzynski et al.  applied trans-splicing and gene replacement techniques to increase the expression of MYBPC3 in iPSC-CMs with MYBPC3c.1358-1359insC mutation. Adenovirus was used to, respectively, transfect iPSC-CMs with 5′ and 3′ trans-spliced molecules and MYBPC3 cDNA. However, both 5′ trans-splicing and 3′ trans-splicing strategies were inefficient with the trans-spliced cMyBPC protein not detectable. In contrast, whole-gene replacement increased the expression level of MYBPC3 to more than 80% in comparison with non-transduced control iPSC-CMs and prevented cell hypertrophy .
Testing of existing drugs
A major advantage of the iPSC technology is the ability to test the efficacy of drugs in mutation-specific or patient-specific iPSC-CMs. Several studies have reported that Ca2+ channel blocker verapamil can significantly improve HCM phenotypes including myocyte hypertrophy, Ca2+ handling abnormalities, and arrhythmia [79, 88]. However, a previous double-blind clinical trial showed that although verapamil improved symptoms in HCM patients, it failed to provide objective clinical benefits, such as exercise capacity . In addition, it was reported that ranolazine ameliorated arrhythmia and reduced the transduction of cellular hypertrophic signaling in iPSC-CMs with MYH7 R453C mutation [166, 170]. However, a randomized, double-blind, phase 2 study showed that ranolazine did not improve exercise capacity, diastolic function, or quality of life in HCM patients . The effects of these drugs at the cellular level did not carry over to the human body, possibly because pathological changes in organs (e.g., myocardial fibrosis) masked the effects of the drugs on cells. Future, more systematic studies will have to determine the validity of this approach.
Discovery of new drugs
Compared with animal models, the usage of disease- or patient-specific iPSC-CMs can better reflect the possible effects of drugs in humans. With further research, we can obtain more and more iPSC lines carrying different sarcomeric mutations, which can be used for high-throughput screening assays for drug discovery . Efficacy studies of drugs in iPSC-CMs of different genetic backgrounds are actually clinical trials in dishes, thereby reducing the high cost of current drug development and improving efficiency. Moreover, we can optimize patient selection for clinical trials by predicting patient responsiveness to drugs to achieve individualized treatment.
Testing drug safety
Human iPSC-CMs can be used to screen for drug-induced cardiovascular toxicity in time, including alterations in cardiac cellular contractility, electrophysiology, and viability . For example, breast cancer patient-derived iPSC-CMs treated with doxorubicin showed decreased cell viability, impaired calcium handling, and mitochondrial function, suggesting that breast cancer patients are more susceptible to doxorubicin-induced cardiotoxicity . In addition, iPSC-CMs derived from healthy people also exhibited cardiotoxicity induced by doxorubicin . Therefore, before clinical trials of new drugs, we can test their safety in iPSC-CMs.
Limitations to iPSC-derived disease models
Although the iPSCs have great therapeutic and translational potential, they show important limitations, such as low reprogramming efficiency, significant differences in the gene expression profiles with embryonic stem cells (ESCs), and teratogenicity.
One limitation is the low reprogramming efficiency of iPSCs, which ranges from 4.4% with modified mRNA and 1% with the retroviruses to as little as 0.001% with the adenovirus and plasmids . By contrast, B cell lines induced by C/EBPalpha can be converted into macrophage-like cells at 100% efficiency within 2–3 days . This shows the fact that the induction of pluripotency by specific factors in contrast to lineage switching faces more barriers, possibly because of the higher degree of epigenetic and transcriptional similarity among mature cells than between mature and pluripotent cell lines .
iPSCs were originally thought to resemble ESCs, given their similarity in morphology, proliferation, differentiation potential, and marker expression . However, this similarity was soon doubted, as differences were discovered in the gene expression profiles. It was found that iPSCs retain donor cell-specific transcriptome, along with the DNA methylation signature [194,195,196,197]. For example, Marchetto et al.  compared the gene expression profiles of iPSCs and ESCs. Their transcriptome analysis showed that iPSCs had insufficient induction of embryonic-specific genes. In addition, a group of genes were upregulated in iPSCs, but they were silenced in ESCs.
The potential teratoma formation after transplantation of iPSCs is another hurdle for clinical application. Incomplete differentiation and difficulty in eliminating undifferentiated cells may lead to potential teratoma formation [198, 199]. However, advances in the culture and differentiation protocol have largely overcome this challenge, resulting in highly pure iPSC-CMs [88, 141]. In addition to the teratogenicity by the undifferentiated cells, differentiated iPSCs still have the intrinsic risk of malignant transformation . The initial use of retroviruses with the integration of viruses into host cell DNA was a contributing factor. Furthermore, the overexpression of c-Myc increases the risk of teratoma formation . To overcome the limitations of retroviral vectors, non-integrating methods have been developed to induce pluripotency, which reduced the risk of genomic integration .
Apart from the above limitations, an important limitation to iPSC-derived disease models is present at the drug screening level, considering the fact that it is unlikely to measure all toxicities or side effects merely at the cellular level . Animal studies and clinical human research are required to investigate drug-induced changes and long-term side effects in nontarget tissues. Besides, with the advances in bioengineering technologies, we will be able to perfuse drug solutions on iPSC derivatives (e.g., cardiac, hepatic, pulmonary, and renal cells) to simulate the multiorgan interactions in the human body, as well as to assess toxicities and side effects.
Since Yamanaka discovered the method of inducing somatic cells into pluripotent stem cells, there has been impressive progress in reprogramming and differentiation protocols. Currently, we can stably acquire a large number of cardiomyocytes from iPSCs, which opens up a new field of HCM research with great potential. Due to the difficulty in obtaining human myocardial tissue and the species differences in animal models, iPSC-CMs have great advantages with their abundant sources. Several studies have demonstrated that iPSC-CM models can facilitate the study of HCM phenotypes and have made important contributions to elucidating molecular mechanisms. In addition, in combination with gene editing technologies such as CRISPR-Cas9 to directly introduce or repair specific HCM mutations, we can directly compare the differences between patient-specific iPSC-CMs and their isogenic control lines to determine the pathogenic role of HCM mutations.
Nonetheless, the research of HCM modeling with iPSC-CMs is still in its infancy. Differences between complex in vivo structures and pathophysiology and simplified in vitro conditions may result in the inability of iPSC-CMs to fully recapitulate the HCM phenotype. Furthermore, one of the challenges is the need to continue to improve the maturity of iPSC-CMs. Previous studies have shown that increasing substrate stiffness , mechanical stress , electrical stimulation , and several other methods can promote the maturation of iPSC-CMs to a certain degree. However, it is unclear how far maturation should improve before iPSC-CMs could be to be generally accepted as a model for studying HCM. Obviously, the use of iPSC-CMs for HCM modeling requires a set of CMs characteristic criteria, including cell morphology, sarcomeric protein isoforms, electrophysiology, calcium handling, mitochondrial function, and metabolism resembling adult cardiomyocytes (as listed in Table 2).
Although the 28 studies reporting iPSC-CMs with HCM mutations vary largely in cell size and no clear consensus is formed in other HCM phenotypes such as contractility, it is too early to conclude that iPSC-CMs cannot provide critical information for HCM models. These differences may reflect the diversity of culture conditions and measurement methods. Therefore, a consensus-implemented approach is needed to standardize the generation and differentiation of iPSCs, normalizing the structure and function properties of iPSC-CMs. We believe that with standardization of culture conditions and culture time, improvement in maturity, and emergence of more comprehensive measurement techniques, more predictive HCM models from iPSC-CMs would be built to better mimic HCM. This will help to provide a better understanding of the pathophysiology of HCM and its individualized treatment.
