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Branco, M. Human Pluripotent Stem Cells and 3D Cardiac Microtissues. Encyclopedia. Available online: https://encyclopedia.pub/entry/17450 (accessed on 20 June 2024).
Branco M. Human Pluripotent Stem Cells and 3D Cardiac Microtissues. Encyclopedia. Available at: https://encyclopedia.pub/entry/17450. Accessed June 20, 2024.
Branco, Mariana. "Human Pluripotent Stem Cells and 3D Cardiac Microtissues" Encyclopedia, https://encyclopedia.pub/entry/17450 (accessed June 20, 2024).
Branco, M. (2021, December 22). Human Pluripotent Stem Cells and 3D Cardiac Microtissues. In Encyclopedia. https://encyclopedia.pub/entry/17450
Branco, Mariana. "Human Pluripotent Stem Cells and 3D Cardiac Microtissues." Encyclopedia. Web. 22 December, 2021.
Human Pluripotent Stem Cells and 3D Cardiac Microtissues
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Cardiovascular disorders (CVDs) are the number one cause of death globally, according to the World Health Organization, being responsible for 17.9 million deaths in 2016, which represented 31% of all global deaths. Human pluripotent stem cells (hPSCs) have aroused attention as a powerful source of cardiac cells that could help to mitigate some of these problems, namely (1) the identification of new mechanisms of action in different cardiac disorders; (2) improving the reliability of cardiotoxicity side effect detection in newly developed compounds; and (3) providing a source of cardiac cells for the development of new regenerative medicine-based therapies.

human pluripotent stem cells (hPSCs) hPSC-derived cardiac cells 3D cardiomyocyte (CM) differentiation 3D engineered cardiac microtissues (MTs) engineered heart tissues (EHT) drug screening platforms cardiotoxicity cardiac disease modeling

1. Introduction

The adult heart is a four-chamber organ, comprising the left and right atria, and the left and right ventricles, that is delimited by a heart wall composed of three different cell layers, the myocardium, the epicardium and the endocardium. The endocardium is the inner endothelial layer of the heart, and the epicardium is the outer epithelial layer that covers the myocardium. The main function of the heart is pumping blood to the entire body. Heart contraction is determined by the cardiac conduction system, which is responsible for the generation and propagation of electrical stimuli to the working myocardium. The cardiac conduction system of the heart consists of (1) sinoatrial node (SAN) cardiomyocytes (CMs), also known as pacemaker cells, which are responsible for the generation of the electric impulse, (2) atrioventricular node (AVN) CMs, and (3) the impulse-propagating His–Purkinje system, which is responsible for the conduction of electric stimuli towards the working myocardium (Figure 1). CMs comprise the majority of the cardiac tissue volume, around 75%, but they account for only around 30% of the total number of cells present in the heart [1][2][3]. The majority of the remaining cells are non-myocytes, predominantly vascular endothelial cells (ECs), which account for 60% of the non-myocyte cell population in terms of cell number, followed by cardiac fibroblasts (CFs), which represent 15% of the non-myocyte cell population [2].
Figure 1. Schematic representation of the human adult heart. Highlighted is the cardiac condition system composed by sinoatrial node (SAN), atrioventricular node (AVN), Left and Right Bundle Branches (LBB and RBB); and heart wall, composed of three different cell layers, the myocardium, the epicardium and the endocardium.
Cardiovascular disorders (CVDs) are the number one cause of death globally, according to the World Health Organization, being responsible for 17.9 million deaths in 2016, which represented 31% of all global deaths [4]. In addition to CVDs, cardiotoxicity linked to newly developed drugs is another serious problem in the cardiac field. Although cardiotoxicity is an important focus of attention during the pre-clinical stage of the pipeline for the development of new drugs, the lack of more reliable and predictive models compromises the accuracy of toxicity detection. In this context, human pluripotent stem cells (hPSCs) have aroused attention as a powerful source of cardiac cells that could help to mitigate some of these problems, namely (1) the identification of new mechanisms of action in different cardiac disorders; (2) improving the reliability of cardiotoxicity side effect detection in newly developed compounds; and (3) providing a source of cardiac cells for the development of new regenerative medicine-based therapies.
Due to the increasing amount of knowledge concerning the embryonic development of the heart, it is now possible to efficiently and robustly direct the differentiation of hPSCs towards CMs and other non-myocyte cardiac cells. These protocols rely on the modulation of important signaling pathways, using growth factors and/or small molecules, in a specific temporal pattern. Despite the major improvements that have been achieved in the field of CM differentiation from hPSCs, these cells still lack the structural, functional and metabolic maturation observed in adult CMs. Different strategies have been implemented to improve hPSC-derived CM maturation in vitro, including long-term culture, electric stimulation, mechanic load, culture with biomaterials, metabolic maturation and co-culture with other cardiac cells (reviewed in [5][6]).

