The epicardium is a mesothelial layer covering the myocardium serving as a progenitor source during cardiac development. The epicardium reactivates upon cardiac injury supporting cardiac repair and regeneration. Fine-tuned balanced signaling regulates cell plasticity and cell-fate decisions of epicardial-derived cells (EPCDs) via epicardial-to-mesenchymal transition (EMT). However, powerful tools to investigate epicardial function, including markers with pivotal roles in developmental signaling, are still lacking. Here, we recapitulated epicardiogenesis using human induced pluripotent stem cells (hiPSCs) and identified type II classical cadherin CDH18 as a biomarker defining lineage specification in human active epicardium. The loss of CDH18 led to the onset of EMT and specific differentiation towards cardiac smooth muscle cells. Furthermore, GATA4 regulated epicardial CDH18 expression. These results highlight the importance of tracing CDH18 expression in hiPSC-derived epicardial cells, providing a model for investigating epicardial function in human development and disease and enabling new possibilities for regenerative medicine.
Cardiovascular diseases are the leading cause of death worldwide. The human heart is incapable of restoring itself and myocardial repair and regeneration processes are still poorly understood. In recent years the epicardium has emerged as a therapeutic target, given its ability to re-activate upon cardiac injury and promote cardiac repair1. The epicardium is a mesothelial layer covering the myocardium and serves as a progenitor source supporting cardiac development, repair, and regeneration2,3,4. Epicardiogenesis is evolutionary conserved and absolutely essential for cardiac development, with failure of epicardial formation being embryonic lethal5,6. After the formation of the proepicardium (PE), a transient cauliflower-like structure at the venous pole of the looping heart, cells of the PE migrate over and cover the looping heart tube completely, subsequently forming the epicardium. The embryonic epicardium is active, epicardial cells are proliferative and have the ability to undergo epicardial epithelial-to-mesenchymal transition (EMT). Those epicardial-derived cells (EPDCs) can proliferate and migrate, invading the underlying myocardium, where they subsequently differentiate into various cardiac cell types like vascular smooth muscle cells (SMC), cardiac fibroblasts (CF), and, to a lesser degree, endothelial cells (EC)7,8,9. The epicardium and EPDCs exhibit extensive developmental plasticity, which is crucial for cardiogenesis, by contributing to coronary vessel formation as well as myocardial growth and maturation1,2,4,10,11. During adulthood, the epicardium enters quiescence and serves as a protective layer. Only upon cardiac injury, the epicardium re-activates and contributes as a quick transient progenitor hub of derivative cells to the cardiovascular system regeneration4,12. This fact provides the epicardium with a pivotal role in mediating regenerative responses. Unlike other species, mammals lack the ability to restore the heart. Nonetheless, re-activation of the epicardium is required for cardiac scar formation as well as coronary vessel growth13. Not only promotes the epicardium myocardial regeneration via paracrine signaling, but it also mediates inflammatory responses14,15. Moreover, engineered patches carrying epicardial follistatin-like 1 (FSTL-1) were able to enhance cardiac regenerative responses16. However, despite ongoing investigation on the epicardium as a therapeutic target tissue, processes governing epicardial development, re-activation and engraftment ability are not well understood, mainly due to lacking comprehension of the fundamental biology behind the epicardium itself, including the expression dynamics of epicardial epitopes mediating tissue responses to homeostasis disruptions and cellular signaling upon cardiac damage.
The epicardium coordinates a complex network of surface remodelers in order to respond to different stimuli driven by organ development and tissue repair11,17. The cell surfaceome describes the whole proteome of surface and transmembrane proteins. During tissue morphogenesis, homeostasis, and regeneration, the surfaceome, particularly surface remodelers, plays an essential role in expansion, migration, and invasion, forming complex cellular structures18. However, cell-surface markers defining functional epicardial cells or regulating epicardial cell-fate decisions have been difficult to identify.
The cadherin family of cell–cell adhesion proteins are important for tissue morphogenesis19. Even though cadherins may have originated to facilitate mechanical cell-cell adhesion, their functions are pleiotropic and have evolved to be imperative for many other aspects, including cell recognition, coordinated cell movements, cell-fate decisions and the maintenance of structural tissue polarity, by controlling diverse signaling pathways20,21,22,23,24,25.
In this study, we identified type II classical cadherin CDH18, formerly known as cadherin 14 (CDH14), as a specific biomarker expressed in the fetal-stage epicardium, defining cellular specification. Human pluripotent stem cells (hPSCs)26,27 and their capacity to differentiate into cardiac cells allow recapitulation of epicardiogenesis in vitro28,29,30,31,32,33,34,35,36. Here we generated human-induced (hi)PSC-derived epicardial-like (EPI) cells. Epicardial identity was associated with sustained expression levels of key epicardial genes and retained CDH18 expression in correlation to GATA4 expression, an essential transcription factor proven to be required to form PE in vivo37. The loss of CDH18 expression led to the activation of cell-fate specific EMT towards SMC differentiation, confirming an important biological function of CDH18, not only to define progressive epicardial lineage specification but also to regulate downstream signaling that drives EPDC derivation. Our study sets a basis for a reproducible model to investigate epicardial function in human cardiac development and disease, ultimately enabling new possibilities in regenerative medicine.
Defined transcriptome of developing epicardial surfaceome revealed the enrichment of CD22 and CDH18 in late-stage populations
To investigate the expression profiles of the epicardium, we compared RNA-Seq dataset of induced epicardial-like cells at day (d) 12 (Epi12) and 48 (Epi48) representing a developmental early- and late-stage respectively and investigated their relationship with the adult quiescent epicardium28. Whole mRNA transcriptome-based principal-component analysis (PCA) demonstrated 84.75% variance between Epi12 and Epi48 populations (Fig. 1a), with the biggest difference observed between Epi12 and adult—quiescent—cells (Supplementary Fig. 1a, b). The active embryonic epicardium is defined by the expression of the transcription factors Wilms’ tumor 1 (WT1) and T-Box 18 (TBX18), followed by aldehyde dehydrogenase 1 member A2 (ALDH1A2), distinguishing it from the myocardium and endocardium, as well as the adult epicardium which lies quiescent (Supplementary Fig. 1c). All three genes were upregulated in correlation to each other over time (Supplementary Fig. 1d, e), confirming the embryonic epicardial identity of the transcriptome profiles. Comparative analysis of genes in correlation to WT1, TBX18 and ALDH1A2, unveiled 2221 common genes in their intersection (Fig. 1b), with 49 genes identified as surface protein-encoding. We also found 825 differentially expressed genes (DEGs) between Epi12 and Epi48 populations (Supplementary Fig. 1f) allowing us to subsequently define a putative transcriptome of epitopes of the active epicardium (Fig. 1c).
Next, we excluded genes that had no significant negative correlation to the endothelial cell markers CDH5 and PECAM1 (Fig. 1d) and selected late-stage upregulated genes (Fig. 1e) that showed a highly positive correlation to epicardial genes (Fig. 1f), obtaining CD22 and CDH18 as candidate cell surface biomarkers in the active epicardium. We then confirmed that both candidates were no longer expressed in adult tissue similar to active epicardial key genes (Fig. 1g and Supplementary Fig. 1g). Next, we verified their absence in two important epicardial derivatives. While CF and SMC markers were upregulated in their respective cellular identities, no mRNA expression of CD22 or CDH18 was detected (Fig. 1h, i). Moreover, we confirmed tissue specificity using independent datasets: Heart CD22 expression ranked #2 after central nervous system (CNS), and heart CDH18 expression ranked #3 after CNS and testis (Supplementary Fig. 1h), two known CDH18-expressing tissues38,39,40. Finally, as an increase of CDH18 is difficult to observe during heart development of mouse embryos due to cellular heterogeneity in the bulk population (Supplementary Fig. 1i), we organotypically cultured explanted epicardial tissue from E14 hearts and documented CDH18 expression in epicardial explants (Supplementary Fig. 1i). We confirmed the epicardial specificity of CDH18 expression in a physiological context of cardiac cells derived from human embryos ranging from different time points of gestation (Fig. 1j, k).