Availability of data and materials
Bone morphogenetic proteins
- [Ca2+]i :
Intracellular Ca2+ concentration
Clustered regularly interspaced short palindromic repeats associated protein
Cysteine- and glycine-rich protein in 3
Cardiac troponin I
Delayed after depolarization
Disordered relaxed state
Engineered heart tissues
Embryonic stem cells
- I Ca :
- I f :
Hyperpolarization-activated pacemaker current
- IGF 1:
Insulin-like growth factor 1
- I K1 :
Inward rectifier potassium current
- I Kr :
Rapid activating component of the delayed rectifier potassium current
- I Ks :
Slow activating component of the delayed rectifier potassium current
- I Na :
Induced pluripotent stem cells
Induced pluripotent stem cells-derived cardiomyocytes
- I to :
Transient outward potassium current
Myosin heavy chain
Myosin-binding protein C
Myosin light chain 2
Myosin light chain 3
Nonhomologous end joining
Nonsense-mediated mRNA decay
Premature termination codons
Slow skeletal troponin I
Transcription activator-like effector nucleases
Ommen SR, Mital S, Burke MA, Day SM, Deswal A, Elliott P, Evanovich LL, Hung J, Joglar JA, Kantor P, et al. 2020 AHA/ACC guideline for the diagnosis and treatment of patients with hypertrophic cardiomyopathy: executive summary: a report of the American college of cardiology/American heart association joint committee on clinical practice guidelines. Circulation. 2020;142(25):e533–57.
Maron BJ. Clinical course and management of hypertrophic cardiomyopathy. N Engl J Med. 2018;379(7):655–68.
Tuohy CV, Kaul S, Song HK, Nazer B, Heitner SB. Hypertrophic cardiomyopathy: the future of treatment. Eur J Heart Fail. 2020;22(2):228–40.
Authors/Task Force m, Elliott PM, Anastasakis A, Borger MA, Borggrefe M, Cecchi F, Charron P, Hagege AA, Lafont A, Limongelli G, et al. 2014 ESC guidelines on diagnosis and management of hypertrophic cardiomyopathy: the task force for the diagnosis and management of hypertrophic cardiomyopathy of the European Society of cardiology (ESC). Eur Heart J. 2014;35(39):2733–79.
Semsarian C, Ingles J, Maron MS, Maron BJ. New perspectives on the prevalence of hypertrophic cardiomyopathy. J Am Coll Cardiol. 2015;65(12):1249–54.
Bick AG, Flannick J, Ito K, Cheng S, Vasan RS, Parfenov MG, Herman DS, DePalma SR, Gupta N, Gabriel SB, et al. Burden of rare sarcomere gene variants in the Framingham and Jackson heart study cohorts. Am J Hum Genet. 2012;91(3):513–9.
Pare JA, Fraser RG, Pirozynski WJ, Shanks JA, Stubington D. Hereditary cardiovascular dysplasia. A form of familial cardiomyopathy. Am J Med. 1961;31:37–62.
Geisterfer-Lowrance AA, Kass S, Tanigawa G, Vosberg HP, McKenna W, Seidman CE, Seidman JG. A molecular basis for familial hypertrophic cardiomyopathy: a beta cardiac myosin heavy chain gene missense mutation. Cell. 1990;62(5):999–1006.
Walsh R, Thomson KL, Ware JS, Funke BH, Woodley J, McGuire KJ, Mazzarotto F, Blair E, Seller A, Taylor JC, et al. Reassessment of mendelian gene pathogenicity using 7,855 cardiomyopathy cases and 60,706 reference samples. Genet Med. 2017;19(2):192–203.
Alfares AA, Kelly MA, McDermott G, Funke BH, Lebo MS, Baxter SB, Shen J, McLaughlin HM, Clark EH, Babb LJ, et al. Results of clinical genetic testing of 2,912 probands with hypertrophic cardiomyopathy: expanded panels offer limited additional sensitivity. Genet Med. 2015;17(11):880–8.
Maron BJ, Maron MS, Semsarian C. Genetics of hypertrophic cardiomyopathy after 20 years: clinical perspectives. J Am Coll Cardiol. 2012;60(8):705–15.
Teekakirikul P, Zhu W, Huang HC, Fung E. Hypertrophic cardiomyopathy: an overview of genetics and management. Biomolecules. 2019;9(12):878.
Mosqueira D, Smith JGW, Bhagwan JR, Denning C. Modeling hypertrophic cardiomyopathy: mechanistic insights and pharmacological intervention. Trends Mol Med. 2019;25(9):775–90.
Millat G, Bouvagnet P, Chevalier P, Dauphin C, Jouk PS, Da Costa A, Prieur F, Bresson JL, Faivre L, Eicher JC, et al. Prevalence and spectrum of mutations in a cohort of 192 unrelated patients with hypertrophic cardiomyopathy. Eur J Med Genet. 2010;53(5):261–7.
Marian AJ, Braunwald E. Hypertrophic cardiomyopathy: genetics, pathogenesis, clinical manifestations, diagnosis, and therapy. Circ Res. 2017;121(7):749–70.
Erdmann J, Daehmlow S, Wischke S, Senyuva M, Werner U, Raible J, Tanis N, Dyachenko S, Hummel M, Hetzer R, et al. Mutation spectrum in a large cohort of unrelated consecutive patients with hypertrophic cardiomyopathy. Clin Genet. 2003;64(4):339–49.
Richard P, Charron P, Carrier L, Ledeuil C, Cheav T, Pichereau C, Benaiche A, Isnard R, Dubourg O, Burban M, et al. Hypertrophic cardiomyopathy: distribution of disease genes, spectrum of mutations, and implications for a molecular diagnosis strategy. Circulation. 2003;107(17):2227–32.
Kimura A, Harada H, Park JE, Nishi H, Satoh M, Takahashi M, Hiroi S, Sasaoka T, Ohbuchi N, Nakamura T, et al. Mutations in the cardiac troponin I gene associated with hypertrophic cardiomyopathy. Nat Genet. 1997;16(4):379–82.
Mogensen J, Klausen IC, Pedersen AK, Egeblad H, Bross P, Kruse TA, Gregersen N, Hansen PS, Baandrup U, Borglum AD. Alpha-cardiac actin is a novel disease gene in familial hypertrophic cardiomyopathy. J Clin Invest. 1999;103(10):R39-43.
Poetter K, Jiang H, Hassanzadeh S, Master SR, Chang A, Dalakas MC, Rayment I, Sellers JR, Fananapazir L, Epstein ND. Mutations in either the essential or regulatory light chains of myosin are associated with a rare myopathy in human heart and skeletal muscle. Nat Genet. 1996;13(1):63–9.
Geier C, Gehmlich K, Ehler E, Hassfeld S, Perrot A, Hayess K, Cardim N, Wenzel K, Erdmann B, Krackhardt F, et al. Beyond the sarcomere: CSRP3 mutations cause hypertrophic cardiomyopathy. Hum Mol Genet. 2008;17(18):2753–65.
Walsh R, Buchan R, Wilk A, John S, Felkin LE, Thomson KL, Chiaw TH, Loong CCW, Pua CJ, Raphael C, et al. Defining the genetic architecture of hypertrophic cardiomyopathy: re-evaluating the role of non-sarcomeric genes. Eur Heart J. 2017;38(46):3461–8.
Marian AJ. The case of “missing causal genes” and the practice of medicine: a Sherlock Holmes approach of deductive reasoning. Circ Res. 2016;119(1):21–4.
Cahill TJ, Ashrafian H, Watkins H. Genetic cardiomyopathies causing heart failure. Circ Res. 2013;113(6):660–75.
Marian AJ. Causality in genetics: the gradient of genetic effects and back to Koch’s postulates of causality. Circ Res. 2014;114(2):e18-21.
MacArthur DG, Manolio TA, Dimmock DP, Rehm HL, Shendure J, Abecasis GR, Adams DR, Altman RB, Antonarakis SE, Ashley EA, et al. Guidelines for investigating causality of sequence variants in human disease. Nature. 2014;508(7497):469–76.
Genomes Project C, Auton A, Brooks LD, Durbin RM, Garrison EP, Kang HM, Korbel JO, Marchini JL, McCarthy S, McVean GA, et al. A global reference for human genetic variation. Nature. 2015;526(7571):68–74.
Genomes Project C, Abecasis GR, Altshuler D, Auton A, Brooks LD, Durbin RM, Gibbs RA, Hurles ME, McVean GA. A map of human genome variation from population-scale sequencing. Nature. 2010;467(7319):1061–73.