2. Cardiovascular Lineages Specification from hPSCs—Lessons from In Vivo Heart Development

The identification of the main steps that occur during the process of heart development during embryogenesis and the knowledge of how the different cell populations are specified have provided the basis for the establishment of directed differentiation protocols to generate the entire repertoire of cardiac cells from hPSCs. Through the step-wise indirect or direct manipulation of different signaling pathways, it has been possible to guide the process of differentiation of hPSCs towards the different cardiac cell fates in vitro.

2.1. Cardiomyocytes

In the past few years, major improvements have been made in the development of directed differentiation methods to generate CMs from hPSCs. Each step involved in in vitro CM differentiation, including the early stages of mid/posterior primitive streak (PS) establishment and lateral/cardiac mesoderm specification, has been deeply studied. Generally speaking, it is known that the combination of Activin/Nodal, BMP, FGF and Wnt activation allows mesoderm induction from hPSCs, and that, later on, the inhibition of Wnt signaling is critical to successfully generate cardiac mesoderm and hPSC-CMs. BMP and Wnt signaling converge in activating cardiac mesoderm specification, whereas FGF and Activin A signaling are additional important pathways that should be active by direct exogenous stimulation or indirectly through Wnt and BMP signal modulation [7][8][9]. The manipulation of these signals using exogenous growth factors and/or small molecules in a specific concentration and at a specific time point has allowed the efficient generation of hPSC-CMs [10][11][12][13]. In addition to the direct exogenous manipulation of the mentioned signaling pathways, through the use of specific agonist and antagonist molecules, other culture parameters can be exogenously controlled to drive cardiac commitment. This is the case of the protocol relying solely on the temporal modulation of Wnt signaling by small molecules in a very specific and controlled manner by firstly activating that pathway for mesendoderm induction followed by a step of Wnt pathway inhibition for cardiac lineage specification [14]. In fact, the success of this differentiation process is dependent on additional parameters that should be taken into consideration, such as (1) the cell confluence at the beginning of the differentiation and the cell density (cells/mL), which impact the concentration of paracrine signals released to the culture medium, (2) the timing of Wnt activation and inhibition, and (3) the concentration of the small molecules [15][16][17][18][19][20]. Since the establishment of this protocol, several adaptations were reported, including the transition to chemically defined and xeno-free conditions [20][21]. Moreover, the impact of Wnt signaling activation duration and the timing for Wnt signaling inhibition were studied, envisaging the optimization of this differentiation process [15][16][20].
Two main limitations have been identified in CMs generated using the aforementioned protocols, namely (1) hPSC-CMs are often relatively immature compared with adult CMs and (2) the obtained hPSC-CMs are often a mixed population of different subtypes of CMs, although most of the time they showed a ventricular-like signature after prolonged time in culture [14][20]. Envisaging the future applicability of these hPSC-derived CMs in clinical applications as well as disease modeling or drug screening studies, new protocols have emerged to specifically generate different subtypes of CMs and also to identify cell surface markers that could help in the selection and purification of those cells.