CDH18 but not CD22 is an epicardial biomarker defining progressive lineage specification
To define the expression pattern of CD22 and CDH18, we first recapitulated epicardiogenesis in vitro. We modified previously reported induction methods for hiPSC-derived epicardial-like (EPI) cells28,32,35 to robustly yield WT1+ cells at a higher rate, and long-term culture of cells was achieved by permanent inhibition of transforming growth factor (TGF)-β (Fig. 2a). As epicardial cells and atrial cardiomyocytes share a common progenitor pool during development3, we excluded the presence of any residual myocyte traces in our system, by generating EPI cells from MYH6 reporter line (MYH6-EIP4)41, obtaining no cardiomyocytes (Supplementary Fig. 2a–c). Moreover, the absence of endothelial markers CD31 and CD144 ensured that no cells of endothelial nature were present (Supplementary Fig. 2d–f). EPI cells showed typical epicardial cobblestone-like morphology (Fig. 2b), and key active epicardial genes (WT1 and TBX18) started to be upregulated as early as d12 continuing so until d24 (Fig. 2c). In agreement with previous models, cells did not upregulate ALDH1A2—a gene highly expressed in the epicardium but not PE—until d24, indicating that the cells at d12 represented an early stage during development comparable to PE cells and specified into later stage epicardium at d24 (fetal-like) (Fig. 2c, d). We found all genes expressed in correlation to each other (Supplementary Fig. 3a) and did not observe significant differences in their expressions after d24 (Supplementary Fig. 3b, c), endorsing that cells had completed differentiation into fetal-stage epicardium by d24. We also confirmed WT1 protein expression on this day by immunofluorescence (Supplementary Fig. 3d–f).
We then checked CD22 expression levels during epicardial differentiation. We found CD22 expression increased but failed to detect CD22 protein in d24 EPI cells, suggesting protein translation insufficiency, and therefore invalidating CD22 as a suitable biomarker (Supplementary Fig. 4).
Next, we investigated CDH18 levels and could verify its expression in EPI cells derived from different hiPSC lines (Fig. 2f and Supplementary Fig. 5a). Both mRNA and protein (Fig. 2e–g) levels increased exponentially over time of epicardial differentiation and CDH18 was co-expressed with WT1 in a highly correlated fashion (Fig. 2g, h and Supplementary Fig. 5), implying a potential role during epicardial specification.
Epicardial cells have the potential to transit into EPDCs via EMT and subsequently differentiate into CF and SMC during development and also after re-activation upon cardiac injury. This functionality has also been well reported in vitro by treatment with basic fibroblast growth factor (bFGF) and TGF-β29,31,34. We initiated an EMT response (Supplementary Fig. 6a) and observed changed morphology as well as loss of WT1 and ZO1 indicating loss of epicardial identity and demonstrating the functionality to transition into EPDCs (Fig. 2i). The identity of induced CFs (iCFs) and induced SMCs (iSMCs) was validated by expression of POSTN and αSMA respectively (Fig. 2i). We also sequenced the whole transcriptome of iCFs and iSMCs, verifying distinct cellular identities by PCA (Supplementary Fig. 6b), as well as enrichment of respective gene signatures (Fig. 2j). We then analyzed CDH18 expression in epicardial derivatives from both PE-like d12 and fetal-like d24 EPI cells. CDH18 was lost in iCF and iSMC (Fig. 2k, l and Supplementary Fig. 6c–f), demonstrating that reduced levels of CDH18 correlated to a loss of EPI identity. Loss of CDH18 upon EMT was also confirmed in murine epicardial MEC1 cell line (Supplementary Fig. 6d).
Taken together, our results demonstrate that CDH18 is a specific active epicardial biomarker and suggest a causal role in specification and maintenance of epicardial identity.
Loss of CDH18 triggers cell-fate specific EMT towards SMCs
To explore the biological relevance of CDH18, we next silenced CDH18 gene expression in d24 EPI cells (d24•si18) (Fig. 3a). The downregulation of CDH18 (Fig. 3b–d) led to visible morphological changes with a fraction of cells becoming elongated and losing the cobblestone-like morphology (Fig. 3d, yellow arrowheads) indicating a switch to mesenchymal cells42. Cells that retained epicardial morphology were in close contact, suggesting that cell–cell interactions might play a role in maintaining EPI identity. Similar results were obtained using a second siRNA (Supplementary Fig. 7a–c). CDH18-silenced cells showed reduced growth (Fig. 3e) suggesting that CDH18 downregulation in fetal-stage epicardium may confer either a proliferative disadvantage impairing epicardium homeostasis and expansion or define the triggering signal for EMT initiation. We observed a gain of SNAI1 expression (Fig. 3f) upon CDH18 downregulation, thus indicating the initiation of EMT. Confirming this functional onset, we observed a strong downregulation of CDH1 (E-cadherin). However, upregulation of CDH2 (N-cadherin) (Fig. 3g) was only modest.
A previous report showed that epicardial cells are more prone to undergo EMT during the early stages of development43,44. We, therefore, questioned, if fetal-like d24 EPI cells were more resistant to initiate EMT than d12 EPI cells, a time-point that portrays a more PE-like state. Accordingly, we silenced CDH18 in d12 EPI cells (d12•si18) finding similar morphofunctional changes as in d24 EPI cells (Fig. 3h–j and Supplementary Fig. 7d–f). Loss of CDH18 was accompanied by gain of SNAI1 (Fig. 3h) and d12•si18 cells showed changed morphology, reduced growth as well as decreased levels of Ki67 (Supplementary Fig. 7f, g and Fig. 3I), thus indicating the onset of EMT. Notably, the switch from CDH1 to CDH2 was much stronger in d12 silenced cells, implying the acquisition of mesenchymal identity (Fig. 3j). Furthermore, CDH18-silenced cells showed enhanced migration (Fig. 3k), endorsing that the downregulation of CDH18 leads to a loss of epicardial identity towards the EPDC-like state. To exclude that our observations were biased due to an early loss of cells we also confirmed that cells displayed reduced proliferation marker expression Ki67, upregulation of SNAI1 and loss of ZO1 3–4 days after silencing (Supplementary Fig. 7h–o), further strengthening, that cells were undergoing EMT upon CDH18 downregulation.
Several pathways are reported to be involved in the regulation of EMT45,46,47,48,49,50,51,52,53,54, with Wnt pathway regulation by Wt1 being linked to epicardial development and cell differentiation6, as well as cardiac regeneration55. Cadherins are known to exert their functions through phosphorylation-mediated protein stabilization of β-catenin in the Wnt signaling pathway56,57. Moreover, a previous report has shown the direct interaction of Cdh18 with β-catenin58. We, therefore, hypothesized that CDH18 might manage its impact on epicardial EMT via the regulation of β-catenin. Indeed, we observed that loss of CDH18 leads to increased levels of β-catenin with decreased phosphorylated (p)-β-catenin as well as an increase in LEF1 (Fig. 3l and Supplementary Fig. 7m, n), indicating the activation of the Wnt pathway.