Duncker DJ, Bakkers J, Brundel BJ, Robbins J, Tardiff JC, Carrier L. Animal and in silico models for the study of sarcomeric cardiomyopathies. Cardiovasc Res. 2015;105(4):439–48.
Geisterfer-Lowrance AA, Christe M, Conner DA, Ingwall JS, Schoen FJ, Seidman CE, Seidman JG. A mouse model of familial hypertrophic cardiomyopathy. Science. 1996;272(5262):731–4.
Carrier L, Knoll R, Vignier N, Keller DI, Bausero P, Prudhon B, Isnard R, Ambroisine ML, Fiszman M, Ross J Jr, et al. Asymmetric septal hypertrophy in heterozygous cMyBP-C null mice. Cardiovasc Res. 2004;63(2):293–304.
Gedicke-Hornung C, Behrens-Gawlik V, Reischmann S, Geertz B, Stimpel D, Weinberger F, Schlossarek S, Precigout G, Braren I, Eschenhagen T, et al. Rescue of cardiomyopathy through U7snRNA-mediated exon skipping in Mybpc3-targeted knock-in mice. EMBO Mol Med. 2013;5(7):1128–45.
McConnell BK, Jones KA, Fatkin D, Arroyo LH, Lee RT, Aristizabal O, Turnbull DH, Georgakopoulos D, Kass D, Bond M, et al. Dilated cardiomyopathy in homozygous myosin-binding protein-C mutant mice. J Clin Invest. 1999;104(9):1235–44.
McConnell BK, Fatkin D, Semsarian C, Jones KA, Georgakopoulos D, Maguire CT, Healey MJ, Mudd JO, Moskowitz IP, Conner DA, et al. Comparison of two murine models of familial hypertrophic cardiomyopathy. Circ Res. 2001;88(4):383–9.
Harris SP, Bartley CR, Hacker TA, McDonald KS, Douglas PS, Greaser ML, Powers PA, Moss RL. Hypertrophic cardiomyopathy in cardiac myosin binding protein-C knockout mice. Circ Res. 2002;90(5):594–601.
Vignier N, Schlossarek S, Fraysse B, Mearini G, Kramer E, Pointu H, Mougenot N, Guiard J, Reimer R, Hohenberg H, et al. Nonsense-mediated mRNA decay and ubiquitin-proteasome system regulate cardiac myosin-binding protein C mutant levels in cardiomyopathic mice. Circ Res. 2009;105(3):239–48.
Mearini G, Stimpel D, Kramer E, Geertz B, Braren I, Gedicke-Hornung C, Precigout G, Muller OJ, Katus HA, Eschenhagen T, et al. Repair of Mybpc3 mRNA by 5′-trans-splicing in a mouse model of hypertrophic cardiomyopathy. Mol Ther Nucleic Acids. 2013;2:e102.
Mearini G, Stimpel D, Geertz B, Weinberger F, Kramer E, Schlossarek S, Mourot-Filiatre J, Stoehr A, Dutsch A, Wijnker PJ, et al. Mybpc3 gene therapy for neonatal cardiomyopathy enables long-term disease prevention in mice. Nat Commun. 2014;5:5515.
van der Velden J, Ho CY, Tardiff JC, Olivotto I, Knollmann BC, Carrier L. Research priorities in sarcomeric cardiomyopathies. Cardiovasc Res. 2015;105(4):449–56.
Eschenhagen T, Carrier L. Cardiomyopathy phenotypes in human-induced pluripotent stem cell-derived cardiomyocytes-a systematic review. Pflugers Arch. 2019;471(5):755–68.
Eschenhagen T, Mummery C, Knollmann BC. Modelling sarcomeric cardiomyopathies in the dish: from human heart samples to iPSC cardiomyocytes. Cardiovasc Res. 2015;105(4):424–38.
Takahashi K, Tanabe K, Ohnuki M, Narita M, Ichisaka T, Tomoda K, Yamanaka S. Induction of pluripotent stem cells from adult human fibroblasts by defined factors. Cell. 2007;131(5):861–72.
Aasen T, Raya A, Barrero MJ, Garreta E, Consiglio A, Gonzalez F, Vassena R, Bilic J, Pekarik V, Tiscornia G, et al. Efficient and rapid generation of induced pluripotent stem cells from human keratinocytes. Nat Biotechnol. 2008;26(11):1276–84.
Zhou T, Benda C, Dunzinger S, Huang Y, Ho JC, Yang J, Wang Y, Zhang Y, Zhuang Q, Li Y, et al. Generation of human induced pluripotent stem cells from urine samples. Nat Protoc. 2012;7(12):2080–9.
Seki T, Yuasa S, Fukuda K. Generation of induced pluripotent stem cells from a small amount of human peripheral blood using a combination of activated T cells and Sendai virus. Nat Protoc. 2012;7(4):718–28.
Buikema JW, Wu SM. Untangling the biology of genetic cardiomyopathies with pluripotent stem cell disease models. Curr Cardiol Rep. 2017;19(4):30.
Nakagawa M, Koyanagi M, Tanabe K, Takahashi K, Ichisaka T, Aoi T, Okita K, Mochiduki Y, Takizawa N, Yamanaka S. Generation of induced pluripotent stem cells without Myc from mouse and human fibroblasts. Nat Biotechnol. 2008;26(1):101–6.
Takahashi K, Yamanaka S. Induction of pluripotent stem cells from mouse embryonic and adult fibroblast cultures by defined factors. Cell. 2006;126(4):663–76.
Okita K, Ichisaka T, Yamanaka S. Generation of germline-competent induced pluripotent stem cells. Nature. 2007;448(7151):313–7.
Blelloch R, Venere M, Yen J, Ramalho-Santos M. Generation of induced pluripotent stem cells in the absence of drug selection. Cell Stem Cell. 2007;1(3):245–7.
Yu J, Vodyanik MA, Smuga-Otto K, Antosiewicz-Bourget J, Frane JL, Tian S, Nie J, Jonsdottir GA, Ruotti V, Stewart R, et al. Induced pluripotent stem cell lines derived from human somatic cells. Science. 2007;318(5858):1917–20.
Brambrink T, Foreman R, Welstead GG, Lengner CJ, Wernig M, Suh H, Jaenisch R. Sequential expression of pluripotency markers during direct reprogramming of mouse somatic cells. Cell Stem Cell. 2008;2(2):151–9.
Soldner F, Hockemeyer D, Beard C, Gao Q, Bell GW, Cook EG, Hargus G, Blak A, Cooper O, Mitalipova M, et al. Parkinson’s disease patient-derived induced pluripotent stem cells free of viral reprogramming factors. Cell. 2009;136(5):964–77.
Okita K, Yamakawa T, Matsumura Y, Sato Y, Amano N, Watanabe A, Goshima N, Yamanaka S. An efficient nonviral method to generate integration-free human-induced pluripotent stem cells from cord blood and peripheral blood cells. Stem Cells. 2013;31(3):458–66.
Zhou W, Freed CR. Adenoviral gene delivery can reprogram human fibroblasts to induced pluripotent stem cells. Stem Cells. 2009;27(11):2667–74.
Fusaki N, Ban H, Nishiyama A, Saeki K, Hasegawa M. Efficient induction of transgene-free human pluripotent stem cells using a vector based on Sendai virus, an RNA virus that does not integrate into the host genome. Proc Jpn Acad Ser B Phys Biol Sci. 2009;85(8):348–62.
Yu J, Hu K, Smuga-Otto K, Tian S, Stewart R, Slukvin II, Thomson JA. Human induced pluripotent stem cells free of vector and transgene sequences. Science. 2009;324(5928):797–801.
Okita K, Nakagawa M, Hyenjong H, Ichisaka T, Yamanaka S. Generation of mouse induced pluripotent stem cells without viral vectors. Science. 2008;322(5903):949–53.
Warren L, Manos PD, Ahfeldt T, Loh YH, Li H, Lau F, Ebina W, Mandal PK, Smith ZD, Meissner A, et al. Highly efficient reprogramming to pluripotency and directed differentiation of human cells with synthetic modified mRNA. Cell Stem Cell. 2010;7(5):618–30.
