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. 2021 Jun 28;9:701175. doi: 10.3389/fcell.2021.701175

Interactions and Feedbacks in E-Cadherin Transcriptional Regulation

Miguel Ramirez Moreno 1, Przemyslaw A Stempor 2, Natalia A Bulgakova 1
PMCID: PMC8273600  PMID: 34262912

Abstract

Epithelial tissues rely on the adhesion between participating cells to retain their integrity. The transmembrane protein E-cadherin is the major protein that mediates homophilic adhesion between neighbouring cells and is, therefore, one of the critical components for epithelial integrity. E-cadherin downregulation has been described extensively as a prerequisite for epithelial-to-mesenchymal transition and is a hallmark in many types of cancer. Due to this clinical importance, research has been mostly focused on understanding the mechanisms leading to transcriptional repression of this adhesion molecule. However, in recent years it has become apparent that re-expression of E-cadherin is a major step in the progression of many cancers during metastasis. Here, we review the currently known molecular mechanisms of E-cadherin transcriptional activation and inhibition and highlight complex interactions between individual mechanisms. We then propose an additional mechanism, whereby the competition between adhesion complexes and heterochromatin protein-1 for binding to STAT92E fine-tunes the levels of E-cadherin expression in Drosophila but also regulates other genes promoting epithelial robustness. We base our hypothesis on both existing literature and our experimental evidence and suggest that such feedback between the cell surface and the nucleus presents a powerful paradigm for epithelial resilience.

Keywords: adhesion, JAK/STAT, heterochromatin, HP1, PAR-3

Introduction

Cells in epithelia adhere to both a common substrate and neighbouring cells (Hagios et al., 1998). The molecules mediating this adhesion (cell adhesion molecules, CAMs) are the driving force behind the tissue architecture (Gumbiner, 1996; Buckley et al., 1998; Hagios et al., 1998; Schock and Perrimon, 2002), whereas adhesion defects often occur during tumour formation and metastasis (Zetter, 1993; Janiszewska et al., 2020). Among CAMs, the calcium-dependent adhesion (cadherin) proteins mediate the direct cell–cell adhesion through homophilic binding of extracellular domains (Perez and Nelson, 2004; Nishiguchi et al., 2016). In this perspective, we focus on Epithelial cadherin (E-cadherin, E-cad), as in epithelial cells, it plays a major role in tissue formation and maintenance (Takeichi, 1977; Halbleib and Nelson, 2006; Nelson, 2008; Kaszak et al., 2020). In contrast, other cadherins such as P-cadherin do also contribute to cell–cell adhesion in epithelia but their actions are restricted to specific areas and developmental stages (Paredes et al., 2012).

The strength of adhesion correlates with the number of E-cad molecules on cell surfaces (Chu et al., 2004). Consequently, the regulation of E-cad surface levels in epithelia is instrumental in many processes, and even mild changes in E-cad levels profoundly affect many processes such as cell rearrangements, proliferation, and tissue architecture (Ciesiolka et al., 2004; Lecuit and Yap, 2015; Mohan et al., 2018; Greig and Bulgakova, 2020). Multiple mechanisms regulate E-cad surface levels including intracellular trafficking, transcriptional regulation, post-translational modifications, and protein degradation (Palacios et al., 2005; Bertocchi et al., 2012; Serrano-Gomez et al., 2016; Brüser and Bogdan, 2017; Cai et al., 2018). In this perspective, we discuss transcriptional regulation, as the decrease in E-cad protein levels often correlates with reduced mRNA abundance (Bringuier et al., 1999).

Transcriptional Repression

The best-studied regulation of E-cad transcription in mammalian cells is silencing by transcription factors (TFs) including SNAIL, SLUG (also known as SNAI1 and SNAI2 in mammals), ZEB1/2, and Twist1/2 (Figure 1A), all of which directly bind conserved E-boxes (CANNTG sequences) in the E-cad promoter (Giroldi et al., 1997; Comijn et al., 2001; Bolós et al., 2003; Wong et al., 2014; Vergara et al., 2016; Russell and Pranjol, 2018). All events of epithelial-to-mesenchymal transition (EMT) during development involve at least one of these TFs, and they were all linked to the tumour progression (Gasparotto et al., 2011; Muenst et al., 2013; da Silva et al., 2014; Vergara et al., 2016; Yang et al., 2020). Despite the overall similarity in the action of these TFs, their different affinities and variable expression allow for a dynamic but tightly regulated expression of E-cad (Bolós et al., 2003; Eger et al., 2005; Schmidt et al., 2005; Vesuna et al., 2008; Mazda et al., 2011). Regulation of E-cad transcription by these TFs is extremely conserved across evolution. While Drosophila and mammalian E-cad differ in their extracellular domains and are products of independent evolution from a common N-cadherin-like ancestor, transcription of the Drosophila E-cad (encoded by the shotgun gene) is also inhibited by the Snail and Twist TFs (Figure 1B; Oda et al., 1998, 2005; Oda and Takeichi, 2011; Nishiguchi et al., 2016). This could be either due to this regulation already being present in the ancestor or a result of the parallel evolution due to the same functional requirements. In Drosophila, while Snail represses E-cad during gastrulation in the embryo, it does not inhibit it in the adult midguts (Oda et al., 1998; Campbell et al., 2019), potentially due to the combinatorial action of other regulators (Figure 1B).

FIGURE 1.

FIGURE 1

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Molecular mechanisms regulating E-cadherin gene expression. (A,B) Summary of the best known negative and positive regulators of E-cad in mammals (A) and Drosophila (B). The expression of E-cad gene (shotgun, shg, in Drosophila) is controlled by transcription factors (ellipses), which inhibit (red) or promote it (green) and interact with the E-cad promoter with varying affinities. Several further proteins modify chromatin at the promoter (squircles) silencing the E-cad expression (red) or increasing it (green). These proteins cooperate, compete, and regulate each other, establishing the intricated network that fine-tune the expression of E-cad. In panel (B) the non-conserved proteins were removed while conserved maintained their relative positions but their names were replaced with those of the Drosophila orthologs. The regulatory roles of transparentized proteins in panels (A,B) were not demonstrated in the respective systems. (C) The proposed model of E-cad modulation of its expression. A subpool of STAT92E translocates into the nucleus independently of the canonical JAK/STAT signalling, where it interacts with heterochromatin protein 1 (HP1). Within euchromatin, HP1 localises at specific loci corresponding to gene promoters, including that of the shg gene. Concurrently, STAT92E is recruited to the cell surface by E-cad and its binding partner Par-3. Therefore, elevated levels of E-cad outcompete STAT92E from the nucleus inhibiting its function in heterochromatin formation and promoter regulation. As the result, elevated E-cad at the cell surface reduces shg expression, ultimately restoring its levels. It is unclear whether this is accompanied by a change in chromatin organisation around shg promoter, indicated by ‘?.’

Multiple epigenetic mechanisms also inhibit E-cad expression in mammalian cells. Hypermethylation of the large CpG island around the promoter is frequently linked with loss of E-cad expression (Yoshiura et al., 1995; van Roy and Berx, 2008). This DNA hypermethylation is often accompanied by histone modifications associated with inactive chromatin (Serrano-Gomez et al., 2016). Most notably, polycomb repressive complex 2 (PRC2) is recruited to the E-cad promoter where its key component, enhancer of zeste homolog 2 (EZH2), places the repressive H3K27me3 mark, deactivating E-cad transcription (Viré et al., 2006; Cao et al., 2008; Fujii and Ochiai, 2008). The histone methyltransferase G9a places the repressive H3K9me2/3 marks on the E-cad promoter (Dong et al., 2012). Histones are subjected to other post-translational modifications that contribute to E-cad silencing, such as the removal of the H3K4me2 mark by the lysine-specific demethylase 1 (LSD1) (Lin et al., 2010), or H3 and H4 deacetylation by the histone deacetylases 1 and 2 (HDAC1/2) (Peinado et al., 2004a). Histone phosphorylation and ubiquitination were also suggested to alter transcription of the mammalian E-cad (Surapaneni et al., 2020; Wang et al., 2020). While there is only a low level of DNA methylation in Drosophila (Lyko et al., 2000) other epigenetic mechanisms are conserved (Figure 1B) and might, similarly to Snail and Twist, regulate E-cad transcription in invertebrates.