Interestingly, we found reduced levels of TCF21 upon CDH18-silencing (Fig. 3l). TCF21 acts as an epicardial marker playing pivotal roles during EMT and EPDC-fate regulation between CF and SMC59,60,61. EPI cells showed increased TCF21 expression at the mRNA and protein levels (Supplementary Fig. 8a, b). Reduced levels of TCF21 imply differentiation towards an SMC rather than a CF fate7,60. To confirm this hypothesis, we compared the expression levels of SMC genes in CDH18-silenced EPI cells and iSMCs. We found that CDH18-silenced cells recapitulated the iSMC expression levels of ACTA2 and CNN1 especially in d12 silenced cells and to a lesser degree also in d24-silenced cells (Fig. 3m and Supplementary Fig. 8c–e). Canonical TGFβ-pathway activation seemed to play a noteworthier role for EMT onset in fetal-like d24 compared to PE-like d12 cells, as TGFβ inhibitor withdrawal does enhance the expression levels of ACTA2 in fetal-like d24 cells (Supplementary Fig. 8e). To assess the role of non-canonical activation, we tested the impact of ROCK, inhibitor impairing RhoA-mediated pathway, on CDH18 downregulation in d12 cells. While no significant change could be observed in SNAI1 expression, ACTA2 expression was reduced by ROCK inhibition, albeit only when TGFβ signaling was not blocked (Supplementary Fig. 8f). The acquisition of an SMC-like identity in CDH18-silenced cells was further verified by the loss of ZO1 and the expression of α-SMA (Fig. 3n).
Next, we aimed to investigate the transcriptional contribution of CDH18-downregulation to the SMC phenotype and the enhanced potential to derive SMC-like cells from d12 PE-like cells compared to d24 fetal-like cells. We, therefore, performed RNA-Seq and showed a transcriptome distribution of up to 99.4% combined variance by PCA (Fig. 4a). PE-like CDH18-silenced cells were allocated within the same arm as iSMCs, mapped closer to iSMC and mapped more DEGs (Fig. 4a, b and Supplementary Fig. 9a, b). The downregulation of CDH18 suppressed epicardial identity both at the fetal-like stage and PE-like stage and activated the SMC gene program, while CF-related genes remained unaffected (Fig. 4c, d and Supplementary Fig. 9c). Specifically, in d12 PE-like stage cells silencing of CDH18 influenced gene categories supporting SMC acquisition (Fig. 4c) and showed a greater expression of important markers of EMT activation (Fig. 4f). The altered expression of important cell cycle checkpoint regulators confirmed reduced proliferation upon CDH18 downregulation (Fig. 4g). Finally, we analyzed gene signatures to identify activated signaling pathways, finding that TGF-β and Wnt pathways, which are known to contribute to the SMC phenotype, were affected and activated upon CDH18 downregulation (Fig. 4h, i).
Altogether, we confirmed that CDH18 is an important regulator of the SMC phenotype. CDH18 downregulation led to a loss of epicardial cell identity under the retained inhibition of TGF-β, a situation that normally sustains epicardial hallmarks. The role of CDH18 is essential in epicardial fate and plasticity determination during human development, especially if the loss of CDH18 occurs at an early stage of epicardial specification.
CDH18 is unable to block TGF-β-driven EMT towards SMC differentiation
To investigate the potential of CDH18 expression to regulate epicardial maintenance and SMC differentiation, we evaluated the capacity of CDH18 to inhibit the initiation of EMT towards SMC differentiation. We ectopically overexpressed human CDH18 cDNA (Supplementary Fig. 10a) in d12 EPI cells, which we assessed via mCherry expression, and subsequently induced EMT towards iSMC formation (Fig. 5a). Unfortunately, EPI cells were difficult to transfect, consistent with a previous report on mouse epicardial-like cells34,48. Optimization of the transfection efficiency (Supplementary Table 1) led to ~25% of cells expressing mCherry at d3 post-transfection (Fig. 5b, c and Supplementary Fig. 10b). To obtain a pure CDH18-overexpressing population, we then sorted mCherry-expressing cells (mCherry+) (Supplementary Fig. 10c). The gain of CDH18 expression was confirmed in unsorted and sorted settings (Fig. 5d), and CDH18 protein gain was confirmed in mCherry+ cells (Fig. 5e).
Next, we analyzed the effect of CDH18 overexpression during iSMC induction. We did not observe morphological differences between conditions (Fig. 5f). Nevertheless, CDH18-overexpressing cells showed fewer iSMCs (Fig. 5f), hinting at a potential role of CDH18 in the maintenance of the active epicardium. All populations showed less CDH18 expression upon treatment with TGF-β and bFGF (Fig. 5g), but transfected cells retained CDH18 expression even after undergoing EMT. In agreement with the morphological SMC phenotype (Fig. 5f), all populations showed increased ACTA2 expression upon iSMC induction (Fig. 5h). Notably, ACTA2 levels were significantly reduced in cells still expressing CDH18. RNA-Seq analysis revealed the reduced expression of several SMC markers in CDH18-overexpressing SMCs (Fig. 5i), highlighting an important role of CDH18 in preventing the fully SMC identity acquisition. During development, not only the differentiation of epicardial cells into SMC, but also their migration is important for successful coronary artery formation62,63,64. To investigate the functional properties of CDH18-overexpressing cells, we next tested their invasion ability. We compared EPI cells to cells undergoing spontaneous EMT in absence of TGF-β inhibitor and CDH18-overexpressing SMCs (Fig. 5j). In all conditions, CDH18-overexpressing cells showed reduced invasion capacity. We, therefore, concluded, that although ectopic CDH18 overexpression cannot fully block the TGF-β-driven differentiation to SMC, it partially impairs the acquisition of SMC identity and their functional properties.
CDH18 expression is under the control of GATA4
Finally, to elucidate the regulation of CDH18 in epicardiogenesis, we identified 23 transcription factors (TFs) out of 522 genes that are in correlation with CDH18 and enriched in the late-stage active epicardium (Fig. 6a, b). Among these TF encoding genes, only GATA4 showed a high relative score to bind the CDH18 promoter (Fig. 6c and Supplementary Fig. 11a). During EPI induction GATA4 is expressed in associative correlation to CDH18 (Supplementary Fig. 11b, c and Fig. 6d). GATA4 is a well-characterized regulator at early cardiogenesis playing a pivotal role in PE formation. GATA4-null mice embryos fail to form the epicardium due to impaired PE development, thus being an essential gene in epicardiogenesis37. We, therefore, silenced GATA4 using two siRNAs (siGata4#1 and siGata4#2) at different time points during epicardial differentiation: at d5 of epicardial induction marking the beginning of cell development towards PE-like stage and at d12 marking the specification towards fetal-like epicardium (Supplementary Fig. 11d). GATA4-silenced cells showed reduced cell survival (Fig. 6e, upper panel), but surviving clones did not alter GATA4 levels (Supplementary Fig. 11e) and were able to reconstitute after 7 days (Fig. 6e, lower panel) showing no significant reduction of either GATA4 or CDH18 expression (Fig. 6e, f). Thus, implying that upon efficient silencing of GATA4, cells were not able to survive. We, therefore, concluded that GATA4 is necessary for cells at d5 to differentiate towards the PE-like state. Next, we downregulated GATA4 in d12 EPI cells and observed some morphological changes after siRNA administration over time (Fig. 6g, yellow arrowheads). The downregulation of GATA4 was accompanied by CDH18 downregulation in a similar fashion on both mRNA (Fig. 6h and Supplementary Fig. 11f) and protein level (Fig. 6i). Thus, these results promote GATA4 as a putative regulator of CDH18 expression, highlighting their relevance for the formation and specification of the active epicardium.
The epicardium is essential for cardiogenesis and its re-activation upon injury during adulthood is indispensable for cardiac regeneration. Exploiting epicardial targeting to model cell therapy-based approaches for cardiac repair and regeneration requires detailed knowledge of the mechanisms that regulate the active epicardium. Here, we recapitulated the human epicardiogenesis using hiPSCs and defined a surfaceome of human active epicardium that is later inactivated in the quiescent (adult) epicardium. We define CDH18 as an epicardial biomarker that is exclusively expressed in epicardial cells compared to their derivatives and other cardiac cell types. The epicardium specificity of CDH18 expression can be used as a specific, rather than associated, active epicardial biomarker to both define biological identity and modulate EMT-specific processes towards SMC. Previous identification of epicardial cells was based on the expression of several transcription factors such as WT1, TBX18 and TCF21. However, their expressions are not restrictive to the epicardium. Our findings provide a molecular alternative for the identification of epicardial cells by tracing CDH18 expression, a biomarker of the active fetal epicardium.