Kim D, Kim CH, Moon JI, Chung YG, Chang MY, Han BS, Ko S, Yang E, Cha KY, Lanza R, et al. Generation of human induced pluripotent stem cells by direct delivery of reprogramming proteins. Cell Stem Cell. 2009;4(6):472–6.
Li W, Ding S. Small molecules that modulate embryonic stem cell fate and somatic cell reprogramming. Trends Pharmacol Sci. 2010;31(1):36–45.
Laflamme MA, Murry CE. Heart regeneration. Nature. 2011;473(7347):326–35.
Hatani T, Miki K, Yoshida Y. Induction of human induced pluripotent stem cells to cardiomyocytes using embryoid bodies. Methods Mol Biol. 2018;1816:79–92.
Zhang J, Wilson GF, Soerens AG, Koonce CH, Yu J, Palecek SP, Thomson JA, Kamp TJ. Functional cardiomyocytes derived from human induced pluripotent stem cells. Circ Res. 2009;104(4):e30-41.
Mummery C, Ward-van Oostwaard D, Doevendans P, Spijker R, van den Brink S, Hassink R, van der Heyden M, Opthof T, Pera M, de la Riviere AB, et al. Differentiation of human embryonic stem cells to cardiomyocytes: role of coculture with visceral endoderm-like cells. Circulation. 2003;107(21):2733–40.
Hamad S, Derichsweiler D, Papadopoulos S, Nguemo F, Saric T, Sachinidis A, Brockmeier K, Hescheler J, Boukens BJ, Pfannkuche K. Generation of human induced pluripotent stem cell-derived cardiomyocytes in 2D monolayer and scalable 3D suspension bioreactor cultures with reduced batch-to-batch variations. Theranostics. 2019;9(24):7222–38.
Arai A, Yamamoto K, Toyama J. Murine cardiac progenitor cells require visceral embryonic endoderm and primitive streak for terminal differentiation. Dev Dyn. 1997;210(3):344–53.
Freund C, Davis RP, Gkatzis K, Ward-van Oostwaard D, Mummery CL. The first reported generation of human induced pluripotent stem cells (iPS cells) and iPS cell-derived cardiomyocytes in the Netherlands. Neth Heart J. 2010;18(1):51–4.
Osafune K, Caron L, Borowiak M, Martinez RJ, Fitz-Gerald CS, Sato Y, Cowan CA, Chien KR, Melton DA. Marked differences in differentiation propensity among human embryonic stem cell lines. Nat Biotechnol. 2008;26(3):313–5.
Wang H, Hao J, Hong CC. Cardiac induction of embryonic stem cells by a small molecule inhibitor of Wnt/beta-catenin signaling. ACS Chem Biol. 2011;6(2):192–7.
Laflamme MA, Chen KY, Naumova AV, Muskheli V, Fugate JA, Dupras SK, Reinecke H, Xu C, Hassanipour M, Police S, et al. Cardiomyocytes derived from human embryonic stem cells in pro-survival factors enhance function of infarcted rat hearts. Nat Biotechnol. 2007;25(9):1015–24.
Yang L, Soonpaa MH, Adler ED, Roepke TK, Kattman SJ, Kennedy M, Henckaerts E, Bonham K, Abbott GW, Linden RM, et al. Human cardiovascular progenitor cells develop from a KDR+ embryonic-stem-cell-derived population. Nature. 2008;453(7194):524–8.
Bhattacharya S, Burridge PW, Kropp EM, Chuppa SL, Kwok WM, Wu JC, Boheler KR, Gundry RL. High efficiency differentiation of human pluripotent stem cells to cardiomyocytes and characterization by flow cytometry. J Vis Exp. 2014;91:52010.
Kadari A, Mekala S, Wagner N, Malan D, Koth J, Doll K, Stappert L, Eckert D, Peitz M, Matthes J, et al. Robust generation of cardiomyocytes from human iPS cells requires precise modulation of BMP and WNT signaling. Stem Cell Rev Rep. 2015;11(4):560–9.
Besser RR, Ishahak M, Mayo V, Carbonero D, Claure I, Agarwal A. Engineered microenvironments for maturation of stem cell derived cardiac myocytes. Theranostics. 2018;8(1):124–40.
Burridge PW, Matsa E, Shukla P, Lin ZC, Churko JM, Ebert AD, Lan F, Diecke S, Huber B, Mordwinkin NM, et al. Chemically defined generation of human cardiomyocytes. Nat Methods. 2014;11(8):855–60.
Tiburcy M, Hudson JE, Balfanz P, Schlick S, Meyer T, Chang Liao ML, Levent E, Raad F, Zeidler S, Wingender E, et al. Defined engineered human myocardium with advanced maturation for applications in heart failure modeling and repair. Circulation. 2017;135(19):1832–47.
Batalov I, Feinberg AW. Differentiation of cardiomyocytes from human pluripotent stem cells using monolayer culture. Biomark Insights. 2015;10(Suppl 1):71–6.
Lan F, Lee AS, Liang P, Sanchez-Freire V, Nguyen PK, Wang L, Han L, Yen M, Wang Y, Sun N, et al. Abnormal calcium handling properties underlie familial hypertrophic cardiomyopathy pathology in patient-specific induced pluripotent stem cells. Cell Stem Cell. 2013;12(1):101–13.
Sala L, Bellin M, Mummery CL. Integrating cardiomyocytes from human pluripotent stem cells in safety pharmacology: Has the time come? Br J Pharmacol. 2017;174(21):3749–65.
Wang J, Li W, Han Y, Chen Y. Different clinical presentation and tissue characterization in a monozygotic twin pair with MYH7 mutation-related hypertrophic cardiomyopathy. Int Heart J. 2019;60(2):477–81.
Dzilic E, Lahm H, Dressen M, Deutsch MA, Lange R, Wu SM, Krane M, Doppler SA. Genome editing redefines precision medicine in the cardiovascular field. Stem Cells Int. 2018;2018:4136473.
Jinek M, Chylinski K, Fonfara I, Hauer M, Doudna JA, Charpentier E. A programmable dual-RNA-guided DNA endonuclease in adaptive bacterial immunity. Science. 2012;337(6096):816–21.
Zetsche B, Gootenberg JS, Abudayyeh OO, Slaymaker IM, Makarova KS, Essletzbichler P, Volz SE, Joung J, van der Oost J, Regev A, et al. Cpf1 is a single RNA-guided endonuclease of a class 2 CRISPR-Cas system. Cell. 2015;163(3):759–71.
Joung JK, Sander JD. TALENs: a widely applicable technology for targeted genome editing. Nat Rev Mol Cell Biol. 2013;14(1):49–55.
Zhang L, Jia R, Palange NJ, Satheka AC, Togo J, An Y, Humphrey M, Ban L, Ji Y, Jin H, et al. Large genomic fragment deletions and insertions in mouse using CRISPR/Cas9. PLoS ONE. 2015;10(3):e0120396.
Long C, Li H, Tiburcy M, Rodriguez-Caycedo C, Kyrychenko V, Zhou H, Zhang Y, Min YL, Shelton JM, Mammen PPA, et al. Correction of diverse muscular dystrophy mutations in human engineered heart muscle by single-site genome editing. Sci Adv. 2014;4(1):eaap9004.
Wu H, Yang H, Rhee JW, Zhang JZ, Lam CK, Sallam K, Chang ACY, Ma N, Lee J, Zhang H, et al. Modelling diastolic dysfunction in induced pluripotent stem cell-derived cardiomyocytes from hypertrophic cardiomyopathy patients. Eur Heart J. 2019;40(45):3685–95.
Parrotta EI, Lucchino V, Scaramuzzino L, Scalise S, Cuda G. Modeling cardiac disease mechanisms using induced pluripotent stem cell-derived cardiomyocytes: progress, promises and challenges. Int J Mol Sci. 2020;21(12):4354.