These mechanisms of E-cad silencing closely interact with each other (Figure 1A). For example, SNAIL recruits the repressive complex containing LSD1 and HDAC1/2 (Peinado et al., 2004a; Lin et al., 2014), which may facilitate the subsequent recruitment of PRC2 (Ai et al., 2017; Jin et al., 2017, p. 1). SNAIL also recruits G9a to the E-cad promoter, whereas both G9a and PRC2 interact with the DNA methyltransferases DNMT1/3A/3B, and thus can promote the CpG methylation (Viré et al., 2006; Dong et al., 2012; Lin et al., 2014). Twist1 can also promote the recruitment of PRC2 (Cakouros et al., 2012; Malouf et al., 2013), whereas ZEB1 recruits histone deacetylases, HDAC1/2, and possibly DNMT3B (Aghdassi et al., 2012; Zhang et al., 2019). Concurrently, EZH2 facilitates the expression of SNAIL and SLUG through an unknown mechanism (Zhang et al., 2017), and increases Twist expression by suppressing the miR-361 (Ihira et al., 2017). Through these interactions, we suggest that individual mechanisms act together to ensure the robust silencing of the E-cad.

Transcriptional Activation

A fluctuation in the activity of a repressive mechanism, for example, due to random changes in fundamentally stochastic expression of regulatory genes, could lead to the E-cad loss (Raj and van Oudenaarden, 2008; Wendt et al., 2011). Protein turnover may counteract the resulting short-term E-cad silencing, whereby the balance between recycling and degradation of endocytosed molecules quickly modulates E-cad surface levels (Bryant and Stow, 2004; Huang et al., 2011; Bulgakova et al., 2013; Erami et al., 2015; Bulgakova and Brown, 2016). However, this might be insufficient to protect tissue integrity from a transient but excessive drop in the E-cad levels. Therefore, the robust E-cad expression requires mechanisms for transcriptional activation.

The traditional view is that E-cad expression is activated by constitutive factors in mammalian cells, which are overcome by repressors (Peinado et al., 2004b). This view was contested upon discovery of the binding of the TF FOXA/HNF3 (Forkhead in Drosophila) to the E-cad promoter, increasing its expression and driving re-epithelization of breast cancer cells (Liu et al., 2005). This activation system interacts with other E-cad regulatory factors and epigenetic machinery. FOXA suppresses the expression of SLUG, combining E-cad activation with the release of repression (Anzai et al., 2017). FOXA also interacts with p300, which promotes E-cad expression with the two p300-binding sites in the human E-cad promoter (Liu et al., 2005). This interaction suggests the potential role of histone modifications in E-cad transcriptional activation due to the intrinsic histone acetyltransferase (HAT) activity of p300 (Ogryzko et al., 1996; Chan and Thangue, 2001). While p300 can acetylate all histones, it preferentially modifies H3 and H4 and competes for binding with HDAC1 (Ogryzko et al., 1996; Li et al., 2014), making p300 an excellent candidate to counteract the histone deacetylation at the E-cad promoter by HDAC1/2. Finally, FOXA drives E-cad downregulation by interacting with Sp1 and RUNX1/AML1 (Liu et al., 2005; D’Costa et al., 2012). Of particular interest for the data presented below is that in mammalian cells RUNX1/AML1 forms a complex with SUV39H1 [Su(var)3–9 in Drosophila], whereas methylation of the H3K9 by SUV39H1 creates a binding site for the heterochromatin protein 1 (HP1) (Lachner et al., 2001; Chakraborty et al., 2003).

Retinoblastoma (Rb) and c-Myc also specifically activate the expression of E-cad promoter through an interaction with the TF AP-2, thus maintaining the epithelial phenotype in mammalian cells (Batsché et al., 1998). Rb interacts with the TF RUNX2, which acts redundantly with closely related RUNX1 in chondrocyte differentiation (Kimura et al., 2010; Gündüz et al., 2012; Komori, 2015). This, alongside RUNX2 and RUNX1 cooperation in tumour cells and RUNX1 recruitment to the E-cad promoter, raises a possibility of similar recruitment of RUNX2, and therefore Rb, to the E-cad promoter (Fowler et al., 2006). Concurrently, Rb depletion induces expression of SLUG and ZEB1, suggesting an indirect effect on E-cad expression (Arima et al., 2008). Additionally to these mechanisms, Rb is likely to modulate epigenetic mechanisms controlling E-cad expression, as it targets the HP1 to gene promoters (Nielsen et al., 2001). This complexity of E-cad transcriptional regulation is further amplified by the involvement of post-transcriptional mechanisms. For example, c-Myc activates E-cad expression, but its substantial overexpression in breast cancer cells leads to post-transcriptional repression of E-cad (Batsché et al., 1998; Cowling and Cole, 2007). While roles of Rb, c-Myc, and Forkhead in the expression of Drosophila E-cad are not known, another TF – Grhl2 (Grainyhead in Drosophila) – activates E-cad expression in both mammalian and invertebrate cells. In the former, intron-bound Grhl2 promotes E-cad transcription through chromatin looping, playing an essential role in determining an epithelial phenotype (Werth et al., 2010; Xiang et al., 2012). Similarly in flies, Grainyhead has four binding sites in the shotgun locus and its overexpression promotes E-cad transcription, as well as that of multiple other genes involved in epithelial maturation (Yao et al., 2017).

The above mechanisms overlook the potential of the cell adhesion molecules (CAMs) in triggering changes in gene expression (Robinson and Moberg, 2011; Dasgupta and McCollum, 2019; Hannezo and Heisenberg, 2019). For example, β-catenin is a TF activated by canonical Wnt signalling (Shang et al., 2017) and induces expression of SLUG in human carcinoma cells (Conacci-Sorrell et al., 2003), meaning that the release of β-catenin from junctions due to E-cad downregulation can reinforce this downregulation. Via α-catenin, E-cad regulates the Hippo pathway by sequestering the YAP protein outside the nucleus, thus explaining the antiproliferative properties of E-cad (Kim et al., 2011; Hannezo and Heisenberg, 2019; Sarpal et al., 2019). α-catenin counteracts β-catenin by inhibiting Wnt-activated transcription (Daugherty et al., 2014). Similarly, p120-catenin antagonises β-catenin-mediated transcriptional activation by interacting with the repressive TF Kaiso (Daniel and Reynolds, 1999; Kourtidis et al., 2013; Schackmann et al., 2013). Curiously, β-catenin expression is inhibited by the Kaiso–p120-catenin complex (Liu et al., 2014), highlighting the reciprocal interactions between transcriptional mechanisms and adhesion molecules. In Drosophila, although Kaiso is not present, p120-catenin is nevertheless involved in transcriptional regulation (Stefanatos et al., 2013). As catenins are recruited by E-cad, their availability for either performing nuclear functions or sequestering other TFs outside the nucleus is dependent on E-cad levels. Indeed, E-cad levels impact on transcription of multiple genes involved in a wide range of processes (Soncin et al., 2011). Furthermore, β-catenin recruits the p300 protein, which promotes E-cad expression (Mosimann et al., 2009). This raises the questions of whether E-cad contributes to own gene expression.

A Feedback Loop Regulating E-Cad Expression Through Par-3, HP1, and STAT92E

The polarity determinant Par-3 (Bazooka in Drosophila) is an evolutionarily conserved protein recruited to E-cad adhesion in both mammals and Drosophila (Harris and Peifer, 2005; Wei et al., 2005; Iden et al., 2006; Wang et al., 2009; Bulgakova et al., 2013; Xue et al., 2013). Par-3 has six protein–protein interaction domains, making it a versatile scaffold for protein recruitment (McKinley et al., 2012). In Drosophila, one of the proteins recruited by Par-3 is the only fly member of the signal transducer and activator of transcription (STAT) protein family, STAT92E (Hou et al., 1996; Yan et al., 1996). This recruitment of STAT92E is necessary for the efficient Janus kinase (JAK)/STAT signalling in Drosophila epithelial cells (Sotillos et al., 2008). In human and mouse cells, the loss of Par-3 activates STAT3 signalling, although it is not known if the loss of STAT3 recruitment to E-cad adhesion sites by Par-3 contributes to this activation (McCaffrey et al., 2012; Guyer and Macara, 2015). However, this is very likely, as the loss of E-cad adhesion also leads to STAT3 activation (del Valle et al., 2013).