Cadherin 18 (CDH18), formerly named cadherin 14, is a type II classical cadherin predominantly expressed in the central nervous system, contributing significantly to axonal development and maintenance38,39,40. This study succeeded in documenting murine CDH18 expression in fetal heart explant culture, known to be derived from the epicardium. Given the thin-shaped nature of the epicardium, its density represents a notable minority in the context of the human heart. We did not detect protein expression in whole trypsinised whole-heart lysates, as mostly cardiomyocytes were isolated. In addition, our study suggests that CDH18 expression shapes the fetal active epicardium transcriptome, but is downregulated in quiescent adult tissue4,65, thus explaining why CDH18 remained undetected in the heart40.
Investigating the expression of CDH18 in human embryonic epicardial cells in vivo is technically difficult due to the lack of technical approaches to trace epicardial development as well as the limited availability of human fetal cells. The use of hiPSC overcomes this issue by enabling recapitulation of epicardiogenesis in vitro. Whereas our study limits our main conclusions to in vitro differentiated epicardial cells and their derivatives, we believe that our key discoveries are physiologically relevant, since RNA-Seq based meta-analysis shows that CDH18 is expressed in the human heart during the third month of gestation, a time point of active development of the cardiovascular system in the human embryo, being in line with the hiPSC-based model shown in this study. Moreover, we documented murine embryonic CDH18 expression in epicardial explant cultures from E14.
Recent reports demonstrate an essential role for WT1 in epicardial cell specification and maturation from the PE, suggesting that junction remodeling is crucial for this transition66,67. We showed that CDH18 is expressed in correlation to WT1 during epicardial development, leading us to hypothesize that CDH18 might play a critical role in the specification and maintenance of epicardial cell identity, similar to WT1.
During human heart formation as well as during cardiac repair epicardial cells also contribute to coronary vessel formation by providing necessary vascular SMC and pericytes, which invade the myocardium. Notably, we provide compelling evidence to indicate that CDH18 is critical for the modulation and establishment of epicardial-derived SMCs. We proved that CDH18 downregulation modulates the expression of EMT markers and impacts epicardial identity. Recent studies characterized a CDH18 tumor-suppressor role in glioma carcinogenesis and progression, demonstrating an active role in invasion and cell migration38, important processes that also relate to cardiac regeneration. The ectopic overexpression of CDH18 was reported in relation to decreased migration capacity in accordance with our study. Most importantly, our study gained mechanistic insights into the biological regulation during the initiation of EMT in epicardial cells. The loss of CDH18 decreased proliferation, CDH1 expression and β-catenin. Interaction with and regulation of stabilization of β-catenin via cadherins is critical for the cellular organization, polarity and development56,57. Indeed, a previous study showed that Cdh14 interacts with and stabilizes β-catenin, similar to Cdh158. Another study reported, that type II Cdh14 (Cdh18) along with type I E-cadherin (Cdh1) and N-cadherin (Cdh2) can bind to G12 family proteins68, which are known to regulate β-catenin translocation from the cell surface, a function associated with Rho-dependent cytoskeletal rearrangement. In our study, we demonstrated that the loss of CDH18 increased β-catenin levels, indicating Wnt activation and the promotion of EMT. The activation of Wnt signaling via β-catenin has long been deemed important for epicardial EMT and SMC differentiation which known to be regulated by Wt16,7,29,53,69,70. Although controversial52, our study supports the necessity of β-catenin for SMC differentiation in vitro70.
TCF21 is another key regulator of epicardial plasticity and EMT, promoting CF over SMC fate60. It is possible that rather than the activation of specific signaling pathways, the expression of TCF21 is of far greater importance for cell-fate decisions. Thus, investigating Wnt and TGF-β signaling pathways and their influence on TCF21 could shed more light on this area, which will also allow understanding cell-fate decisions during cardiac regeneration, in particular those physiological stimuli that presumably force cells to assume a CF rather than pericyte phenotype. In addition, TCF21 is known to be induced by retinoic acid (RA), thereby delaying SMC differentiation60,71,72. We showed that the expression of ALDH1A2 is not upregulated until d24 and is low in d12 EPI cells. Thus, our observation of d24-silenced cells expressing lower levels of the SMC marker ACTA2 could be linked to the higher presence of RA and TCF21 in fetal-like cells compared to PE-like cells. During cardiac repair, the expression of ALDH1A2 is re-activated, which might explain the preferred CF fate of epicardial cells that undergo EMT upon re-activation. Here, we showed loss of TCF21 upon loss of CDH18, suggesting a role for β-catenin-mediated Wnt activation in SMC differentiation. Furthermore, this observation occurred under the inhibition of the TGF-β co-receptor ALK5, which is known to impair EMT73. Our findings provide new insights into the regulation of the EMT process as a crucial step mediating epicardial-driven regenerative responses.
The exact mechanisms governing epicardial EMT and subsequent cell-fate decision of EPDCs in active epicardium are not well understood. Most experiments have used chick, avian or mouse models, which can differ considerably from humans. We observed the activation of downstream TGF-β signal targets, confirmed by RNA-Seq, upon CDH18 silencing albeit treatment with SB431542. Our study endorses the link between TGF-β signaling and SMC phenotype29,31,34,70, as we observed SMC differentiation and TGF-β activation upon lower CDH18 expression. Although CDH18 overexpression could not completely block TGF-β-driven SMC differentiation, we found a partial prevention in the acquisition of SMC identity, indicated by lower levels of ACTA2. Altogether, our findings indicate a causal role of CDH18 in TGF-β pathway during the acquisition of SMC identity.
CDH18 is a cell surface protein linked to G12 family proteins68, and RhoA pathway-mediated non-canonical TGF-β pathway, which has been linked to EMT and cell invasion45,48,49,50,74,75, might be activated upon the silencing. Interestingly, one study showed that inhibition of the RhoA pathway does not influence canonical TGF-β activation but still impairs SMC formation76. In our study, however, for those experiments that kept ALK5 inhibition conditions, ROCK inhibitor did not alter SMC marker expression upon CDH18 silencing. Only when ALK5 inhibitor was absent, did inhibition of ROCK lead to a decrease in ACTA2.
Recently, a new study linked a TGFβ-independent causal role of the extracellular matrix (ECM) protein agrin to epicardial EMT mediated by dystroglycan17. This report also showed that their findings occur via Wnt signaling activation, a phenotype that could have common denominators with our study. Future studies are needed to shed more light into the different behavior of cells during human epicardial development. Such studies will also help deepen our understanding of epicardium re-activation upon cardiac injury and advance the field of cardiac regenerative medicine.
Finally, we demonstrated the correlative relationship between CDH18 and GATA4 expression, proposing GATA4 as a putative regulator of CDH18. This effect likely occurs by static binding to the CDH18 promoter near the transcriptional starting site, reinforcing GATA4 as an important activator of the embryonic epicardium program.
CDH18 is a highly conserved protein in higher mammals and species that evolutionary developed a double closed circulatory system, maintaining an amino acid alignment of over 95% identity and 100% query sequence coverage relative to humans. The conserved sequence suggests that CDH18 has remained relatively unchanged far back up the phylogenetic tree in species developing a higher compartmentalized system to create a physical separation of oxygenated and deoxygenated blood (Supplementary Fig. 12). Higher demand for SMC might explain CDH18 conservation in development (Supplementary Fig. 13). Altogether, these findings indicate a specific role for CDH18 in epicardial regeneration and development, both processes that are under the control of evolutionarily conserved pathways
In conclusion, our work defines a biological function of CDH18 in the epicardial context, enabling roads for the manipulation and therapeutic gene modulation of the active epicardium for cardiac repair and regeneration.