Gallagher DN, Haber JE. Repair of a site-specific DNA cleavage: old-school lessons for Cas9-mediated gene editing. ACS Chem Biol. 2018;13(2):397–405.
Anderson DM, Anderson KM, Chang CL, Makarewich CA, Nelson BR, McAnally JR, Kasaragod P, Shelton JM, Liou J, Bassel-Duby R, et al. A micropeptide encoded by a putative long noncoding RNA regulates muscle performance. Cell. 2015;160(4):595–606.
Jaffre F, Miller CL, Schanzer A, Evans T, Roberts AE, Hahn A, Kontaridis MI. Inducible pluripotent stem cell–derived cardiomyocytes reveal aberrant extracellular regulated kinase 5 and mitogen-activated protein kinase kinase 1/2 signaling concomitantly promote hypertrophic cardiomyopathy in RAF1-associated Noonan syndrome. Circulation. 2019;140(3):207–24.
Xiang X, Li C, Chen X, Dou H, Li Y, Zhang X, Luo Y. CRISPR/Cas9-mediated gene tagging: a step-by-step protocol. Methods Mol Biol. 2019;1961:255–69.
Roberts B, Hendershott MC, Arakaki J, Gerbin KA, Malik H, Nelson A, Gehring J, Hookway C, Ludmann SA, Yang R, et al. Fluorescent gene tagging of transcriptionally silent genes in hiPSCs. Stem Cell Rep. 2019;12(5):1145–58.
Liu M, Rehman S, Tang X, Gu K, Fan Q, Chen D, Ma W. Methodologies for improving HDR efficiency. Front Genet. 2018;9:691.
Pawelczak KS, Gavande NS, VanderVere-Carozza PS, Turchi JJ. Modulating DNA repair pathways to improve precision genome engineering. ACS Chem Biol. 2018;13(2):389–96.
Smirnikhina SA, Anuchina AA, Lavrov AV. Ways of improving precise knock-in by genome-editing technologies. Hum Genet. 2019;138(1):1–19.
Savic N, Ringnalda FC, Lindsay H, Berk C, Bargsten K, Li Y, Neri D, Robinson MD, Ciaudo C, Hall J, et al. Covalent linkage of the DNA repair template to the CRISPR-Cas9 nuclease enhances homology-directed repair. Elife. 2018;7:e33761.
Savic N, Ringnalda FC, Berk C, Bargsten K, Hall J, Jinek M, Schwank G. In vitro generation of CRISPR-Cas9 complexes with covalently bound repair templates for genome editing in mammalian cells. Bio Protoc. 2019;9(1):e3136.
Komor AC, Kim YB, Packer MS, Zuris JA, Liu DR. Programmable editing of a target base in genomic DNA without double-stranded DNA cleavage. Nature. 2016;533(7603):420–4.
Gerdes AM, Kellerman SE, Moore JA, Muffly KE, Clark LC, Reaves PY, Malec KB, McKeown PP, Schocken DD. Structural remodeling of cardiac myocytes in patients with ischemic cardiomyopathy. Circulation. 1992;86(2):426–30.
Ribeiro AJ, Ang YS, Fu JD, Rivas RN, Mohamed TM, Higgs GC, Srivastava D, Pruitt BL. Contractility of single cardiomyocytes differentiated from pluripotent stem cells depends on physiological shape and substrate stiffness. Proc Natl Acad Sci U S A. 2015;112(41):12705–10.
Hwang HS, Kryshtal DO, Feaster TK, Sanchez-Freire V, Zhang J, Kamp TJ, Hong CC, Wu JC, Knollmann BC. Comparable calcium handling of human iPSC-derived cardiomyocytes generated by multiple laboratories. J Mol Cell Cardiol. 2015;85:79–88.
Parikh SS, Blackwell DJ, Gomez-Hurtado N, Frisk M, Wang L, Kim K, Dahl CP, Fiane A, Tonnessen T, Kryshtal DO, et al. Thyroid and glucocorticoid hormones promote functional T-tubule development in human-induced pluripotent stem cell-derived cardiomyocytes. Circ Res. 2017;121(12):1323–30.
Mollova M, Bersell K, Walsh S, Savla J, Das LT, Park SY, Silberstein LE, Dos Remedios CG, Graham D, Colan S, et al. Cardiomyocyte proliferation contributes to heart growth in young humans. Proc Natl Acad Sci U S A. 2013;110(4):1446–51.
Hunkeler NM, Kullman J, Murphy AM. Troponin I isoform expression in human heart. Circ Res. 1991;69(5):1409–14.
Bedada FB, Chan SS, Metzger SK, Zhang L, Zhang J, Garry DJ, Kamp TJ, Kyba M, Metzger JM. Acquisition of a quantitative, stoichiometrically conserved ratiometric marker of maturation status in stem cell-derived cardiac myocytes. Stem Cell Rep. 2014;3(4):594–605.
Samak M, Hinkel R. Stem cells in cardiovascular medicine: historical overview and future prospects. Cells. 2019;8(12):1530.
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(4):505–13.
Streckfuss-Bomeke K, Tiburcy M, Fomin A, Luo X, Li W, Fischer C, Ozcelik C, Perrot A, Sossalla S, Haas J, et al. Severe DCM phenotype of patient harboring RBM20 mutation S635A can be modeled by patient-specific induced pluripotent stem cell-derived cardiomyocytes. J Mol Cell Cardiol. 2017;113:9–21.
Ma J, Guo L, Fiene SJ, Anson BD, Thomson JA, Kamp TJ, Kolaja KL, Swanson BJ, January CT. High purity human-induced pluripotent stem cell-derived cardiomyocytes: electrophysiological properties of action potentials and ionic currents. Am J Physiol Heart Circ Physiol. 2011;301(5):H2006–17.
Karakikes I, Ameen M, Termglinchan V, Wu JC. Human induced pluripotent stem cell-derived cardiomyocytes: insights into molecular, cellular, and functional phenotypes. Circ Res. 2015;117(1):80–8.
Davis RP, Casini S, van den Berg CW, Hoekstra M, Remme CA, Dambrot C, Salvatori D, Oostwaard DW, Wilde AA, Bezzina CR, et al. Cardiomyocytes derived from pluripotent stem cells recapitulate electrophysiological characteristics of an overlap syndrome of cardiac sodium channel disease. Circulation. 2012;125(25):3079–91.
Yazawa M, Hsueh B, Jia X, Pasca AM, Bernstein JA, Hallmayer J, Dolmetsch RE. Using induced pluripotent stem cells to investigate cardiac phenotypes in timothy syndrome. Nature. 2011;471(7337):230–4.
Bers DM. Cardiac excitation-contraction coupling. Nature. 2002;415(6868):198–205.
Itzhaki I, Rapoport S, Huber I, Mizrahi I, Zwi-Dantsis L, Arbel G, Schiller J, Gepstein L. Calcium handling in human induced pluripotent stem cell derived cardiomyocytes. PLoS ONE. 2011;6(4): e18037.
Sun X, Nunes SS. Biowire platform for maturation of human pluripotent stem cell-derived cardiomyocytes. Methods. 2016;101:21–6.
Rao C, Prodromakis T, Kolker L, Chaudhry UA, Trantidou T, Sridhar A, Weekes C, Camelliti P, Harding SE, Darzi A, et al. The effect of microgrooved culture substrates on calcium cycling of cardiac myocytes derived from human induced pluripotent stem cells. Biomaterials. 2013;34(10):2399–411.
Lee YK, Ng KM, Lai WH, Chan YC, Lau YM, Lian Q, Tse HF, Siu CW. Calcium homeostasis in human induced pluripotent stem cell-derived cardiomyocytes. Stem Cell Rev Rep. 2011;7(4):976–86.
Gherghiceanu M, Barad L, Novak A, Reiter I, Itskovitz-Eldor J, Binah O, Popescu LM. Cardiomyocytes derived from human embryonic and induced pluripotent stem cells: comparative ultrastructure. J Cell Mol Med. 2011;15(11):2539–51.
Schaper J, Meiser E, Stammler G. Ultrastructural morphometric analysis of myocardium from dogs, rats, hamsters, mice, and from human hearts. Circ Res. 1985;56(3):377–91.