The canonical signalling pathway comprises STAT translocation into the nucleus after its phosphorylation by the ligand-activated JAK. However, there is growing evidence of non-canonical signalling in both Drosophila and mammals, whereby non-phosphorylated STAT stabilises heterochromatin by binding HP1 (Li, 2008; Hixson et al., 2019). HP1 is an evolutionarily conserved non-histone chromatin protein whose best-characterised role is the formation and propagation of heterochromatin (Lomberk et al., 2006; Dialynas et al., 2008; Xu et al., 2014). HP1 is recruited to H3K9me2/3 through SUV39H1, and then itself recruits more SUV39H1 molecules, propagating H3K9 methylation (Lomberk et al., 2006). This is accompanied by DNA methylation in mammalian cells, as both HP1 and SUV39H1 bind DNMTs (Fuks et al., 2003). HP1 was also linked to other functions including transcriptional regulation, where HP1 usually represses transcription by facilitating methylation of H3K9. However, HP1 also localises at specific loci within euchromatin (Dialynas et al., 2008), and there is growing evidence of its association with transcriptionally active regions from C. elegans to Drosophila and humans (Lu et al., 2000; Piacentini et al., 2003; Mateescu et al., 2008; McMurchy et al., 2017). In Drosophila, the active euchromatic regions with HP1 recruitment include developmentally-regulated and heat-shock genes, whose activity correlates with HP1 dosage (Piacentini et al., 2003). Non-phosphorylated STAT92E and HP1 co-localise at euchromatic regions of Drosophila polytene chromosomes (Shi et al., 2008). HP1 disperses from chromosomes following hyperactivation of the canonical signalling, suggesting an intricate cross-talk between the two modes of signalling, likely through the availability of non-phosphorylated STAT92E (Xu et al., 2014).

As STAT92E promotes heterochromatin gene silencing in a dose-dependent manner (Shi et al., 2008), we hypothesised that an increase of E-cad levels would lead to STAT92E sequestration at adhesion sites, making it unavailable to bind HP1 and promote heterochromatin formation alongside other functions (Figure 1C). We examined the effects of E-cad overexpression on heterochromatic gene silencing monitored using position-effect variegation (PEV). In this assay, a gene is translocated into a peri-heterochromatic region where its expression is silenced in a stochastic pattern (Figure 2A). This silencing depends on the spreading of heterochromatin: if a protein normally promotes heterochromatin formation, its loss would reduce the amount of heterochromatin and suppress the variegation (=loss of silencing of the translocated gene, Figure 2A; Elgin and Reuter, 2013). We overexpressed E-cad in the Drosophila eye using the UAS–Gal4 system with GMR::Gal4, which expresses Gal4 in all retinal cells (Hay et al., 1997). We used the variegating allele of the brown gene (bwPEV), necessary to produce the red pigment pteridine. Overexpression of E-cad increased the normalised eye pigmentation with pteridine to 104% from 69% in the control (Figures 2B,C and Supplementary Figure 1). Therefore, E-cad suppresses PEV in the eye, supporting that elevated E-cad inhibits heterochromatin formation.

FIGURE 2.

FIGURE 2

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The crosstalk between E-cadherin, STAT92E and HP1. (A) A diagram of position effect variegation (PEV) in relation to chromatin compaction. (B,C) E-cad overexpression suppresses PEV: examples (B) and quantification (C). Scale bar – 0.2 mm, ***p = 0.0001 (unpaired t-test). (D) Overlapping box-and-violin plots showing the quantification of rBEADS normalised HP1 signal on larval-specific (red) and common (blue) promoters. The p-value above – the significance calculated using U-test. The value below the boxplots – the mean of the signal. (E) The distribution of the rBEADS normalised HP1 signal over transcription start sites (TSS) loci of larval-specific (red) and common (blue) genes. The vertical grey line represents the location of TSS, the plots span 1 kb upstream and downstream from annotated TSS. (F) A Venn diagram showing the overlaps between peak loci in the larval dataset and two independent embryonic (16–24 h) replicates. All three peak sets were filtered from peaks overlapping centromeric heterochromatin clusters and non-mappable, repetitive regions, defined based on GEM mappability analyses. modENCODE peak calls have been lifted from dm3 to dm6 genome assembly to match the analyses of larval HP1 peaks. The IDR method with 0.05 p-vale threshold was used to collapse the replicates to a common, high confidence peak set. (G) shotgun expression following overexpression of non-phosphorylatable STAT92EY704F measured using RT-qPCR and normalised to the expression of house-keeping gene RpL32. Four biological replicates are shown as distinct dots with three technical replicates each. *p = 0.02 (one-sample t-test comparing to 1). (H–J) E-cad-GFP expressed from the endogenous promoter is downregulated following E-cad overexpression (E-cad OS) from a heterologous UAS promoter. A representative image (H) shows endogenously expressed E-cad-GFP visualised with native GFP fluorescence (green, left; grey, middle) and total E-cad visualised with antibody staining (magenta, left; grey, right), scale bar – 10 μm. Levels of endogenously expressed E-cad-GFP are quantified in panel (I) and total E-cad levels in panel (J). **p = 0.0018 and ***p = 0.0003 (paired t-test).

Next, we sought to determine what genes might be regulated through our hypothetical mechanism and examined the HP1 localisation using publicly available ChIP-seq datasets generated using third instar larvae (Riddle et al., 2011), which include epithelial monolayers of imaginal discs. There were distinct peaks of HP1 overlapping with promoters of 456 genes outside of heterochromatin clusters and non-mappable (non-unique, repeatable) regions (Figures 2D,E and Supplementary Figure 2). We compared the larval HP1 peaks with two independent replicates from an earlier developmental stage – embryos at 16–24 h of development (modENCODE Consortium, Roy et al., 2010). Roughly half of the HP1 peaks in the larval dataset (155 peaks mapping to 151 loci) were common to all three datasets, while another half (187 peaks/loci) were larval-specific (Figure 2F). The common peaks (potentially, developmentally conserved) mostly mapped to bi-directional promoters, so that 155 peaks mapped to a total of 280 genes. These genes are enriched for housekeeping terms: regulation of primary and macromolecule metabolic processes (Supplementary Figure 3, terms 7–11); and mitotic cell cycle and cell cycle regulation (Supplementary Figure 3, terms 2–6). In contrast, the larval-specific peaks, which map to promoters of 178 genes, are enriched for terms related to epithelial tissue: morphogenesis of epithelium, actin filament organisation, epithelium development and imaginal disc development (Supplementary Figure 3, terms 19–24). The shotgun promoter overlapped a larval-specific HP1 peak with a signal value of 9.83 (207th promoter rank). This peak is consistent with two potential mechanisms for HP1 recruitment – via Su(var)3–9 or one of the two fly Rb-family proteins. Therefore, the modulation of HP1 function downstream of E-cad and non-canonical STAT signalling might alter the E-cad expression.

Indeed, overexpression of the non-phosphorylatable STAT92E (STAT92EY704F), which acts only in the non-canonical signalling (Karsten et al., 2006; Xu et al., 2014), increased expression of the shotgun gene (Figure 2G). Such an increase is consistent with the model whereby an unprogrammed drop in E-cad levels releases STAT92E, which leads to a consequent increase of E-cad expression and thus restores E-cad levels (Figure 1C). We observed that overexpression of E-cad from a heterologous UAS promoter leads to downregulation of the protein expressed from the endogenous locus (Figures 2H–J). This might be accompanied by simultaneous upregulation of the canonical JAK/STAT signalling as observed for STAT3 activation in some human cancers (Loh et al., 2019). We postulate that other genes with larval-specific HP1 peaks behave similarly to E-cad. Then, the activation of non-canonical STAT signalling following a stochastic drop in E-cad levels would reinforce the epithelial nature of the cells as many of these genes are involved in epithelial morphogenesis.