2.2. Vascular Cardiac Cells

The protocols that have been described to obtain vascular cells in vitro from hPSCs (Figure 2A–C) are normally organized in three main steps, the first comprising cardiac mesoderm induction, followed by vascular progenitor cell specification and proliferation, and finally by EC or pericyte/vascular SMC (vSMC) commitment and expansion. Since the first attempts to generate cardiovascular progenitor cells, and afterwards vascular cells, were not highly efficient, MACS or FACS-based purification methods were normally integrated with the differentiation protocols.
Figure 2. Current in vitro strategies to generate non-myocyte cardiac cells from hPSCs and respective molecular markers. Protocols for generation of (A) vascular progenitor cells and endothelial cells (ECs); (B) cardiac fibroblasts (CFs) and vascular smooth muscle cells (vSMCs) from cardiac mesoderm progenitors; (C) CFs and vSMCs from pro-epicardial-like cells. VECs, venous endothelial cells; AECs, arterial endothelial cells; P, passage; AA, ascorbic acid; DMEM/F12+SP, DMEM/F12 + chemically defined lipid concentrate + ITS + Glutamax + monothiol glycerol + AA; ECGM-MV2, Endothelial Cell Growth Medium MV2, PromoCell; ESFM, Endothelial Serum-Free Medium, GIBCO; ECGM-2, Endothelial Cell Growth Medium 2, PromoCell.

2.3. Cardiac Fibroblasts (CFs)

It has been described that CF cells can arise from different cell population sources including (1) the epicardium, which is described as the major source of resident CFs present in the myocardium (~80%), (2) the embryonic endothelium, localized in particular regions of the ventricular septum and the right ventricle (~10–20%), and (3) the neural crest lineages, which represent a small fraction [22][23]. CFs present a highly variable morphology in culture and are mostly sheet or spindle-like shaped. Although there is not a specific marker for the identification of cardiac fibroblasts, a combination of cytoskeletal, cell membrane, nuclear and extracellular markers has been used to identify these cells. Periostin (POSTN), fibronectin, and collagen types I, III, V and VI are the main structural components produced by CFs present in cardiac tissue. Moreover, vimentin has been used as a cytoskeletal marker, TCF21 as a nuclear marker and PDGFRα, DDR2 and CD90 as cell membrane markers.
Cardiac fibroblasts are normally present as a minority among the overall cell population generated in optimized hPSC-CMs differentiation protocols. Thus, differentiation protocols have been developed to specifically derive CFs from hPSCs. In a recent study from Zhang and colleagues [24], a protocol for the generation of CFs from SHF progenitors (ISL1+/CXCR4+) was reported. After generating mesoderm progenitor cells (Bry+/CD90+) by Wnt signaling activation, this population was exposed to FGF2 for 18 days, thus generating more than 70% CFs (TE-7+) (Figure 2B). Indirect differentiation protocols, which comprise a pre-differentiation stage of hPSCs into epicardial cells followed by a stage of differentiation into CFs, have been also explored to generate these types of cardiac cells [25][26][27][28] (Figure 2C). For this purpose, the hPSC-derived epicardial cells (>90% WT1+cells) are exposed to FGF2 for 6–14 days to generate an almost pure population of POSTN+ CF-like cells.
To assess CF function, the main evaluated parameter is the capacity of the cells for producing ECM proteins, particularly fibronectin and collagen I. Additionally, the capacity of CFs to convert into myofibroblasts (SMA+ population with prominent fibers) upon injury can be evaluated, which has been recreated in vitro through TGFβ1 stimulation [24].