Paik DT, Chandy M, Wu JC. Patient and disease-specific induced pluripotent stem cells for discovery of personalized cardiovascular drugs and therapeutics. Pharmacol Rev. 2020;72(1):320–42.
Chung S, Dzeja PP, Faustino RS, Perez-Terzic C, Behfar A, Terzic A. Mitochondrial oxidative metabolism is required for the cardiac differentiation of stem cells. Nat Clin Pract Cardiovasc Med. 2007;4(Suppl 1):S60–7.
Hattori F, Chen H, Yamashita H, Tohyama S, Satoh YS, Yuasa S, Li W, Yamakawa H, Tanaka T, Onitsuka T, et al. Nongenetic method for purifying stem cell-derived cardiomyocytes. Nat Methods. 2010;7(1):61–6.
Lopaschuk GD, Jaswal JS. Energy metabolic phenotype of the cardiomyocyte during development, differentiation, and postnatal maturation. J Cardiovasc Pharmacol. 2010;56(2):130–40.
Kim C, Wong J, Wen J, Wang S, Wang C, Spiering S, Kan NG, Forcales S, Puri PL, Leone TC, et al. Studying arrhythmogenic right ventricular dysplasia with patient-specific iPSCs. Nature. 2013;494(7435):105–10.
Lundy SD, Zhu WZ, Regnier M, Laflamme MA. Structural and functional maturation of cardiomyocytes derived from human pluripotent stem cells. Stem Cells Dev. 2013;22(14):1991–2002.
Kuppusamy KT, Jones DC, Sperber H, Madan A, Fischer KA, Rodriguez ML, Pabon L, Zhu WZ, Tulloch NL, Yang X, et al. Let-7 family of microRNA is required for maturation and adult-like metabolism in stem cell-derived cardiomyocytes. Proc Natl Acad Sci U S A. 2015;112(21):E2785–94.
Mills RJ, Titmarsh DM, Koenig X, Parker BL, Ryall JG, Quaife-Ryan GA, Voges HK, Hodson MP, Ferguson C, Drowley L, et al. Functional screening in human cardiac organoids reveals a metabolic mechanism for cardiomyocyte cell cycle arrest. Proc Natl Acad Sci U S A. 2017;114(40):E8372–81.
Yang X, Pabon L, Murry CE. Engineering adolescence: maturation of human pluripotent stem cell-derived cardiomyocytes. Circ Res. 2014;114(3):511–23.
Abilez OJ, Tzatzalos E, Yang H, Zhao MT, Jung G, Zollner AM, Tiburcy M, Riegler J, Matsa E, Shukla P, et al. Passive stretch induces structural and functional maturation of engineered heart muscle as predicted by computational modeling. Stem Cells. 2018;36(2):265–77.
Chan YC, Ting S, Lee YK, Ng KM, Zhang J, Chen Z, Siu CW, Oh SK, Tse HF. Electrical stimulation promotes maturation of cardiomyocytes derived from human embryonic stem cells. J Cardiovasc Transl Res. 2013;6(6):989–99.
Ebert A, Joshi AU, Andorf S, Dai Y, Sampathkumar S, Chen H, Li Y, Garg P, Toischer K, Hasenfuss G, et al. Proteasome-dependent regulation of distinct metabolic states during long-term culture of human iPSC-derived cardiomyocytes. Circ Res. 2019;125(1):90–103.
Ribeiro MC, Tertoolen LG, Guadix JA, Bellin M, Kosmidis G, D’Aniello C, Monshouwer-Kloots J, Goumans MJ, Wang YL, Feinberg AW, et al. Functional maturation of human pluripotent stem cell derived cardiomyocytes in vitro–correlation between contraction force and electrophysiology. Biomaterials. 2015;51:138–50.
van den Berg CW, Okawa S, de Sousa C, Lopes SM, van Iperen L, Passier R, Braam SR, Tertoolen LG, del Sol A, Davis RP, Mummery CL. Transcriptome of human foetal heart compared with cardiomyocytes from pluripotent stem cells. Development. 2015;142(18):3231–8.
Small EM, Olson EN. Pervasive roles of microRNAs in cardiovascular biology. Nature. 2011;469(7330):336–42.
Lee DS, Chen JH, Lundy DJ, Liu CH, Hwang SM, Pabon L, Shieh RC, Chen CC, Wu SN, Yan YT, et al. Defined MicroRNAs induce aspects of maturation in mouse and human embryonic-stem-cell-derived cardiomyocytes. Cell Rep. 2015;12(12):1960–7.
White MC, Pang L, Yang X. MicroRNA-mediated maturation of human pluripotent stem cell-derived cardiomyocytes: Towards a better model for cardiotoxicity? Food Chem Toxicol. 2016;98(Pt A):17–24.
Feyen DAM, McKeithan WL, Bruyneel AAN, Spiering S, Hormann L, Ulmer B, Zhang H, Briganti F, Schweizer M, Hegyi B, et al. Metabolic maturation media improve physiological function of human iPSC-derived cardiomyocytes. Cell Rep. 2020;32(3):107925.
Correia C, Koshkin A, Duarte P, Hu D, Teixeira A, Domian I, Serra M, Alves PM. Distinct carbon sources affect structural and functional maturation of cardiomyocytes derived from human pluripotent stem cells. Sci Rep. 2017;7(1):8590.
Knight WE, Cao Y, Lin YH, Chi C, Bai B, Sparagna GC, Zhao Y, Du Y, Londono P, Reisz JA, et al. Maturation of pluripotent stem cell-derived cardiomyocytes enables modeling of human hypertrophic cardiomyopathy. Stem Cell Rep. 2021;16(3):519–33.
Gupta MK, Walthall JM, Venkataraman R, Crowder SW, Jung DK, Yu SS, Feaster TK, Wang X, Giorgio TD, Hong CC, et al. Combinatorial polymer electrospun matrices promote physiologically-relevant cardiomyogenic stem cell differentiation. PLoS ONE. 2011;6(12):e28935.
Chun YW, Balikov DA, Feaster TK, Williams CH, Sheng CC, Lee JB, Boire TC, Neely MD, Bellan LM, Ess KC, et al. Combinatorial polymer matrices enhance in vitro maturation of human induced pluripotent stem cell-derived cardiomyocytes. Biomaterials. 2015;67:52–64.
Feaster TK, Cadar AG, Wang L, Williams CH, Chun YW, Hempel JE, Bloodworth N, Merryman WD, Lim CC, Wu JC, et al. Matrigel mattress: a method for the generation of single contracting human-induced pluripotent stem cell-derived cardiomyocytes. Circ Res. 2015;117(12):995–1000.
Chaicharoenaudomrung N, Kunhorm P, Noisa P. Three-dimensional cell culture systems as an in vitro platform for cancer and stem cell modeling. World J Stem Cells. 2019;11(12):1065–83.
Eder A, Vollert I, Hansen A, Eschenhagen T. Human engineered heart tissue as a model system for drug testing. Adv Drug Deliv Rev. 2016;96:214–24.
Al-Haque S, Miklas JW, Feric N, Chiu LL, Chen WL, Simmons CA, Radisic M. Hydrogel substrate stiffness and topography interact to induce contact guidance in cardiac fibroblasts. Macromol Biosci. 2012;12(10):1342–53.
Silbernagel N, Korner A, Balitzki J, Jaggy M, Bertels S, Richter B, Hippler M, Hellwig A, Hecker M, Bastmeyer M, et al. Shaping the heart: Structural and functional maturation of iPSC-cardiomyocytes in 3D-micro-scaffolds. Biomaterials. 2020;227:119551.
Tu C, Chao BS, Wu JC. Strategies for improving the maturity of human induced pluripotent stem cell-derived cardiomyocytes. Circ Res. 2018;123(5):512–4.
Ronaldson-Bouchard K, Ma SP, Yeager K, Chen T, Song L, Sirabella D, Morikawa K, Teles D, Yazawa M, Vunjak-Novakovic G. Advanced maturation of human cardiac tissue grown from pluripotent stem cells. Nature. 2018;556(7700):239–43.