Outlook/Perspective

E-cad levels at the cell surface play an important role in determining cellular properties: cell shape, response to signalling, cell division, and neighbour exchange to name just a few (Lecuit and Lenne, 2007; Rübsam et al., 2017; Mendonsa et al., 2018). Not surprisingly, E-cad expression is subjected to complex regulation by multiple interconnected mechanisms. However, to respond to the cell’s needs and modulate E-cad levels accordingly, information about the E-cad levels at the surface must be transferred to the nucleus. Here, we propose one such mechanism, whereby surface E-cad feeds back to the nucleus through competition for STAT92E binding. We hypothesise that this feedback stabilises E-cad transcription enabling robust and resilient cell–cell adhesion. Importantly, this provides the cell with a tumour-suppressive mechanism, as heterochromatin formation, which is likely to follow a drop in E-cad levels, is linked with cellular senescence (Narita et al., 2003). This contrasts another feedback mechanism whereby the release of β-catenin from adhesion sites leads to SLUG expression and reinforces E-cad silencing in human cells. Therefore, depending on the context E-cad expression may be stabilised or silenced through feedback interactions. It is tempting to speculate that further mechanisms of information transfer from the cell surface to the nucleus exist – not unlike the multiple and intertwined mechanisms for regulation of E-cad transcription. A comprehensive understanding of E-cad transcriptional regulation requires the discovery and in-depth analysis of such mechanisms.

Data Availability Statement

The datasets presented in this study can be found in online repositories. The names of the repository/repositories and accession number(s) can be found below: http://data.modencode.org/, 3187; http://data.modencode.org/, 3188; http://data.modencode.org/, 3391; http://data.modencode.org/, 3392.

Author Contributions

NAB and MRM performed the experiments and wrote the manuscript. NAB and PAS did the bioinformatics analyses. All authors contributed to the article and approved the submitted version.

Conflict of Interest

PAS was a shareholder and the managing director of the company SmartImmune Ltd. The remaining authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Acknowledgments

We thank Martin Zeidler for STAT92EY704F flies and the University of Sheffield’s Wolfson Light Microscopy and Drosophila Facillities for their technical support.

Footnotes

Funding. This work was supported by a grant from the UKRI BBSRC (BB/P007503/1) to NAB.

Supplementary Material

The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fcell.2021.701175/full#supplementary-material

Click here for additional data file. (1.7MB, pdf)