Cell lines and culture conditions
The hiPSC lines 201B7 and 409B2 (retroviral reprogramming by Yamanaka factors)27 as well as a MYH6-eGFP reporter cell line (MYH6-EIP4)41 were cultured in ReproCell ES media containing 4 ng/ml bFGF on irradiated MEF feeder cells. For removal of the feeder layer, the cells were treated with CTK. Feeder-free 201B7 were cultured in complete StemFit® AK02N media on iMatrix-511 (Matrixome) coated dishes. All cell lines used were female. MEC1 Mouse Embryonic Epicardial Cell Line SCC187 (Merck Millipore) was cultured and maintained in DMEM supplemented with 10% FBS. Mycoplasma testing was performed on a regular basis to exclude contamination.
For epicardial differentiation, single cell suspension of hiPSC was generated using Accutase and subsequently cells were plated onto low-attachment HEMA-coated plates (6000-8000/96-well) to form embryoid bodies (EBs). Differentiation media was composed of complete StemPro®-34 media supplemented with 50 μg/ml ascorbic acid, 2 mM L-glutamine, 0.4 μM monothioglycerol and 150 mg/ml transferrin. To initiate differentiation, 0.5% Matrigel, 10 μM Y-27632 and 2 ng/ml human recombinant (hr)BMP4 were added to the differentiation media. After 24 h (h) EBs were cultured in differentiation media with a final concentration of 10 ng/ml hrBMP4, 2 ng/ml Activin A and 5 ng/ml hrbFGF. After 84 h, EBs were collected, dissociated using Accutase, and plated onto 0.1% gelatin-coated dishes (0.3 × 105 cells/cm2) in differentiation media containing 3 mM CHIR99021, 30 ng/ml hrBMP4, 5 ng/ml hrVEGF and 10 µM SB431542. From d7 onwards, induced hiPSC-derived epicardial-like (EPI) cells were maintained in maintenance media (DMEM containing 10% FBS and 10 µM SB431542). For cell passaging, EPI cells were detached by Accutase treatment for 5 min and replated as described above.
For the silencing experiments, Silencer® Select siRNAs (ThermoFisher) were diluted to a 10 µmol stock solution: siCdh18#1 (s2816), siCdh18#2 (s2817), siGata4#1 (s535120) and siGata4#2 (s535121). To silence cells, 10 µl siRNA stock was diluted in 500 µl Opti-MEM, and 10 µl RNAiMax was added (silencing solution) followed by 10 min incubation at room temperature. The solution was added dropwise into one well of a six-well plate containing 2 ml maintenance media and 0.2–0.3 × 105 cells seeded a day prior. The media was exchanged 24 h later, and the cells were cultured as usual thereafter. For the immunocytochemistry experiments, 260 µl silencing solution was added dropwise into one well of a 12-well plate containing 1 ml maintenance media and 0.05 × 105 cells seeded a day prior to silencing.
For CDH18 cDNA overexpression experiments, the cells were seeded to be 60–70% confluent at least one day prior to the transfection. For optimization of transfection protocol, transfection reagents and ratios as well as the plasmid amount were used as indicated in Supplementary Table 1. Unless otherwise stated, 6 µg plasmid per one well of a six-well plate or 34 µg plasmid per 10 cm dish were transfected using FuGENE®6 at an agent-to-DNA ratio of 2:1. Amounts of reagents were calculated using FuGENE®HD Protocol Database web tool. Cell sorting was performed by FACS via assessment of mCherry expression (Texas-Red positive cells).
Induction of EMT
For the induction of EMT, EPI cells were passaged one day prior, at a density of 0.1–0.2 × 105 per well of a six-well plate. For differentiation towards CF, the cells were treated with 10 ng/ml bFGF in 10% FBS DMEM for at least 8 days. For differentiation towards SMC cells were treated with 5 ng/ml TGF-β for 4 days followed by 10 ng/ml bFGF for another 4 days in 10% FBS DMEM.
Isolation of mouse fetal hearts and whole-heart lysate
Embryos at E14 were collected into pre-warmed PBS and any extraembryonic tissue was removed using tweezers. Embryos were decapitated, the chest cavity was opened by ripping the anterior side apart and fetal hearts were isolated mechanically into a separate dish with pre-warmed PBS. Whole heart protein lysate was prepared by firstly cutting hearts into small pieces and treatment with trypsin for 20 min at 37 °C. The cell suspension was re-suspended, treated for an additional 20 min at 37 °C and supernatant was collected after centrifugation at 300 × g for 5 min.
Ex vivo explant assay
Collected hearts at E14 were cut into 4–8 pieces and placed onto 0.1% gelatin-coated dished in DMEM (low glucose) supplemented with 15% FBS. Outgrowth cultures formed as early as 24 h post placing. After 72 h heart pieces were manually removed with tweezers. Explant cells were passage by trypsinization the following day and cultured in 10% FBS-DMEM (low glucose) supplemented with 10 µM SB431542. Cells were collected 1–3 days after passage. EMT was induced as described above in 10% FBS-DMEM (low glucose).
For RNA extraction, live cells were collected as du- or triplicate in QIAzol lysis reagent, and total RNA isolation was performed using the RNAeasy micro kit (Qiagen) according to the manufacturer’s manual. The generation of cDNA was performed by using the ReverTra Ace system (Toyobo BIOTECH) according to the manufacturer’s manual. QRT-PCR was performed in du- or triplicate either using TaqMan gene expression assays (WT1 Hs01103750_g1; TBX18 Hs01385457_m1; ALDH1A2 Hs00108254_m1; TCF21 Hs00162646_m1) in TaqMan master mix solution or SYBR green (ThermoFisher) with primers against identified candidates (Supplementary Table 2) in SYBR green master mix solution according to the manufacturers’ respective manuals. Data acquisition was carried out by StepOne Plus (AppliedBiosystems). Analysis was performed using Microsoft Excel and GraphPad Prism. Values without readout were not included. The distribution of values was verified by a box plot depiction to determine the appropriate statistical analysis. For direct comparison of dataset normally distributed data were analyzed by students t-test, as indicated in figure legends, otherwise, the Mann–Whitney test was used. Comparisons of more than two datasets were carried out via ANOVA, according to experimental conditions, as indicated in the figure legends. All error bars represent the standard error of the mean (SEM). For co-expression analysis, qRT-PCR data was used to generate a regression line by GraphPad Prism v7 and higher.
Cells were fixed by 4% PFA treatment for 15 min and stored in PBS at 4 °C. Blocking was performed for 30–45 min in blocking buffer: 1% BSA and 0.5% Triton X for nuclear staining or 0.1% Tween 20 for surface or intracellular staining and 0.1 M glycine in PBS. Following three PBS washes, primary antibody was added in blocking buffer without glycine and incubated overnight at 4 °C. The primary antibodies used are as follows: Anti-WT1 1:200 (Abcam; ab89901), Anti-ZO1 1:100–300 (Invitrogen; ZO1-1A12), Anti-POSTN 1:200 (ThermoFisher; PA5-98301), Anti-α-SMA 1:200 (Abcam; ab7817) and Anti-Ki67 1:200 (BioLegend; 350502). The next day, the cells were washed thrice with PBS, secondary antibody (Invitrogen, 1:1000–2000) was added in 1% BSA-PBS and incubated for 2 h at room temperature. The secondary antibodies were used are as follows: goat-anti-mouse-Alexa488 (A11001), goat-anti-rabbit-Alexa546 (A11010), goat-anti-mouse-Alexa546 (A11030), goat-anti-mouse-Alexa594 (A11032), goat-anti-rabbit-Alexa647 (A21245), goat-anti-mouse-Alexa647 (A21236). Following three washes with PBS, Hoechst or DAPI (5 ng/ml, 1:10,000) was added for nuclear counterstaining.