Ruan JL, Tulloch NL, Razumova MV, Saiget M, Muskheli V, Pabon L, Reinecke H, Regnier M, Murry CE. Mechanical stress conditioning and electrical stimulation promote contractility and force maturation of induced pluripotent stem cell-derived human cardiac tissue. Circulation. 2016;134(20):1557–67.
Sewanan LR, Campbell SG. Modelling sarcomeric cardiomyopathies with human cardiomyocytes derived from induced pluripotent stem cells. J Physiol. 2020;598(14):2909–22.
Tanaka A, Yuasa S, Mearini G, Egashira T, Seki T, Kodaira M, Kusumoto D, Kuroda Y, Okata S, Suzuki T, et al. Endothelin-1 induces myofibrillar disarray and contractile vector variability in hypertrophic cardiomyopathy-induced pluripotent stem cell-derived cardiomyocytes. J Am Heart Assoc. 2014;3(6):e001263.
Dambrot C, Braam SR, Tertoolen LG, Birket M, Atsma DE, Mummery CL. Serum supplemented culture medium masks hypertrophic phenotypes in human pluripotent stem cell derived cardiomyocytes. J Cell Mol Med. 2014;18(8):1509–18.
Han L, Li Y, Tchao J, Kaplan AD, Lin B, Li Y, Mich-Basso J, Lis A, Hassan N, London B, et al. Study familial hypertrophic cardiomyopathy using patient-specific induced pluripotent stem cells. Cardiovasc Res. 2014;104(2):258–69.
Birket MJ, Ribeiro MC, Kosmidis G, Ward D, Leitoguinho AR, van de Pol V, Dambrot C, Devalla HD, Davis RP, Mastroberardino PG, et al. Contractile defect caused by mutation in MYBPC3 revealed under conditions optimized for human PSC-cardiomyocyte function. Cell Rep. 2015;13(4):733–45.
Ojala M, Prajapati C, Polonen RP, Rajala K, Pekkanen-Mattila M, Rasku J, Larsson K, Aalto-Setala K. Mutation-specific phenotypes in hiPSC-derived cardiomyocytes carrying either myosin-binding protein C or alpha-tropomyosin mutation for hypertrophic cardiomyopathy. Stem Cells Int. 2016;2016:1684792.
Pioner JM, Racca AW, Klaiman JM, Yang KC, Guan X, Pabon L, Muskheli V, Zaunbrecher R, Macadangdang J, Jeong MY, et al. Isolation and mechanical measurements of myofibrils from human induced pluripotent stem cell-derived cardiomyocytes. Stem Cell Rep. 2016;6(6):885–96.
Ross SB, Fraser ST, Nowak N, Semsarian C. Generation of induced pluripotent stem cells (iPSCs) from a hypertrophic cardiomyopathy patient with the pathogenic variant p.Val698Ala in beta-myosin heavy chain (MYH7) gene. Stem Cell Res. 2017;20:88–90.
Prondzynski M, Kramer E, Laufer SD, Shibamiya A, Pless O, Flenner F, Muller OJ, Munch J, Redwood C, Hansen A, et al. Evaluation of MYBPC3 trans-splicing and gene replacement as therapeutic options in human iPSC-derived cardiomyocytes. Mol Ther Nucleic Acids. 2017;7:475–86.
Wang L, Kryshtal DO, Kim K, Parikh S, Cadar AG, Bersell KR, He H, Pinto JR, Knollmann BC. Myofilament calcium-buffering dependent action potential triangulation in human-induced pluripotent stem cell model of hypertrophic cardiomyopathy. J Am Coll Cardiol. 2017;70(20):2600–2.
Viswanathan SK, Puckelwartz MJ, Mehta A, et al. Association of cardiomyopathy with MYBPC3 D389V and MYBPC3Delta25bpIntronic deletion in South Asian descendants. JAMA Cardiol. 2018;3(6):481–8.
Li S, Pan H, Tan C, et al. Mitochondrial dysfunctions contribute to hypertrophic cardiomyopathy in patient iPSC-derived cardiomyocytes with MT-RNR2 mutation. Stem Cell Rep. 2018;10(3):808–21.
Ma N, Zhang JZ, Itzhaki I, et al. Determining the pathogenicity of a genomic variant of uncertain significance using CRISPR/Cas9 and human-induced pluripotent stem cells. Circulation. 2018;138(23):2666–81.
Wang L, Kim K, Parikh S, Cadar AG, Bersell KR, He H, Pinto JR, Kryshtal DO, Knollmann BC. Hypertrophic cardiomyopathy-linked mutation in troponin T causes myofibrillar disarray and pro-arrhythmic action potential changes in human iPSC cardiomyocytes. J Mol Cell Cardiol. 2018;114:320–7.
Mosqueira D, Mannhardt I, Bhagwan JR, Lis-Slimak K, Katili P, Scott E, Hassan M, Prondzynski M, Harmer SC, Tinker A, et al. CRISPR/Cas9 editing in human pluripotent stem cell-cardiomyocytes highlights arrhythmias, hypocontractility, and energy depletion as potential therapeutic targets for hypertrophic cardiomyopathy. Eur Heart J. 2018;39(43):3879–92.
Prajapati C, Ojala M, Aalto-Setala K. Divergent effects of adrenaline in human induced pluripotent stem cell-derived cardiomyocytes obtained from hypertrophic cardiomyopathy. Dis Model Mech. 2018;11(2):dmm032896.
Chang ACY, Chang ACH, Kirillova A, et al. Telomere shortening is a hallmark of genetic cardiomyopathies. Proc Natl Acad Sci U S A. 2018;115(37):9276–81.
Yang KC, Breitbart A, De Lange WJ, et al. Novel adult-onset systolic cardiomyopathy due to MYH7 E848G mutation in patient-derived induced pluripotent stem cells. JACC Basic Transl Sci. 2018;3(6):728–40.
Smith JGW, Owen T, Bhagwan JR, Mosqueira D, Scott E, Mannhardt I, Patel A, Barriales-Villa R, Monserrat L, Hansen A, et al. Isogenic pairs of hiPSC-CMs with hypertrophic cardiomyopathy/LVNC-associated ACTC1 E99K mutation unveil differential functional deficits. Stem Cell Rep. 2018;11(5):1226–43.
Cohn R, Thakar K, Lowe A, Ladha FA, Pettinato AM, Romano R, Meredith E, Chen YS, Atamanuk K, Huey BD, et al. A contraction stress model of hypertrophic cardiomyopathy due to sarcomere mutations. Stem Cell Rep. 2019;12(1):71–83.
Seeger T, Shrestha R, Lam CK, Chen C, McKeithan WL, Lau E, Wnorowski A, McMullen G, Greenhaw M, Lee J, et al. A premature termination codon mutation in MYBPC3 causes hypertrophic cardiomyopathy via chronic activation of nonsense-mediated decay. Circulation. 2019;139(6):799–811.
Prondzynski M, Lemoine MD, Zech AT, Horvath A, Di Mauro V, Koivumaki JT, Kresin N, Busch J, Krause T, Kramer E, et al. Disease modeling of a mutation in alpha-actinin 2 guides clinical therapy in hypertrophic cardiomyopathy. EMBO Mol Med. 2019;11(12):e11115.
Zhou W, Bos JM, Ye D, et al. Induced pluripotent stem cell-derived cardiomyocytes from a patient with MYL2-R58Q-mediated apical hypertrophic cardiomyopathy show hypertrophy, myofibrillar disarray, and calcium perturbations. J Cardiovasc Transl Res. 2019;12(5):394–403.
Bhagwan JR, Mosqueira D, Chairez-Cantu K, Mannhardt I, Bodbin SE, Bakar M, Smith JGW, Denning C. Isogenic models of hypertrophic cardiomyopathy unveil differential phenotypes and mechanism-driven therapeutics. J Mol Cell Cardiol. 2020;145:43–53.
Dainis A, Zaleta-Rivera K, Ribeiro A, Chang ACH, Shang C, Lan F, Burridge PW, Liu WR, Wu JC, Chang ACY, et al. Silencing of MYH7 ameliorates disease phenotypes in human iPSC-cardiomyocytes. Physiol Genomics. 2020;52(7):293–303.