2.4. Epicardial Cells

In vivo, the epicardium derives from a transient cluster of mesothelial cells, the proepicardium (PE) organ (PEO), which develops at the base of the venous inflow tract of the heart tube and in close proximity with the liver. PE cells migrate, attach and spread along the outer surface of the developing myocardium, ultimately giving rise to an epithelial sheet, the epicardium. The most common markers used to identify PE cells and epicardial cells are WT1, TCF21, TBX18, SCX and SEMA3D [29][30][31].
Witty and colleagues reported one of the first protocols for efficient differentiation of epicardial-like cells from hPSC [25]. They demonstrated that the combined activation of BMP and WNT signaling in a KDR+/PDGFRA+ mesoderm cardiac progenitor population promotes the generation of more than 80% WT1+/TBX18+ epicardial-like cells (Figure 2C). After several passages in culture, these cells showed characteristic properties of the in vivo epicardium, namely the formation of epithelial-like sheets with tight junctions expressing ZO1 and exhibiting the expression of ALDH1A2, which is correlated with aldehyde dehydrogenase activity, an indication of their ability to synthesize RAc. An alternative method to obtain epicardial-like cells was reported by Lyer et al. [28], who proposed the combination of WNT, BMP and RAc signaling activation in lateral plate mesoderm progenitor cells (KDR+/ISL1+) towards the generation of more than 60% WT1+ cells after 15 days of differentiation. These epicardial-like cells showed epithelial cell morphology and expression of the epicardial markers TBX18, WT1 and TCF21. At the same time, Boa and colleagues [26] showed that the temporal modulation of the canonical WNT signaling via small molecules was sufficient for epicardial induction from hPSCs in chemically defined, xeno-free conditions. The activation of WNT signaling for 2 days in an NKX2.5+/ISL1+ cardiac progenitor population was sufficient to generate more than 90% WT1+ epicardial-like cells. They also demonstrated that TGF-β signaling inhibition allows the long-term maintenance of self-renewing epicardial cells. Zhao and colleagues [27] reinforced the idea that the sole combination of RAc and WNT signaling pathway activation, in an ISL1+/KDR+ cardiac progenitor population of cells, was sufficient to generate more than 90% WT1+-epicardial-like cells.
One of the major features of pro-epicardial cells in vivo is their capacity for migration and spreading, surrounding the myocardium of the developing heart. After the establishment of this layer, some cells undergo epithelial-mesenchymal transition (EMT) to differentiate into vSMCs and CFs. Thus, the potential of these cells to undergo EMT has been the main functional test for epicardial cells in vitro [25][28].