References

  1. Aghdassi A., Sendler M., Guenther A., Mayerle J., Behn C.-O., Heidecke C.-D., et al. (2012). Recruitment of histone deacetylases HDAC1 and HDAC2 by the transcriptional repressor ZEB1 downregulates E-cadherin expression in pancreatic cancer. Gut 61 439–448. 10.1136/gutjnl-2011-300060 [DOI] [PubMed] [Google Scholar]
  2. Ai S., Peng Y., Li C., Gu F., Yu X., Yue Y., et al. (2017). EED orchestration of heart maturation through interaction with HDACs is H3K27me3-independent. eLife 6:e24570. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Anzai E., Hirata K., Shibazaki M., Yamada C., Morii M., Honda T., et al. (2017). FOXA1 induces E-Cadherin expression at the protein level via suppression of slug in epithelial breast cancer cells. Biol. Pharm. Bull. 40 1483–1489. 10.1248/bpb.b17-00307 [DOI] [PubMed] [Google Scholar]
  4. Arima Y., Inoue Y., Shibata T., Hayashi H., Nagano O., Saya H., et al. (2008). Rb depletion results in deregulation of E-cadherin and induction of cellular phenotypic changes that are characteristic of the epithelial-to-mesenchymal transition. Cancer Res. 68 5104–5112. 10.1158/0008-5472.can-07-5680 [DOI] [PubMed] [Google Scholar]
  5. Batsché E., Muchardt C., Behrens J., Hurst H. C., Crémisi C. (1998). RB and c-Myc activate expression of the E-cadherin gene in epithelial cells through interaction with transcription factor AP-2. Mol. Cell. Biol. 18 3647–3658. 10.1128/mcb.18.7.3647 [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Bertocchi C., Vaman Rao M., Zaidel-Bar R. (2012). Regulation of adherens junction dynamics by phosphorylation switches. J. Signal. Transduct. 2012:125295. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Bolós V., Peinado H., Pérez-Moreno M. A., Fraga M. F., Esteller M., Cano A. (2003). The transcription factor Slug represses E-cadherin expression and induces epithelial to mesenchymal transitions: a comparison with Snail and E47 repressors. J. Cell Sci. 116 499–511. 10.1242/jcs.00224 [DOI] [PubMed] [Google Scholar]
  8. Bringuier P. P., Giroldi L. A., Umbas R., Shimazui T., Schalken J. A. (1999). Mechanisms associated with abnormal E-cadherin immunoreactivity in human bladder tumors. Int. J. Cancer 83 591–595. [DOI] [PubMed] [Google Scholar]
  9. Brüser L., Bogdan S. (2017). Adherens junctions on the move—membrane trafficking of E-Cadherin. Cold Spring Harb. Perspect. Biol. 9:a029140. 10.1101/cshperspect.a029140 [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Bryant D. M., Stow J. L. (2004). The ins and outs of E-cadherin trafficking. Trends Cell Biol. 14 427–434. 10.1016/j.tcb.2004.07.007 [DOI] [PubMed] [Google Scholar]
  11. Buckley C. D., Rainger G. E., Bradfield P. F., Nash G. B., Simmons D. L. (1998). Cell adhesion: more than just glue (Review). Mol. Membr. Biol. 15 167–176. 10.3109/09687689709044318 [DOI] [PubMed] [Google Scholar]
  12. Bulgakova N. A., Brown N. H. (2016). Drosophila p120-catenin is crucial for endocytosis of the dynamic E-cadherin-Bazooka complex. J. Cell Sci. 129 477–482. [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Bulgakova N. A., Grigoriev I., Yap A. S., Akhmanova A., Brown N. H. (2013). Dynamic microtubules produce an asymmetric E-cadherin-Bazooka complex to maintain segment boundaries. J. Cell Biol. 201 887–901. 10.1083/jcb.201211159 [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Cai J., Culley M. K., Zhao Y., Zhao J. (2018). The role of ubiquitination and deubiquitination in the regulation of cell junctions. Protein Cell 9 754–769. 10.1007/s13238-017-0486-3 [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Cakouros D., Isenmann S., Cooper L., Zannettino A., Anderson P., Glackin C., et al. (2012). Twist-1 Induces Ezh2 recruitment regulating histone methylation along the Ink4A/Arf locus in mesenchymal stem cells. Mol. Cell. Biol. 32 1433–1441. 10.1128/mcb.06315-11 [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Campbell K., Rossi F., Adams J., Pitsidianaki I., Barriga F. M., Garcia-Gerique L., et al. (2019). Collective cell migration and metastases induced by an epithelial-to-mesenchymal transition in Drosophila intestinal tumors. Nat. Commun. 10:2311. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Cao Q., Yu J., Dhanasekaran S. M., Kim J. H., Mani R.-S., Tomlins S. A., et al. (2008). Repression of E-cadherin by the polycomb group protein EZH2 in cancer. Oncogene 27 7274–7284. 10.1038/onc.2008.333 [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Chakraborty S., Sinha K. K., Senyuk V., Nucifora G. (2003). SUV39H1 interacts with AML1 and abrogates AML1 transactivity. AML1 is methylated in vivo. Oncogene 22 5229–5237. 10.1038/sj.onc.1206600 [DOI] [PubMed] [Google Scholar]
  19. Chan H. M., Thangue N. B. L. (2001). p300/CBP proteins: HATs for transcriptional bridges and scaffolds. J. Cell Sci. 114 2363–2373. 10.1242/jcs.114.13.2363 [DOI] [PubMed] [Google Scholar]
  20. Chu Y.-S., Thomas W. A., Eder O., Pincet F., Perez E., Thiery J. P., et al. (2004). Force measurements in E-cadherin–mediated cell doublets reveal rapid adhesion strengthened by actin cytoskeleton remodeling through Rac and Cdc42. J. Cell Biol. 167 1183–1194. 10.1083/jcb.200403043 [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Ciesiolka M., Delvaeye M., Imschoot G. V., Verschuere V., McCrea P., Roy F., et al. (2004). p120 catenin is required for morphogenetic movements involved in the formation of the eyes and the craniofacial skeleton in Xenopus. J. Cell Sci. 117 4325–4339. 10.1242/jcs.01298 [DOI] [PubMed] [Google Scholar]
  22. Comijn J., Berx G., Vermassen P., Verschueren K., van Grunsven L., Bruyneel E., et al. (2001). The two-handed e box binding zinc finger protein SIP1 downregulates E-Cadherin and induces invasion. Mol. Cell 7 1267–1278. 10.1016/s1097-2765(01)00260-x [DOI] [PubMed] [Google Scholar]
  23. Conacci-Sorrell M., Simcha I., Ben-Yedidia T., Blechman J., Savagner P., Ben-Ze’ev A. (2003). Autoregulation of E-cadherin expression by cadherin-cadherin interactions: the roles of beta-catenin signaling. Slug, and MAPK. J. Cell Biol. 163 847–857. 10.1083/jcb.200308162 [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Cowling V. H., Cole M. D. (2007). E-cadherin repression contributes to c-Myc-induced epithelial cell transformation. Oncogene 26 3582–3586. 10.1038/sj.onc.1210132 [DOI] [PubMed] [Google Scholar]
  25. D’Costa Z. J., Jolly C., Androphy E. J., Mercer A., Matthews C. M., Hibma M. H. (2012). Transcriptional repression of E-Cadherin by human papillomavirus Type 16 E6. PLoS One 7:e48954. 10.1371/journal.pone.0048954 [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. da Silva S. D., Alaoui-Jamali M. A., Soares F. A., Carraro D. M., Brentani H. P., Hier M., et al. (2014). TWIST1 is a molecular marker for a poor prognosis in oral cancer and represents a potential therapeutic target. Cancer 120 352–362. 10.1002/cncr.28404 [DOI] [PubMed] [Google Scholar]
  27. Daniel J. M., Reynolds A. B. (1999). The Catenin p120ctn interacts with Kaiso, a Novel BTB/POZ domain zinc finger transcription factor. Mol. Cell Biol. 19 3614–3623. 10.1128/mcb.19.5.3614 [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Dasgupta I., McCollum D. (2019). Control of cellular responses to mechanical cues through YAP/TAZ regulation. J. Biol. Chem. 294 17693–17706. 10.1074/jbc.rev119.007963 [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Daugherty R. L., Serebryannyy L., Yemelyanov A., Flozak A. S., Yu H.-J., Kosak S. T., et al. (2014). α-Catenin is an inhibitor of transcription. Proc. Natl. Acad. Sci. U. S. A. 111 5260–5265. [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. del Valle I., Rudloff S., Carles A., Li Y., Liszewska E., Vogt R., et al. (2013). E-cadherin is required for the proper activation of the Lifr/Gp130 signaling pathway in mouse embryonic stem cells. Development 140 1684–1692. 10.1242/dev.088690 [DOI] [PubMed] [Google Scholar]