Jia WW, Lu JZ, Zhang L, et al. An induced pluripotent stem cell line (EHTJUi003-A) generated from a neonate with c.1377delC mutation in the gene MYBPC3 causing hypertrophic cardiomyopathy. Stem Cell Res. 2021;53:102328.
Ramachandra CJA, Kp MMJ, Chua J, Hernandez-Resendiz S, Liehn EA, Knoll R, Gan LM, Michaelsson E, Jonsson MKB, Ryden-Markinhuhta K, et al. Inhibiting cardiac myeloperoxidase alleviates the relaxation defect in hypertrophic cardiomyocytes. Cardiovasc Res. 2022;118(2):517–30.
Bensley JG, De Matteo R, Harding R, Black MJ. Three-dimensional direct measurement of cardiomyocyte volume, nuclearity, and ploidy in thick histological sections. Sci Rep. 2016;6:23756.
Ho CY, Day SM, Colan SD, Russell MW, Towbin JA, Sherrid MV, Canter CE, Jefferies JL, Murphy AM, Cirino AL, et al. The burden of early phenotypes and the influence of wall thickness in hypertrophic cardiomyopathy mutation carriers: findings from the HCMNet study. JAMA Cardiol. 2017;2(4):419–28.
Bers DM. Calcium cycling and signaling in cardiac myocytes. Annu Rev Physiol. 2008;70:23–49.
Backx PH, Gao WD, Azan-Backx MD, Marban E. The relationship between contractile force and intracellular [Ca2+] in intact rat cardiac trabeculae. J Gen Physiol. 1995;105(1):1–19.
Keller DI, Coirault C, Rau T, Cheav T, Weyand M, Amann K, Lecarpentier Y, Richard P, Eschenhagen T, Carrier L. Human homozygous R403W mutant cardiac myosin presents disproportionate enhancement of mechanical and enzymatic properties. J Mol Cell Cardiol. 2004;36(3):355–62.
Witjas-Paalberends ER, Guclu A, Germans T, Knaapen P, Harms HJ, Vermeer AM, Christiaans I, Wilde AA, Dos Remedios C, Lammertsma AA, et al. Gene-specific increase in the energetic cost of contraction in hypertrophic cardiomyopathy caused by thick filament mutations. Cardiovasc Res. 2014;103(2):248–57.
Toepfer CN, Garfinkel AC, Venturini G, Wakimoto H, Repetti G, Alamo L, Sharma A, Agarwal R, Ewoldt JF, Cloonan P, et al. Myosin sequestration regulates sarcomere function, cardiomyocyte energetics, and metabolism, informing the pathogenesis of hypertrophic cardiomyopathy. Circulation. 2020;141(10):828–42.
Gilligan DM, Chan WL, Joshi J, Clarke P, Fletcher A, Krikler S, Oakley CM. A double-blind, placebo-controlled crossover trial of nadolol and verapamil in mild and moderately symptomatic hypertrophic cardiomyopathy. J Am Coll Cardiol. 1993;21(7):1672–9.
Olivotto I, Camici PG, Merlini PA, Rapezzi C, Patten M, Climent V, Sinagra G, Tomberli B, Marin F, Ehlermann P, et al. Efficacy of ranolazine in patients with symptomatic hypertrophic cardiomyopathy: the RESTYLE-HCM randomized, double-blind, placebo-controlled study. Circ Heart Fail. 2018;11(1):e004124.
Sharma A, McKeithan WL, Serrano R, Kitani T, Burridge PW, Del Alamo JC, Mercola M, Wu JC. Use of human induced pluripotent stem cell-derived cardiomyocytes to assess drug cardiotoxicity. Nat Protoc. 2018;13(12):3018–41.
Burridge PW, Li YF, Matsa E, Wu H, Ong SG, Sharma A, Holmstrom A, Chang AC, Coronado MJ, Ebert AD, et al. Human induced pluripotent stem cell-derived cardiomyocytes recapitulate the predilection of breast cancer patients to doxorubicin-induced cardiotoxicity. Nat Med. 2016;22(5):547–56.
Chaudhari U, Nemade H, Wagh V, Gaspar JA, Ellis JK, Srinivasan SP, Spitkovski D, Nguemo F, Louisse J, Bremer S, et al. Identification of genomic biomarkers for anthracycline-induced cardiotoxicity in human iPSC-derived cardiomyocytes: an in vitro repeated exposure toxicity approach for safety assessment. Arch Toxicol. 2016;90(11):2763–77.
Robinton DA, Daley GQ. The promise of induced pluripotent stem cells in research and therapy. Nature. 2012;481(7381):295–305.
Bussmann LH, Schubert A, Vu Manh TP, De Andres L, Desbordes SC, Parra M, Zimmermann T, Rapino F, Rodriguez-Ubreva J, Ballestar E, et al. A robust and highly efficient immune cell reprogramming system. Cell Stem Cell. 2009;5(5):554–66.
Hochedlinger K, Jaenisch R. Induced pluripotency and epigenetic reprogramming. Cold Spring Harb Perspect Biol. 2015;7(12):a019448.
Marchetto MC, Yeo GW, Kainohana O, Marsala M, Gage FH, Muotri AR. Transcriptional signature and memory retention of human-induced pluripotent stem cells. PLoS ONE. 2009;4(9):e7076.
Kim K, Doi A, Wen B, Ng K, Zhao R, Cahan P, Kim J, Aryee MJ, Ji H, Ehrlich LI, et al. Epigenetic memory in induced pluripotent stem cells. Nature. 2010;467(7313):285–90.
Lister R, Pelizzola M, Kida YS, Hawkins RD, Nery JR, Hon G, Antosiewicz-Bourget J, O’Malley R, Castanon R, Klugman S, et al. Hotspots of aberrant epigenomic reprogramming in human induced pluripotent stem cells. Nature. 2011;471(7336):68–73.
Ohi Y, Qin H, Hong C, Blouin L, Polo JM, Guo T, Qi Z, Downey SL, Manos PD, Rossi DJ, et al. Incomplete DNA methylation underlies a transcriptional memory of somatic cells in human iPS cells. Nat Cell Biol. 2011;13(5):541–9.
Tang C, Lee AS, Volkmer JP, Sahoo D, Nag D, Mosley AR, Inlay MA, Ardehali R, Chavez SL, Pera RR, et al. An antibody against SSEA-5 glycan on human pluripotent stem cells enables removal of teratoma-forming cells. Nat Biotechnol. 2011;29(9):829–34.
Ben-David U, Gan QF, Golan-Lev T, Arora P, Yanuka O, Oren YS, Leikin-Frenkel A, Graf M, Garippa R, Boehringer M, et al. Selective elimination of human pluripotent stem cells by an oleate synthesis inhibitor discovered in a high-throughput screen. Cell Stem Cell. 2013;12(2):167–79.
Lee AS, Tang C, Rao MS, Weissman IL, Wu JC. Tumorigenicity as a clinical hurdle for pluripotent stem cell therapies. Nat Med. 2013;19(8):998–1004.
Schlaeger TM, Daheron L, Brickler TR, Entwisle S, Chan K, Cianci A, DeVine A, Ettenger A, Fitzgerald K, Godfrey M, et al. A comparison of non-integrating reprogramming methods. Nat Biotechnol. 2015;33(1):58–63.
Youssef AA, Ross EG, Bolli R, Pepine CJ, Leeper NJ, Yang PC. The promise and challenge of induced pluripotent stem cells for cardiovascular applications. JACC Basic Transl Sci. 2016;1(6):510–23.
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Li, J., Feng, X. & Wei, X. Modeling hypertrophic cardiomyopathy with human cardiomyocytes derived from induced pluripotent stem cells. Stem Cell Res Ther 13, 232 (2022). https://doi.org/10.1186/s13287-022-02905-0
- Hypertrophic cardiomyopathy
- Induced pluripotent stem cells
- iPSC-derived cardiomyocytes
- Disease modeling