3. Impact of 3D Environment on Cardiomyocyte Differentiation of hPSCs

The first attempts to perform CM differentiation from hPSCs in a 3D environment were based on the embryoid body (EB) differentiation method. Despite the low efficiency of this process, due to the lack of specific cues to induce cardiac differentiation, this system allowed the acquisition of valuable biological information that later on was taken into consideration for the development of more robust and efficient 3D-based directed differentiation protocols.
Taking advantage of micropatterned and forced aggregation platforms to generate size-controlled 3D aggregates of hPSCs, different studies have been performed to evaluate the impact of culturing hPSCs in a 3D environment in comparison with 2D culture. Several reports have shown that EB differentiation was influenced by hPSC aggregate size at the beginning of the process [32][33][34][35]. hPSC aggregate diameters in the range of 250–350 µm were described as the optimal size to maximize cardiac gene expression and thus cardiac differentiation efficiency. Additionally, it was also stated that this controlled method for aggregate generation increased hPSC aggregate homogeneity at the beginning of differentiation, which resulted in a higher reproducibility and less variability between runs, compared to standard enzymatic methods to form EBs [35].
Although the size of hPSC aggregates had been shown to influence the differentiation outcome for different lineages, the molecular factors behind that effect were not clear at the beginning. To achieve a deeper understanding of these effects, different studies were performed [36][37][38][39]. Using the EB differentiation protocol, Azarin and co-workers [39] suggested that cell-to-cell interactions experienced by the hPSCs in the undifferentiated state in a 3D environment through, for example, E-cadherin interactions could be responsible for the modulation of different signaling pathways that could later impact the differentiation outcome. Particularly, they observed that upon EB differentiation, hPSCs that had been previously cultured in micropatterned platforms (300 × 300 µm) showed an upregulation of the canonical Wnt signaling pathway during the first few days of differentiation when compared with 2D cultured hPSCs, which resulted in a higher expression of genes associated with primitive streak, mesoderm and cardiac lineage commitment. In another study from the same group [38], it was suggested that hPSCs cultured in microwells showed an upregulation of BMP signaling and less transcriptional activity of genes involved in the Activin/Nodal pathway, which they proposed to result in a priming of hPSCs towards differentiation, and consequently influenced the exit from pluripotency and germ lineage specification upon the beginning of differentiation. Through the analysis of important genes involved in cardiac differentiation, they observed that EB differentiation using microwell cultured hPSCs showed a strong peak of expression of Brachyury, a transcription factor that controls PS induction, which was not observed in EB differentiation performed with 2D cultured hPSCs. Similarly, mesendoderm lineage and early cardiogenesis genes, such as ISL1 and NKX2.5, were upregulated in EB differentiation using microwell cultured hPSCs. Moreover, upregulation of the genes responsive to Wnt signaling, such as WNT3A, WNT8A and LEF1, Notch pathway genes such as NOTCH1 and DELTA1, and representative genes from the TGFβ superfamily, such as BMP2, BMP7, NOGGIN and NODAL, were also upregulated in EBs obtained from microwell cultured hPSCs.
The effect of 3D aggregate size on CM differentiation from hPSCs was later on confirmed by Bauwens and colleagues under serum-free conditions and using a directed CM differentiation protocol based on the manipulation of BMP4, Activin A and FGF for mesoderm induction and a second stage of DKK1 and VEGF media supplementation for cardiac mesoderm and CM specification [37]. As the cellular mechanism, they suggested that the aggregate size influences the extension of endoderm layer development during differentiation as the main reason behind the impact of aggregate size on cardiac induction and differentiation efficiency (endoderm-secreted factors). They additionally stated that the control of aggregate size at the beginning of differentiation allows the achievement of consistent and efficient cardiac induction runs.
More recently, other groups have demonstrated the impact of hPSC-aggregate size on CM differentiation when using solely the temporal modulation of Wnt signaling by using a forced aggregation platform [40][41] or a dynamic system [42] for generation of hPSC aggregates. In a recent study, a possible impact, at the transcriptional level, of forced aggregation of hPSCs when compared to 2D cultured hPSCs was revealed [41]. In this study, it was demonstrated that by culturing hPSCs under 3D conditions, in the presence of hPSC expansion medium, for 3 days in a microwell platform (approx. 300 µm in diameter), these cells exhibited a priming for mesendoderm commitment, which further resulted in a faster exit from the pluripotency stage and primitive streak commitment upon cardiac differentiation induction, culminating in a faster CM differentiation progression and maturation when compared with 2D monolayer. It was suggested that these differences could be related not only to the higher degree of cell-to-cell interactions observed in 3D aggregates, but also to the oxygen gradients inside the spheroids, which, all together, culminated in a stabilization of the TGF-β/Nodal pathway, upregulation of the MAPK/JNK/ERK pathway and increased glycolysis metabolism, when compared with 2D-cultured hPSCs.
Different 3D culture platforms have been described to successfully generate CMs from hPSCs. Depending on the purpose, the generation of the 3D aggregates of hPSCs for cardiac differentiation may rely on dynamic systems, including different bioreactor configurations [16][42][43][44][45][46] or static conditions, where forced aggregation platforms, including microwell plates [40][41], U- and V- 96-well plates [18][47] and micropatterned surfaces [48], have been used. Since the pre-differentiation stage and the early differentiation phase have a substantial impact on cardiomyocyte differentiation outcomes, as discussed before, the control of the initial population of 3D-hPSC aggregates in terms of aggregate size and homogeneity, synergistically combined with optimized concentrations of growth factors and/or small molecules to induce the first commitment of hPSCs, is crucial for the success of the differentiation process. The use of microwell plates allows the generation of size controlled and homogenous populations of aggregates only through cell seeding density manipulation (number of cells/aggregates) [40][41]. In the case of dynamic systems, the size of aggregates and, depending on the platform, aggregate homogeneity, can be also controlled by cell seeding density, agitation rate and time in culture [42]. However, the variability in the size of the aggregates is generally higher in this type of dynamic system when compared with forced aggregation platforms, which can comprise the reproducibility between biological runs. Both dynamic and microwell systems have been proven to generate highly pure populations of CMs (>80% of CMs) within 10/16 days of differentiation (Table 1). Although the aforementioned parameters (pre-differentiation period and concentration of small molecules and/or growth factors), highlighted in Table 1, are critical for the success of CM differentiation in a 3D environment, they should be synergistically optimized for the specific platform that is being used. Different hPSC-aggregate sizes at day 0 of differentiation or different concentrations of factors in different culture platforms may result in identical efficiencies. Since there are already robust and efficient systems to generate CMs in a 3D environment, the selection of the best protocol will depend on the final aim. Dynamic systems should be preferred for large-scale production of CMs, for example, for regenerative medicine applications. On the other hand, to produce CMs or 3D cardiac MTs for in vitro applications, protocols that rely on microwell plates may be advantageous, since these are simpler and do not require sophisticated equipment nor specific expertise in bioprocessing and may allow an easier integration with a medium- to high- throughput screening platform.
Table 1. Summary of studies reporting CM differentiation from hPSCs under 3D conditions. The table highlights 3D-CM differentiation platforms in dynamic systems (bioreactors and spinner flasks) and static conditions (microwell plates and ULA attachment plates). Ø, diameter; ULA, Ultra Low Attachment Plate.