Flow cytometry analysis/FACS
The cell was dissociated into single-cell suspension by Accutase treatment, washed twice and fixed using 4% PFA for 15 min. For nuclear staining, the cells were permeabilized by 0.1% Triton X solution. Conjugated antibodies were diluted 1:50 in FACS buffer (5% FBS-PBS) and incubated at room temperature for 30–45 min. The conjugated antibodies used are as follows: WT1-Alexa488 (Abcam; ab202635), rb-IgG-Alexa-488 (Abcam; ab199091), CDH18-FITC (Biorbyt Ltd; orb7854), CD22-APC (BD; 562860) CD31-APC (BioLegend; 303116) and CD144-FITC (BD; 560874).
For sorting, a single cell suspension was generated as described above, and counterstaining with 1:1,000 DAPI (5 ng/ml) was performed for dead cell exclusion. Sorting gates were set as described.
Analysis was performed using BD FACSDiva v6 or v8 and FlowJo v10. The cell population was identified by FSC/SSC gating and doublets discrimination was performed. Negative gates were established using unstained samples or isotype controls and negative control samples. Positive gates were set to contain no negative signal.
Cells were detached using Accutase and lysed in Mammalian Protein Extraction Reagent (M-PER) (Thermo Scientific; 78501) buffer. The amount of protein was determined by the Bradford assay using BSA as the standard. The primary antibodies (dilution 1:1000) used are as follows: Anti-CDH18 antibody (Proteintech 13091-1-AP), Anti-GATA4 (CST; 369665S (D3A3M)), Anti-SNAI1 (Abcam; ab63371), Anti-α-SMA (Abcam; ab11952), Anti-β-actin (Sigma; A5441), Anti-WT1 (Abcam; ab89901), Anti-CD22 (Abcam; ab207727), Anti-TCF21 (Abcam; ab32981), Anti-β-catenin (CST; 8814), Anti-phosporylated-β-catenin (S33/S37/T41) (CST; 9561), Anti-TNNI1 (Abcam; ab203515). The secondary antibodies (dilution 1:5000) used were as follows: goat anti-rabbit-horseradish peroxidase (HRP) (Abcam; ab97051), rabbit anti-mouse-HRP (Abcam; ab97046). Blots derive from the same experiment and were processed in parallel. Unprocessed blots are shown in Supplementary Figs. 14–18.
A time-course curve of parental (scr) and siRNA-expressing cells was generated by seeding 4.5 × 103 cells into 6-cm bottom-well diameter dishes. After 24 h, fresh medium was replaced and cells were fixed to be referred as a standard for relative growth (day 0). Relative growth was assessed every 48 h (2 days), fixing in 2% PFA and stained with 1% crystal violet (Sigma; C6158-50G). After extensive cell washing, crystal violet was solubilized in 20% acetic acid (Sigma) and quantified absorbance at 595 nm as a relative assessment of cell number (PerkinElmer; EnVision 2104 Multilabel Reader). Relative values in Fig. 4 represent cell growth of daughter the indicated medium. Percentage zero (0%) refers to initial growth at day 0.
Cell were silenced as described above and collected 4 days later. 2.5 × 104 cells were seeded onto a Corning® Biocoat® Matrigel® Invasion chamber and incubated for 24 or 48 h. The removal of non-invasive cells was carried out as stated in the Corning® Biocoat® Matrigel® Invasion manual, and the cells were fixed for 2 min in methanol followed by staining using crystal violet overnight and finally washed with distilled water. Pictures were taken using BZ-X710 (Keyence) and cell counting was performed with ImageJ. Cell counting was performed blinded using ImageJ v1.52. Images were converted to 8-bit greyscale format, and the same threshold was set for all images to highlight cell structures. Potential cell clusters were separated by the watershed function, and only structures 0.01-0.1 pixel2 in size were counted. Graphs were generated using GraphPad Prism v8.
Retrospective analysis of CDH18 gene expression was done using GSE84085 (GEO) for epicardium development and GSE7307 for cardiovascular-derived tissues (subset of aorta, coronary artery, heart, heart atrium and heart ventricle) [GEO and R2; genomic analysis and visualization platform (http://r2.amc.nl/)].
Promoter sequences were identified by Ensembl and then screened for transcription factor binding sites with PROMO-ALGGEN (based on TRANSFAC v8.3) and JASPAR 2016.
Image acquisition, processing and analysis
Microscopy images were taken by BZ-X710 and processed by BZ-X Analyzer (Keyence); pseudo-coloring was used as indicated in figure legends. For phase-contrast and fluorescence pictures, a representative section was chosen, cropped and magnified. Western blot data were recorded by LAS4000 (Cytiva). General image analysis was performed using Microsoft powerpoint as well as ImageJ v1.52.
Data normalization was carried out using NOISeq. The final processed data and raw fastq files generated de novo in this study were submitted to Gene Expression Omnibus (GEO) under the accession code GSE165450. The raw data were analyzed by using RStudio for further analysis on gene expression. Gene expression data for reading, exploring and pre-processing were conducted using the Bioconductor package NOISeq pipeline to perform data and differential expression analysis for RNA-Seq. A hierarchical derived cluster dendrogram was generated by using hclust, stats package, and agnes in the cluster package. Distances were assessed by using Manhattan city-block distance algorithm. K-means were calculated with the kmeans function. Distance and correlation matrices were computed and plotted by using get_dist, fviz_dist included in the factoextra package. The fviz_cluster function was used to compute cluster scatter plots. Heatmaps were depicted to cluster the expression data for DEGs using R script. Statistical analysis and visualization of the functional profiles of genes as well as gene clusters for GO terms were conducted using the DOSE and clusterProfiler. Ingenuity pathway analysis (IPA) was used to develop an upstream pathway analysis as well as pathway activity patterns. Chord diagrams were generated by using GOplot.
The CDH18 overexpression plasmid used in this study is available from VectorBuilder [VB200615-1011cyh]. Other materials are available from the corresponding author upon reasonable request, but we may require payment and/or a Materials Transfer Agreement.
All experiments involving animals were approved by the Kyoto University Animal Experimentation Committee and carried out in accordance to the Guidelines for Animal Experiments of Kyoto University and Guide to the Care and Use of Laboratory Animals by the Institute of Animal Resources.
Further information on research design is available in the Nature Research Reporting Summary linked to this article.
All data supporting this study are available within this publication and its supplementary information or can be provided upon request. The RNA-Seq dataset generated in this study is deposited at Gene Expression Omnibus (GEO) with the accession code GSE165450.
Bargehr, J. et al. Epicardial cells derived from human embryonic stem cells augment cardiomyocyte-driven heart regeneration. Nat. Biotechnol. 37, 895–906 (2019).
Pérez-Pomares, J. M. & de la Pompa, J. L. Signaling during epicardium and coronary vessel development. Circ. Res. 109, 1429–1442 (2011).
van Wijk, B. & van den Hoff, M. Epicardium and myocardium originate from a common cardiogenic precursor pool. Trends Cardiovasc. Med. 20, 1–7 (2010).
Zhou, B. et al. Adult mouse epicardium modulates myocardial injury by secreting paracrine factors. J. Clin. Investig. 121, 1894–1904 (2011).
Gittenberger-de Groot, A. C., Vrancken Peeters, M. P., Mentink, M. M., Gourdie, R. G. & Poelmann, R. E. Epicardium-derived cells contribute a novel population to the myocardial wall and the atrioventricular cushions. Circ. Res. 82, 1043–1052 (1998).
von Gise, A. et al. WT1 regulates epicardial epithelial to mesenchymal transition through β-catenin and retinoic acid signaling pathways. Dev. Biol. 356, 421–431 (2011).