  31. Dialynas G. K., Vitalini M. W., Wallrath L. L. (2008). Linking heterochromatin protein 1 (HP1) to cancer progression. Mutat. Res. 647 13–20. 10.1016/j.mrfmmm.2008.09.007 [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Dong C., Wu Y., Yao J., Wang Y., Yu Y., Rychahou P. G., et al. (2012). G9a interacts with Snail and is critical for Snail-mediated E-cadherin repression in human breast cancer. J. Clin. Invest. 122 1469–1486. 10.1172/jci57349 [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Eger A., Aigner K., Sonderegger S., Dampier B., Oehler S., Schreiber M., et al. (2005). DeltaEF1 is a transcriptional repressor of E-cadherin and regulates epithelial plasticity in breast cancer cells. Oncogene 24 2375–2385. 10.1038/sj.onc.1208429 [DOI] [PubMed] [Google Scholar]
  34. Elgin S. C. R., Reuter G. (2013). Position-effect variegation, heterochromatin formation, and gene silencing in Drosophila. Cold Spring Harb. Perspect. Biol. 5:a017780. 10.1101/cshperspect.a017780 [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Erami Z., Timpson P., Yao W., Zaidel-Bar R., Anderson K. I. (2015). There are four dynamically and functionally distinct populations of E-cadherin in cell junctions. Biol. Open 4 1481–1489. 10.1242/bio.014159 [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Fowler M., Borazanci E., McGhee L., Pylant S. W., Williams B. J., Glass J., et al. (2006). RUNX1 (AML-1) and RUNX2 (AML-3) cooperate with prostate-derived Ets factor to activate transcription from the PSA upstream regulatory region. J. Cell. Biochem. 97 1–17. 10.1002/jcb.20664 [DOI] [PubMed] [Google Scholar]
  37. Fujii S., Ochiai A. (2008). Enhancer of zeste homolog 2 downregulates E-cadherin by mediating histone H3 methylation in gastric cancer cells. Cancer Sci. 99 738–746. 10.1111/j.1349-7006.2008.00743.x [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Fuks F., Hurd P. J., Deplus R., Kouzarides T. (2003). The DNA methyltransferases associate with HP1 and the SUV39H1 histone methyltransferase. Nucleic Acids Res. 31 2305–2312. 10.1093/nar/gkg332 [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Gasparotto D., Polesel J., Marzotto A., Colladel R., Piccinin S., Modena P., et al. (2011). Overexpression of TWIST2 correlates with poor prognosis in head and neck squamous cell carcinomas. Oncotarget 2 1165–1175. 10.18632/oncotarget.390 [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Giroldi L. A., Bringuier P.-P., de Weijert M., Jansen C., van Bokhoven A., Schalken J. A. (1997). Role of E boxes in the repression of E-cadherin expression. Biochem. Biophys. Res. Commun. 241 453–458. 10.1006/bbrc.1997.7831 [DOI] [PubMed] [Google Scholar]
  41. Greig J., Bulgakova N. A. (2020). Interplay between actomyosin and E-cadherin dynamics regulates cell shape in the Drosophila embryonic epidermis. J. Cell Sci. 133:jcs242321. [DOI] [PubMed] [Google Scholar]
  42. Gumbiner B. M. (1996). Cell adhesion: the molecular basis of tissue architecture and morphogenesis. Cell 84 345–357. 10.1016/s0092-8674(00)81279-9 [DOI] [PubMed] [Google Scholar]
  43. Gündüz V., Kong E., Bryan C. D., Hinds P. W. (2012). Loss of the retinoblastoma tumor suppressor protein in murine calvaria facilitates immortalization of osteoblast-adipocyte bipotent progenitor cells characterized by low expression of N-Cadherin. Mol. Cell Biol. 32 2561–2569. 10.1128/mcb.06453-11 [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Guyer R. A., Macara I. G. (2015). Loss of the polarity protein PAR3 activates STAT3 signaling via an atypical protein Kinase C (aPKC)/NF-κB/Interleukin-6 (IL-6) Axis in mouse mammary cells. J. Biol. Chem. 290 8457–8468. 10.1074/jbc.m114.621011 [DOI] [PMC free article] [PubMed] [Google Scholar]
  45. Hagios C., Lochter A., Bissell M. J. (1998). Tissue architecture: the ultimate regulator of epithelial function? Philos. Trans. R Soc. Lond. B Biol. Sci. 353 857–870. 10.1098/rstb.1998.0250 [DOI] [PMC free article] [PubMed] [Google Scholar]
  46. Halbleib J. M., Nelson W. J. (2006). Cadherins in development: cell adhesion, sorting, and tissue morphogenesis. Genes Dev. 20 3199–3214. 10.1101/gad.1486806 [DOI] [PubMed] [Google Scholar]
  47. Hannezo E., Heisenberg C.-P. (2019). Mechanochemical feedback loops in development and disease. Cell 178 12–25. 10.1016/j.cell.2019.05.052 [DOI] [PubMed] [Google Scholar]
  48. Harris T. J. C., Peifer M. (2005). The positioning and segregation of apical cues during epithelial polarity establishment in Drosophila. J. Cell Biol. 170 813–823. 10.1083/jcb.200505127 [DOI] [PMC free article] [PubMed] [Google Scholar]
  49. Hay B. A., Maile R., Rubin G. M. (1997). P element insertion-dependent gene activation in the Drosophila eye. PNAS 94 5195–5200. 10.1073/pnas.94.10.5195 [DOI] [PMC free article] [PubMed] [Google Scholar]
  50. Hixson K. M., Cogswell M., Brooks-Kayal A. R., Russek S. J. (2019). Evidence for a non-canonical JAK/STAT signaling pathway in the synthesis of the brain’s major ion channels and neurotransmitter receptors. BMC Genom. 20:677. [DOI] [PMC free article] [PubMed] [Google Scholar]
  51. Hou X. S., Melnick M. B., Perrimon N. (1996). Marelle acts downstream of the Drosophila HOP/JAK kinase and encodes a protein similar to the mammalian STATs. Cell 84 411–419. 10.1016/s0092-8674(00)81286-6 [DOI] [PubMed] [Google Scholar]
  52. Huang J., Huang L., Chen Y.-J., Austin E., Devor C. E., Roegiers F., et al. (2011). Differential regulation of adherens junction dynamics during apical–basal polarization. J. Cell Sci. 124 4001–4013. 10.1242/jcs.086694 [DOI] [PMC free article] [PubMed] [Google Scholar]
  53. Iden S., Rehder D., August B., Suzuki A., Wolburg-Buchholz K., Wolburg H., et al. (2006). A distinct PAR complex associates physically with VE-cadherin in vertebrate endothelial cells. EMBO Rep. 7 1239–1246. 10.1038/sj.embor.7400819 [DOI] [PMC free article] [PubMed] [Google Scholar]
  54. Ihira K., Dong P., Xiong Y., Watari H., Konno Y., Hanley S. J., et al. (2017). EZH2 inhibition suppresses endometrial cancer progression via miR-361/Twist axis. Oncotarget 8 13509–13520. 10.18632/oncotarget.14586 [DOI] [PMC free article] [PubMed] [Google Scholar]
  55. Janiszewska M., Primi M. C., Izard T. (2020). Cell adhesion in cancer: beyond the migration of single cells. J. Biol. Chem. 295 2495–2505. 10.1074/jbc.rev119.007759 [DOI] [PMC free article] [PubMed] [Google Scholar]
  56. Jin Y., Huo B., Fu X., Hao T., Zhang Y., Guo Y., et al. (2017). LSD1 collaborates with EZH2 to regulate expression of interferon-stimulated genes. Biomed. Pharmacother. 88 728–737. 10.1016/j.biopha.2017.01.055 [DOI] [PubMed] [Google Scholar]
  57. Karsten P., Plischke I., Perrimon N., Zeidler M. P. (2006). Mutational analysis reveals separable DNA binding and trans-activation of Drosophila STAT92E. Cell. Signal. 18 819–829. 10.1016/j.cellsig.2005.07.006 [DOI] [PubMed] [Google Scholar]
  58. Kaszak I., Witkowska-Piłaszewicz O., Niewiadomska Z., Dworecka-Kaszak B., Ngosa Toka F., Jurka P. (2020). Role of cadherins in cancer-a review. Int. J. Mol. Sci. 21:7624. 10.3390/ijms21207624 [DOI] [PMC free article] [PubMed] [Google Scholar]
  59. Kim N.-G., Koh E., Chen X., Gumbiner B. M. (2011). E-cadherin mediates contact inhibition of proliferation through Hippo signaling-pathway components. PNAS 108 11930–11935. 10.1073/pnas.1103345108 [DOI] [PMC free article] [PubMed] [Google Scholar]
  60. Kimura A., Inose H., Yano F., Fujita K., Ikeda T., Sato S., et al. (2010). Runx1 and Runx2 cooperate during sternal morphogenesis. Development 137 1159–1167. 10.1242/dev.045005 [DOI] [PMC free article] [PubMed] [Google Scholar]
  61. Komori T. (2015). The functions of Runx family transcription factors and Cbfb in skeletal development. Oral Sci. Int. 12 83–93. [Google Scholar]
  62. Kourtidis A., Ngok S. P., Anastasiadis P. Z. (2013). p120 catenin: an essential regulator of cadherin stability, adhesion-induced signaling, and cancer progression. Prog. Mol. Biol. Transl. Sci. 116 409–432. [DOI] [PMC free article] [PubMed] [Google Scholar]