Reference

Pre-Differentiation

Differentiation

Platform

Time

Media

Aggregate Ø at D0

Platform

Media

Molecules

Duration

Efficiency

Halloin et al., 2019

Stirred Bioreactor

2 Days

E8

±125 µm

Stirred Bioreactor (500 mL)

CDM3 *1

CHIR

D0–D1

5 µM

10 Days

±1 × 106 CMs/mL

IWP2

D1–D3

5 µM

93 ± 5% CMs

Chen et al., 2015

Spinner Flask

2 Days

StemPro hESC SFM + FGF2

200 ± 20 μm

Spinner Flask (125 mL–1L)

RPMI + B27-INS

D0–D4

CHIR

D0–D1

6/12 µM *2

16 Days

±1 × 106 CMs/mL (1L)

±2 × 106 CMs/mL (500 mL)

RPMI + B27

D4–D16

IWP4

D2–D4

5 µM

>90% CMs

Fonoudi et al., 2015

Stirred Bioreactor

5 Days

DMEM/F12+ FGF2

175 ± 25 µm

Stirred Bioreactor (100 mL)

RPMI + B27

D0–D15

CHIR

D0–D1

12 µM

15 Days

0.8 × 106 CMs/mL

SB + Pur + IWP2

D2–D4

5 µM each

>80% CMs

Branco et al., 2019

AggrewellTM800

3 Days

mTeSR1

±300 µm

AggrewellTM800

D0–D7

RPMI + B27-INS

D0–D7

CHIR

D0–D1

11 µM

12 Days

±20 × 106 CMs/plate

ULA 6-well plate

D0–D12

RPMI + B27

D7–D12

IWP4

D3–D5

5 µM

>85% CMs

Burridge et al., 2011

-

*6

96-V ULA plate

D0–D4

RPMI

D0–D10

*5

10 Days

±0.4 × 106 CMs/plate

96-U ULA plate

D4–D10

>80% CMs

Dahlmann et al., 2013

Agarose Microwell plate

1 Day

FCM *3

400–500 µm *4

(±220 µm (D-3))