Braitsch, C. M. & Yutzey, K. E. Transcriptional control of cell lineage development in epicardium-derived cells. J. Dev. Biol. 1, 92–111 (2013).
Katz, T. C. et al. Distinct compartments of the proepicardial organ give rise to coronary vascular endothelial cells. Dev. Cell 22, 639–650 (2012).
Zhou, B. et al. Epicardial progenitors contribute to the cardiomyocyte lineage in the developing heart. Nature 454, 109–113 (2008).
Krainock, M. et al. Epicardial epithelial-to-mesenchymal transition in heart development and disease. J. Clin. Med. 5, 27 (2016).
Smart, N., Dube, K. N. & Riley, P. R. Coronary vessel development and insight towards neovascular therapy. Int. J. Exp. Pathol. 90, 262–283 (2009).
van Wijk, B., Gunst, Q. D., Moorman, A. F. & van den Hoff, M. J. Cardiac regeneration from activated epicardium. PLoS ONE 7, e44692 (2012).
Rao, K. S. & Spees, J. L. Harnessing epicardial progenitor cells and their derivatives for rescue and repair of cardiac tissue after myocardial infarction. Curr. Mol. Biol. Rep. 3, 149–158 (2017).
Cao, Y. & Cao, J. Covering and re-covering the heart: development and regeneration of the epicardium. J. Cardiovasc. Dev. Dis. 6, 3 (2018).
Ramjee, V. et al. Epicardial YAP/TAZ orchestrate an immunosuppressive response following myocardial infarction. J. Clin. Investig. 127, 899–911 (2017).
Wei, K. et al. Epicardial FSTL1 reconstitution regenerates the adult mammalian heart. Nature 525, 479–485 (2015).
Sun, X. et al. The extracellular matrix protein agrin is essential for epicardial epithelial-to-mesenchymal transition during heart development. Development 148, dev197525 (2021).
Walma, D. A. C. & Yamada, K. M. The extracellular matrix in development. Development 147, dev175596 (2020).
Halbleib, J. M. & Nelson, W. J. Cadherins in development: cell adhesion, sorting, and tissue morphogenesis. Genes Dev. 20, 3199–3214 (2006).
Foty, R. A. & Steinberg, M. S. The differential adhesion hypothesis: a direct evaluation. Dev. Biol. 278, 255–263 (2005).
Hogan, C. et al. Rap1 regulates the formation of E-cadherin-based cell–cell contacts. Mol. Cell. Biol. 24, 6690–6700 (2004).
Johnson, E., Theisen, C. S., Johnson, K. R. & Wheelock, M. J. R-cadherin influences cell motility via Rho family GTPases. J. Biol. Chem. 279, 31041–31049 (2004).
Luo, J., Treubert-Zimmermann, U. & Redies, C. Cadherins guide migrating Purkinje cells to specific parasagittal domains during cerebellar development. Mol. Cell. Neurosci. 25, 138–152 (2004).
Maeda, M., Johnson, K. R. & Wheelock, M. J. Cadherin switching: essential for behavioral but not morphological changes during an epithelium-to-mesenchyme transition. J. Cell Sci. 118, 873–887 (2005).
Reintsch, W. E., Habring-Mueller, A., Wang, R. W., Schohl, A. & Fagotto, F. beta-Catenin controls cell sorting at the notochord-somite boundary independently of cadherin-mediated adhesion. J. Cell Biol. 170, 675–686 (2005).
Takahashi, K. et al. Induction of pluripotent stem cells from adult human fibroblasts by defined factors. Cell 131, 861–872 (2007).
Takahashi, K. & Yamanaka, S. Induction of pluripotent stem cells from mouse embryonic and adult fibroblast cultures by defined factors. Cell 126, 663–676 (2006).
Bao, X. et al. Long-term self-renewing human epicardial cells generated from pluripotent stem cells under defined xeno-free conditions. Nat. Biomed. Eng. 1, 0003 (2016).
Bax, N. A. M. et al. In vitro epithelial-to-mesenchymal transformation in human adult epicardial cells is regulated by TGFβ-signaling and WT1. Basic Res. Cardiol. 106, 829–847 (2011).
Burridge, P. W., Keller, G., Gold, J. D. & Wu, J. C. Production of de novo cardiomyocytes: human pluripotent stem cell differentiation and direct reprogramming. Cell Stem Cell 10, 16–28 (2012).
Compton, L. A., Potash, D. A., Mundell, N. A. & Barnett, J. V. Transforming growth factor-β induces loss of epithelial character and smooth muscle cell differentiation in epicardial cells. Dev. Dyn. 235, 82–93 (2006).
Iyer, D. et al. Robust derivation of epicardium and its differentiated smooth muscle cell progeny from human pluripotent stem cells. Development 142, 1528–1541 (2015).
Mummery, C. L. et al. Differentiation of human embryonic stem cells and induced pluripotent stem cells to cardiomyocytes: a methods overview. Circ. Res. 111, 344–358 (2012).
Tuyn, J. V. et al. Epicardial cells of human adults can undergo an epithelial-to-mesenchymal transition and obtain characteristics of smooth muscle cells in vitro. STEM CELLS 25, 271–278 (2007).
Witty, A. D. et al. Generation of the epicardial lineage from human pluripotent stem cells. Nat. Biotechnol. 32, 1026–1035 (2014).
Zhao, J. et al. Efficient differentiation of TBX18(+)/WT1(+) epicardial-like cells from human pluripotent stem cells using small molecular compounds. Stem Cells Dev. 26, 528–540 (2017).
Watt, A. J., Battle, M. A., Li, J. & Duncan, S. A. GATA4 is essential for formation of the proepicardium and regulates cardiogenesis. Proc. Natl Acad. Sci. USA 101, 12573–12578 (2004).
Bai, Y. H. et al. A novel tumor-suppressor, CDH18, inhibits glioma cell invasiveness via UQCRC2 and correlates with the prognosis of glioma patients. Cell. Physiol. Biochem. 48, 1755–1770 (2018).
Klein, M. E. et al. PDLIM7 and CDH18 regulate the turnover of MDM2 during CDK4/6 inhibitor therapy-induced senescence. Oncogene 37, 5066–5078 (2018).
Shibata, T., Shimoyama, Y., Gotoh, M. & Hirohashi, S. Identification of human cadherin-14, a novel neurally specific type II cadherin, by protein interaction cloning. J. Biol. Chem. 272, 5236–5240 (1997).
Funakoshi, S. et al. Enhanced engraftment, proliferation, and therapeutic potential in heart using optimized human iPSC-derived cardiomyocytes. Sci. Rep. 6, 19111 (2016).
Yang, J. et al. Guidelines and definitions for research on epithelial-mesenchymal transition. Nat. Rev. Mol. Cell Biol. 21, 341–352 (2020).
Moerkamp, A. T. et al. Human fetal and adult epicardial-derived cells: a novel model to study their activation. Stem Cell Res. Ther. 7, 174 (2016).
Wei, X. et al. Biological characteristics of embryonic epicardial cells in vitro correlate with embryonic day. Acta Biochim. Biophys. Sin. 49, 14–24 (2017).
Dokic, D. & Dettman, R. W. VCAM-1 inhibits TGFβ stimulated epithelial–mesenchymal transformation by modulating Rho activity and stabilizing intercellular adhesion in epicardial mesothelial cells. Dev. Biol. 299, 489–504 (2006).
Dronkers, E., Wauters, M. M. M., Goumans, M. J. & Smits, A. M. Epicardial TGFβ and BMP signaling in cardiac regeneration: what lesson can we learn from the developing heart? Biomolecules 10, 404 (2020).