  63. Lachner M., O’Carroll D., Rea S., Mechtler K., Jenuwein T. (2001). Methylation of histone H3 lysine 9 creates a binding site for HP1 proteins. Nature 410 116–120. 10.1038/35065132 [DOI] [PubMed] [Google Scholar]
  64. Lecuit T., Lenne P.-F. (2007). Cell surface mechanics and the control of cell shape, tissue patterns and morphogenesis. Nat. Rev. Mol. Cell Biol. 8 633–644. 10.1038/nrm2222 [DOI] [PubMed] [Google Scholar]
  65. Lecuit T., Yap A. S. (2015). E-cadherin junctions as active mechanical integrators in tissue dynamics. Nat. Cell Biol. 17 533–539. 10.1038/ncb3136 [DOI] [PubMed] [Google Scholar]
  66. Li W. X. (2008). Canonical and non-canonical JAK–STAT signaling. Trends Cell Biol. 18 545–551. 10.1016/j.tcb.2008.08.008 [DOI] [PMC free article] [PubMed] [Google Scholar]
  67. Li X., Yang H., Huang S., Qiu Y. (2014). Histone deacetylase 1 and p300 can directly associate with chromatin and compete for binding in a mutually exclusive manner. PLoS One 9:e94523. 10.1371/journal.pone.0094523 [DOI] [PMC free article] [PubMed] [Google Scholar]
  68. Lin T., Ponn A., Hu X., Law B. K., Lu J. (2010). Requirement of the histone demethylase LSD1 in snai1-mediated transcriptional repression during epithelial-mesenchymal transition. Oncogene 29 4896–4904. 10.1038/onc.2010.234 [DOI] [PMC free article] [PubMed] [Google Scholar]
  69. Lin Y., Dong C., Zhou B. P. (2014). Epigenetic regulation of EMT: the snail story. Curr. Pharm. Des. 20 1698–1705. 10.2174/13816128113199990512 [DOI] [PMC free article] [PubMed] [Google Scholar]
  70. Liu Y., Dong Q.-Z., Wang S., Xu H.-T., Miao Y., Wang L., et al. (2014). Kaiso interacts with p120-catenin to regulate β-catenin expression at the transcriptional level. PLoS One 9:e87537. 10.1371/journal.pone.0087537 [DOI] [PMC free article] [PubMed] [Google Scholar]
  71. Liu Y.-N., Lee W.-W., Wang C.-Y., Chao T.-H., Chen Y., Chen J. H. (2005). Regulatory mechanisms controlling human E-cadherin gene expression. Oncogene 24 8277–8290. 10.1038/sj.onc.1208991 [DOI] [PubMed] [Google Scholar]
  72. Loh C.-Y., Chai J. Y., Tang T. F., Wong W. F., Sethi G., Shanmugam M. K., et al. (2019). The E-Cadherin and N-Cadherin switch in epithelial-to-mesenchymal transition: signaling, therapeutic implications, and challenges. Cells 8:1118. 10.3390/cells8101118 [DOI] [PMC free article] [PubMed] [Google Scholar]
  73. Lomberk G., Wallrath L., Urrutia R. (2006). The heterochromatin protein 1 family. Genome Biol. 7:228. [DOI] [PMC free article] [PubMed] [Google Scholar]
  74. Lu B. Y., Emtage P. C., Duyf B. J., Hilliker A. J., Eissenberg J. C. (2000). Heterochromatin protein 1 is required for the normal expression of two heterochromatin genes in Drosophila. Genetics 155 699–708. 10.1093/genetics/155.2.699 [DOI] [PMC free article] [PubMed] [Google Scholar]
  75. Lyko F., Ramsahoye B. H., Jaenisch R. (2000). DNA methylation in Drosophila melanogaster. Nature 408 538–540. 10.1038/35046205 [DOI] [PubMed] [Google Scholar]
  76. Malouf G. G., Taube J. H., Lu Y., Roysarkar T., Panjarian S., Estecio M. R., et al. (2013). Architecture of epigenetic reprogramming following Twist1-mediated epithelial-mesenchymal transition. Genome Biol. 14:R144. [DOI] [PMC free article] [PubMed] [Google Scholar]
  77. Mateescu B., Bourachot B., Rachez C., Ogryzko V., Muchardt C. (2008). Regulation of an inducible promoter by an HP1beta-HP1gamma switch. EMBO Rep. 9 267–272. 10.1038/embor.2008.1 [DOI] [PMC free article] [PubMed] [Google Scholar]
  78. Mazda M., Nishi K., Naito Y., Ui-Tei K. (2011). E-Cadherin is transcriptionally activated via suppression of ZEB1 transcriptional repressor by small RNA-mediated gene silencing. PLoS One 6:e28688. 10.1371/journal.pone.0028688 [DOI] [PMC free article] [PubMed] [Google Scholar]
  79. McCaffrey L. M., Montalbano J., Mihai C., Macara I. G. (2012). Loss of the Par3 polarity protein promotes breast tumorigenesis and metastasis. Cancer Cell 22 601–614. 10.1016/j.ccr.2012.10.003 [DOI] [PMC free article] [PubMed] [Google Scholar]
  80. McKinley R. F. A., Yu C. G., Harris T. J. C. (2012). Assembly of Bazooka polarity landmarks through a multifaceted membrane-association mechanism. J. Cell Sci. 125 1177–1190. 10.1242/jcs.091884 [DOI] [PubMed] [Google Scholar]
  81. McMurchy A. N., Stempor P., Gaarenstroom T., Wysolmerski B., Dong Y., Aussianikava D., et al. (2017). A team of heterochromatin factors collaborates with small RNA pathways to combat repetitive elements and germline stress. eLife 6:e21666. [DOI] [PMC free article] [PubMed] [Google Scholar]
  82. Mendonsa A., Na T.-Y., Gumbiner B. M. (2018). E-cadherin in contact inhibition and cancer. Oncogene 37 4769–4780. 10.1038/s41388-018-0304-2 [DOI] [PMC free article] [PubMed] [Google Scholar]
  83. modENCODE Consortium, Roy S., Ernst J., Kharchenko P. V., Kheradpour P., Negre N., et al. (2010). Identification of functional elements and regulatory circuits by Drosophila modENCODE. Science 330 1787–1797. [DOI] [PMC free article] [PubMed] [Google Scholar]
  84. Mohan A., Schlue K. T., Kniffin A. F., Mayer C. R., Duke A. A., Narayanan V., et al. (2018). Spatial proliferation of epithelial cells is regulated by E-Cadherin force. Biophys. J. 115 853–864. 10.1016/j.bpj.2018.07.030 [DOI] [PMC free article] [PubMed] [Google Scholar]
  85. Mosimann C., Hausmann G., Basler K. (2009). β-Catenin hits chromatin: regulation of Wnt target gene activation. Nat. Rev. Mol. Cell Biol. 10 276–286. 10.1038/nrm2654 [DOI] [PubMed] [Google Scholar]
  86. Muenst S., Däster S., Obermann E. C., Droeser R. A., Weber W. P., von Holzen U., et al. (2013). Nuclear expression of snail is an independent negative prognostic factor in human breast cancer. Dis. Markers 35 337–344. 10.1155/2013/902042 [DOI] [PMC free article] [PubMed] [Google Scholar]
  87. Narita M., Nunez S., Heard E., Narita M., Lin A. W., Hearn S. A., et al. (2003). Rb-mediated heterochromatin formation and silencing of E2F target genes during cellular senescence. Cell 113 703–716. 10.1016/s0092-8674(03)00401-x [DOI] [PubMed] [Google Scholar]
  88. Nelson W. J. (2008). Regulation of cell-cell adhesion by the cadherin-catenin complex. Biochem. Soc. Trans. 36 149–155. 10.1042/bst0360149 [DOI] [PMC free article] [PubMed] [Google Scholar]
  89. Nielsen S. J., Schneider R., Bauer U. M., Bannister A. J., Morrison A., O’Carroll D., et al. (2001). Rb targets histone H3 methylation and HP1 to promoters. Nature 412 561–565. 10.1038/35087620 [DOI] [PubMed] [Google Scholar]
  90. Nishiguchi S., Yagi A., Sakai N., Oda H. (2016). Divergence of structural strategies for homophilic E-cadherin binding among bilaterians. J. Cell Sci. 129 3309–3319. [DOI] [PubMed] [Google Scholar]
  91. Oda H., Takeichi M. (2011). Structural and functional diversity of cadherin at the adherens junction. J. Cell Biol. 193 1137–1146. 10.1083/jcb.201008173 [DOI] [PMC free article] [PubMed] [Google Scholar]
  92. Oda H., Tagawa K., Akiyama-Oda Y. (2005). Diversification of epithelial adherens junctions with independent reductive changes in cadherin form: identification of potential molecular synapomorphies among bilaterians. Evol. Dev. 7 376–389. 10.1111/j.1525-142x.2005.05043.x [DOI] [PubMed] [Google Scholar]
  93. Oda H., Tsukita S., Takeichi M. (1998). Dynamic behavior of the cadherin-based cell-cell adhesion system during Drosophila gastrulation. Dev. Biol. 203 435–450. 10.1006/dbio.1998.9047 [DOI] [PubMed] [Google Scholar]
  94. Ogryzko V. V., Schiltz R. L., Russanova V., Howard B. H., Nakatani Y. (1996). The transcriptional coactivators p300 and CBP are histone acetyltransferases. Cell 87 953–959. 10.1016/s0092-8674(00)82001-2 [DOI] [PubMed] [Google Scholar]
  95. Palacios F., Tushir J. S., Fujita Y., D’Souza-Schorey C. (2005). Lysosomal targeting of E-Cadherin: a unique mechanism for the down-regulation of cell-cell adhesion during epithelial to mesenchymal transitions. Mol. Cell Biol. 25 389–402. 10.1128/mcb.25.1.389-402.2005 [DOI] [PMC free article] [PubMed] [Google Scholar]