ULA 6-well plate

RPMI + B27-INS

D0–D7

CHIR

D0–D1

8 µM

10 Days

*6

ULA 6-well plate

3 Days

RPMI + B27

D7–D10

IWR1

D3–D5

4 µM

Up to 65% CMs

*1 RPMI 1640 (+2 mM Glutamine) + 495 µg/mL Recombinant Human Albumin + 213 µg/mL Ascorbic Acid. *2 Depending on the cell line. *3 DMEM/F12 + GlutaMAX + 20% (v/v) Knockout serum replacement + 1% (v/v) non-essential amino acids + 0.1 mM β mercaptoethanol + 10 ng/mL FGF-2. *4 Determined by bright field image analysis. *5 D0-D2: BMP4 (25 ng/mL); FGF2 (5 ng/mL); PVA (4 mg/mL); h-Insulin (10 µg/mL). D2-D4: HAS (5 mg/mL); 280 µM L-ascorbic acid. D4-foward: h-Insulin (10 µg/mL). *6 Not specified*.

4. Engineering 3D Cardiac Microtissues to Better Mimic the Human Heart Environment

In the in vivo cardiac microenvironment, CMs are organized in a 3D structure, the integrity of which is maintained by ECM produced mainly by CFs, and they are in close proximity with cardiac vascular cells, which play a critical role not only during the first stages of embryonic heart development but also in myocardium structural and functional maturation. In addition to ECM production and remodeling, CFs have been also described to modulate and interfere with the electrical behavior of CMs [49]. One of the main limitations that started to stand out in hPSC-CMs was the lack of structural and functional maturation compared with adult CMs. The development of more complex 3D cardiac MTs in vitro, combining different cardiac cells in a 3D structure, has emerged as an interesting alternative to better mimic the complexity and dynamic network of interactions and signals that are present in human heart tissue [48][50], towards the development of more reliable cardiac models for different applications. In fact, cardiac models in which only CMs are present do not recapitulate the in vivo environment of the heart, where CFs and vascular cells interact with and strongly impact CM behavior.
Different approaches to generating 3D cardiac MTs have been explored and reported in the literature. The two most promising approaches for in vitro applications are (1) hydrogel-based engineered heart tissues (EHT), and (2) cardiac spheroids obtained through self-assembly of cells in a scaffold-free environment (3D multicellular MTs). In both models, different cardiac cells, including ECs and CFs, which can be hPSC-derived or primary cultured cells, are combined with hPSC-derived CMs at a specific ratio. In the case of the EHT models, the composition of the hydrogel and the concentration of the ECM used are also important parameters to ensure tissue structure and functionality [51].

5. In Vitro Applications of hPSC-Derived 3D Cardiac Microtissues (MTs)

The improvements achieved in the field of hPSC-derived cardiac MTs have increased the interest in applying those models to different in vitro applications, such as modeling of cardiac disorders, cardiotoxicity tests and studying the therapeutic effects of developing drugs in the context of diseased phenotypes (Figure 3). Importantly, to increase the applicability of the hPSC-derived cardiac models, namely in the pharmaceutical industry, the integration of those models in medium- to high-throughput screening settings coupled with high content analytical setups has also been a focus of attention. The miniaturization of 3D cardiac MTs is particularly relevant in cardiotoxicity and drug screening applications since normally a considerable number of new compounds are tested at different concentrations and times of exposure, and, in this way, it is possible to maximize the readouts with smaller and more cost-effective setups.
Figure 3. hiPSC-derived 3D cardiac models—applications and challenges. (A) hPSCs, and specifically hiPSCs, are a powerful technology to generate wild-type and patient-specific cardiac models to be applied in drug screening, cardiotoxicity tests and disease modeling assays. (B) The most widely used cardiac models for in vitro applications still present some limitations that can be solved by using different tissue engineering strategies.

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