Goumans, M. -J. & Ten Dijke, P. TGF-β signaling in control of cardiovascular function. Cold Spring Harbor Perspect. Biol. 10, a022210 (2018).
Hill, C. R. et al. BMP2 signals loss of epithelial character in epicardial cells but requires the Type III TGFβ receptor to promote invasion. Cell. Signal. 24, 1012–1022 (2012).
Ishii, Y., Garriock, R. J., Navetta, A. M., Coughlin, L. E. & Mikawa, T. BMP signals promote proepicardial protrusion necessary for recruitment of coronary vessel and epicardial progenitors to the heart. Dev. Cell 19, 307–316 (2010).
Lu, J. et al. Coronary smooth muscle differentiation from proepicardial cells requires rhoA-mediated actin reorganization and p160 rho-kinase activity. Dev. Biol. 240, 404–418 (2001).
Mack, C. P. Signaling mechanisms that regulate smooth muscle cell differentiation. Arterioscler. Thromb. Vasc. Biol. 31, 1495–1505 (2011).
Rudat, C., Norden, J., Taketo, M. M. & Kispert, A. Epicardial function of canonical Wnt-, Hedgehog-, Fgfr1/2-, and Pdgfra-signalling. Cardiovasc. Res. 100, 411–421 (2013).
Smith, C. L., Baek, S. T., Sung, C. Y. & Tallquist, M. D. Epicardial-derived cell epithelial-to-mesenchymal transition and fate specification require PDGF receptor signaling. Circ. Res. 108, e15–e26 (2011).
Zhou, B. P. et al. Dual regulation of Snail by GSK-3beta-mediated phosphorylation in control of epithelial–mesenchymal transition. Nat. Cell Biol. 6, 931–940 (2004).
Duan, J. et al. Wnt1/betacatenin injury response activates the epicardium and cardiac fibroblasts to promote cardiac repair. EMBO J. 31, 429–442 (2012).
Heuberger, J. & Birchmeier, W. Interplay of cadherin-mediated cell adhesion and canonical Wnt signaling. Cold Spring Harb. Perspect. Biol. 2, a002915 (2010).
Nelson, W. J. & Nusse, R. Convergence of Wnt, beta-catenin, and cadherin pathways. Science 303, 1483–1487 (2004).
Shimoyama, Y., Takeda, H., Yoshihara, S., Kitajima, M. & Hirohashi, S. Biochemical characterization and functional analysis of two type II classic cadherins, cadherin-6 and -14, and comparison with E-cadherin. J. Biol. Chem. 274, 11987–11994 (1999).
Acharya, A. et al. The bHLH transcription factor Tcf21 is required for lineage-specific EMT of cardiac fibroblast progenitors. Development 139, 2139–2149 (2012).
Braitsch, C. M., Combs, M. D., Quaggin, S. E. & Yutzey, K. E. Pod1/Tcf21 is regulated by retinoic acid signaling and inhibits differentiation of epicardium-derived cells into smooth muscle in the developing heart. Dev. Biol. 368, 345–357 (2012).
Hu, H., Lin, S., Wang, S. & Chen, X. The role of transcription factor 21 in epicardial cell differentiation and the development of coronary heart disease. Front. Cell Dev. Biol. 8, 457 (2020).
Lupu, I. E., De Val, S. & Smart, N. Coronary vessel formation in development and disease: mechanisms and insights for therapy. Nat. Rev. Cardiol. 17, 790–806 (2020).
Tevosian, S. G. et al. FOG-2, a cofactor for GATA transcription factors, is essential for heart morphogenesis and development of coronary vessels from epicardium. Cell 101, 729–739 (2000).
Zhuge, Y. et al. Role of smooth muscle cells in cardiovascular disease. Int. J. Biol. Sci. 16, 2741–2751 (2020).
Wu, M. et al. Epicardial spindle orientation controls cell entry into the myocardium. Dev. Cell 19, 114–125 (2010).
Risebro, C. A., Vieira, J. M., Klotz, L. & Riley, P. R. Characterisation of the human embryonic and foetal epicardium during heart development. Development 142, 3630–3636 (2015).
Velecela, V. et al. Epicardial cell shape and maturation are regulated by Wt1 via transcriptional control of Bmp4. Development 146, ev178723 (2019).
Meigs, T. E., Fields, T. A., McKee, D. D. & Casey, P. J. Interaction of Gα12 and Gα13 with the cytoplasmic domain of cadherin provides a mechanism for β-catenin release. PNAS 98, 519–524 (2001).
Rudat, C. & Kispert, A. Wt1 and epicardial fate mapping. Circ. Res. 111, 165–169 (2012).
Zamora, M., Männer, J. & Ruiz-Lozano, P. Epicardium-derived progenitor cells require β-catenin for coronary artery formation. PNAS 104, 18109–18114 (2007).
Azambuja Ana, P. et al. Retinoic aid and VEGF delay smooth muscle relative to endothelial differentiation to coordinate inner and outer coronary vessel wall morphogenesis. Circ. Res. 107, 204–216 (2010).
Guadix, J. A. et al. Wt1 controls retinoic acid signalling in embryonic epicardium through transcriptional activation of Raldh2. Development 138, 1093–1097 (2011).
Sridurongrit, S., Larsson, J., Schwartz, R., Ruiz-Lozano, P. & Kaartinen, V. Signaling via the Tgf-beta type I receptor Alk5 in heart development. Dev. Biol. 322, 208–218 (2008).
Sánchez, N. S. & Barnett, J. V. TGFβ and BMP-2 regulate epicardial cell invasion via TGFβR3 activation of the Par6/Smurf1/RhoA pathway. Cell. Signal. 24, 539–548 (2012).
Zhao, W. et al. Effect of TGF-β1 on the migration and recruitment of mesenchymal stem cells after vascular balloon injury: involvement of matrix metalloproteinase-14. Sci. Rep. 6, 21176 (2016).
Tao, J., Barnett, J. V., Watanabe, M. & Ramírez-Bergeron, D. Hypoxia supports epicardial cell differentiation in vascular smooth muscle cells through the activation of the TGFβ pathway. J. Cardiovasc. Dev. Dis. 5, (2018).
We would like to thank Professor Shinya Yamanaka and all members of the Yoshida laboratory for their constructive feedback on this study. J.J. thanks the Japanese Government (MEXT) scholarship for its financial support. This work was supported by a grant from the Leducq Foundation (18CVD05) and JSPS KAKENHI Grants (20K17078 (A.L.-C.), 21H02912 (Y.Y.), and 17H04176 (Y.Y.)). Funding is also provided by the Research Center Network for Realization of Regenerative Medicine, Japan Agency of Medical Research and Development (AMED) (JP19bm0104001, JP19bm0804008, JP20bm0204003, and JP20bm0804022) (Y.Y.), the Research on Regulatory Science of Pharmaceuticals and Medical Devices, AMED (JP19mk0104117) (Y.Y.), the Research Project for Practical Applications of Regenerative Medicine, AMED (JP19bk0104095) (Y.Y.), the iPS Cell Research Fund (A.L.-C. and Y.Y.) and the SECOM Science and Technology Foundation (Y.Y.). We would like to express our heartfelt gratitude to Peter Karagiannis for proofreading and advice in manuscript writing, Kazuhisa Chonabayashi for providing control cells, Yoko Uematsu, Kaoru Shimizu and Mikako Marx Mori for their administrative support.
J.J., T.Y., Y.K., E.M.V.-S., M.N. and A.L.-C. declare no competing interest. Y.Y. is a scientific advisor of Orizuru Therapeutics.
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Junghof, J., Kogure, Y., Yu, T. et al. CDH18 is a fetal epicardial biomarker regulating differentiation towards vascular smooth muscle cells. npj Regen Med 7, 14 (2022). https://doi.org/10.1038/s41536-022-00207-w