  96. Paredes J., Figueiredo J., Albergaria A., Oliveira P., Carvalho J., Ribeiro A. S., et al. (2012). Epithelial E- and P-cadherins: role and clinical significance in cancer. Biochim. Biophys. Acta 1826 297–311. [DOI] [PubMed] [Google Scholar]
  97. Peinado H., Ballestar E., Esteller M., Cano A. (2004a). Snail mediates E-cadherin repression by the recruitment of the Sin3A/histone deacetylase 1 (HDAC1)/HDAC2 complex. Mol. Cell Biol. 24 306–319. 10.1128/mcb.24.1.306-319.2004 [DOI] [PMC free article] [PubMed] [Google Scholar]
  98. Peinado H., Portillo F., Cano A. (2004b). Transcriptional regulation of cadherins during development and carcinogenesis. Int. J. Dev. Biol. 48 365–375. 10.1387/ijdb.041794hp [DOI] [PubMed] [Google Scholar]
  99. Perez T. D., Nelson W. J. (2004). Cadherin adhesion: mechanisms and molecular interactions. Handb. Exp. Pharmacol. 2004 3–21. 10.1007/978-3-540-68170-0_1 [DOI] [PMC free article] [PubMed] [Google Scholar]
  100. Piacentini L., Fanti L., Berloco M., Perrini B., Pimpinelli S. (2003). Heterochromatin protein 1 (HP1) is associated with induced gene expression in Drosophila euchromatin. J. Cell Biol. 161 707–714. 10.1083/jcb.200303012 [DOI] [PMC free article] [PubMed] [Google Scholar]
  101. Raj A., van Oudenaarden A. (2008). Stochastic gene expression and its consequences. Cell 135 216–226. 10.1016/j.cell.2008.09.050 [DOI] [PMC free article] [PubMed] [Google Scholar]
  102. Riddle N. C., Minoda A., Kharchenko P. V., Alekseyenko A. A., Schwartz Y. B., Tolstorukov M. Y., et al. (2011). Plasticity in patterns of histone modifications and chromosomal proteins in Drosophila heterochromatin. Genome Res. 21 147–163. 10.1101/gr.110098.110 [DOI] [PMC free article] [PubMed] [Google Scholar]
  103. Robinson B. S., Moberg K. H. (2011). Cell–cell junctions: α-Catenin and E-Cadherin help fence in Yap1. Curr. Biol. 21 R890–R892. [DOI] [PMC free article] [PubMed] [Google Scholar]
  104. Rübsam M., Mertz A. F., Kubo A., Marg S., Jüngst C., Goranci-Buzhala G., et al. (2017). E-cadherin integrates mechanotransduction and EGFR signaling to control junctional tissue polarization and tight junction positioning. Nat. Commun. 8:1250. [DOI] [PMC free article] [PubMed] [Google Scholar]
  105. Russell H., Pranjol M. Z. I. (2018). Transcription factors controlling E-cadherin down-regulation in ovarian cancer. Biosci. Horizons Int. J. Student Res. 11:hzy010. [Google Scholar]
  106. Sarpal R., Yan V., Kazakova L., Sheppard L., Yu J. C., Fernandez-Gonzalez R., et al. (2019). Role of α-Catenin and its mechanosensing properties in regulating Hippo/YAP-dependent tissue growth. PLoS Genet. 15:e1008454. 10.1371/journal.pgen.1008454 [DOI] [PMC free article] [PubMed] [Google Scholar]
  107. Schackmann R. C. J., Tenhagen M., Ven R. A. H., van de, Derksen P. W. B. (2013). p120-catenin in cancer – mechanisms, models and opportunities for intervention. J. Cell Sci. 126 3515–3525. 10.1242/jcs.134411 [DOI] [PubMed] [Google Scholar]
  108. Schmidt C. R., Gi Y. J., Patel T. A., Coffey R. J., Beauchamp R. D., Pearson A. S. (2005). E-cadherin is regulated by the transcriptional repressor SLUG during Ras-mediated transformation of intestinal epithelial cells. Surgery 138 306–312. 10.1016/j.surg.2005.06.007 [DOI] [PubMed] [Google Scholar]
  109. Schock F., Perrimon N. (2002). Molecular mechanisms of epithelial morphogenesis. Annu. Rev. Cell Dev. Biol. 18 463–493. [DOI] [PubMed] [Google Scholar]
  110. Serrano-Gomez S. J., Maziveyi M., Alahari S. K. (2016). Regulation of epithelial-mesenchymal transition through epigenetic and post-translational modifications. Mol. Cancer 15:18. [DOI] [PMC free article] [PubMed] [Google Scholar]
  111. Shang S., Hua F., Hu Z.-W. (2017). The regulation of β-catenin activity and function in cancer: therapeutic opportunities. Oncotarget 8 33972–33989. 10.18632/oncotarget.15687 [DOI] [PMC free article] [PubMed] [Google Scholar]
  112. Shi S., Larson K., Guo D., Lim S. J., Dutta P., Yan S.-J., et al. (2008). Drosophila STAT is required for directly maintaining HP1 localization and heterochromatin stability. Nat. Cell Biol. 10 489–496. 10.1038/ncb1713 [DOI] [PMC free article] [PubMed] [Google Scholar]
  113. Soncin F., Mohamet L., Ritson S., Hawkins K., Bobola N., Zeef L., et al. (2011). E-cadherin acts as a regulator of transcripts associated with a wide range of cellular processes in mouse embryonic stem cells. PLoS One 6:e21463. 10.1371/journal.pone.0021463 [DOI] [PMC free article] [PubMed] [Google Scholar]
  114. Sotillos S., Díaz-Meco M. T., Moscat J., Castelli-Gair Hombría J. (2008). Polarized subcellular localization of Jak/STAT components is required for efficient signaling. Curr. Biol. 18 624–629. 10.1016/j.cub.2008.03.055 [DOI] [PubMed] [Google Scholar]
  115. Stefanatos R. K., Bauer C., Vidal M. (2013). p120 catenin is required for the stress response in Drosophila. PLoS One 8:e83942. 10.1371/journal.pone.0083942 [DOI] [PMC free article] [PubMed] [Google Scholar]
  116. Surapaneni S. K., Bhat Z. R., Tikoo K. (2020). MicroRNA-941 regulates the proliferation of breast cancer cells by altering histone H3 Ser 10 phosphorylation. Sci. Rep. 10:17954. [DOI] [PMC free article] [PubMed] [Google Scholar]
  117. Takeichi M. (1977). Functional correlation between cell adhesive properties and some cell surface proteins. J. Cell Biol. 75 464–474. 10.1083/jcb.75.2.464 [DOI] [PMC free article] [PubMed] [Google Scholar]
  118. van Roy F., Berx G. (2008). The cell-cell adhesion molecule E-cadherin. Cell Mol. Life Sci. 65 3756–3788. 10.1007/s00018-008-8281-1 [DOI] [PMC free article] [PubMed] [Google Scholar]
  119. Vergara D., Simeone P., Franck J., Trerotola M., Giudetti A., Capobianco L., et al. (2016). Translating epithelial mesenchymal transition markers into the clinic: novel insights from proteomics. EuPA Open Proteom 10 31–41. 10.1016/j.euprot.2016.01.003 [DOI] [PMC free article] [PubMed] [Google Scholar]
  120. Vesuna F., van Diest P., Chen J. H., Raman V. (2008). Twist is a transcriptional repressor of E-cadherin gene expression in breast cancer. Biochem. Biophys. Res. Commun. 367 235–241. 10.1016/j.bbrc.2007.11.151 [DOI] [PMC free article] [PubMed] [Google Scholar]
  121. Viré E., Brenner C., Deplus R., Blanchon L., Fraga M., Didelot C., et al. (2006). The Polycomb group protein EZH2 directly controls DNA methylation. Nature 439 871–874. 10.1038/nature04431 [DOI] [PubMed] [Google Scholar]
  122. Wang D., Wang Y., Wu X., Kong X., Li J., Dong C. (2020). RNF20 is critical for snail-mediated E-Cadherin repression in human breast cancer. Front. Oncol. 10:613470. [DOI] [PMC free article] [PubMed] [Google Scholar]
  123. Wang Z., Sandiford S., Wu C., Li S. S.-C. (2009). Numb regulates cell-cell adhesion and polarity in response to tyrosine kinase signalling. EMBO J. 28 2360–2373. 10.1038/emboj.2009.190 [DOI] [PMC free article] [PubMed] [Google Scholar]
  124. Wei S.-Y., Escudero L. M., Yu F., Chang L.-H., Chen L.-Y., Ho Y.-H., et al. (2005). Echinoid is a component of adherens junctions that cooperates with DE-Cadherin to mediate cell adhesion. Dev. Cell 8 493–504. 10.1016/j.devcel.2005.03.015 [DOI] [PubMed] [Google Scholar]
  125. Wendt M. K., Taylor M. A., Schiemann B. J., Schiemann W. P. (2011). Down-regulation of epithelial cadherin is required to initiate metastatic outgrowth of breast cancer. Mol. Biol. Cell 22 2423–2435. 10.1091/mbc.e11-04-0306 [DOI] [PMC free article] [PubMed] [Google Scholar]
  126. Werth M., Walentin K., Aue A., Schönheit J., Wuebken A., Pode-Shakked N., et al. (2010). The transcription factor grainyhead-like 2 regulates the molecular composition of the epithelial apical junctional complex. Development 137 3835–3845. 10.1242/dev.055483 [DOI] [PubMed] [Google Scholar]
  127. Wong T.-S., Gao W., Chan J. Y.-W. (2014). Transcription regulation of E-Cadherin by Zinc Finger E-Box binding homeobox proteins in solid tumors. Biomed. Res. Int. 2014:921564. [DOI] [PMC free article] [PubMed] [Google Scholar]
  128. Xiang X., Deng Z., Zhuang X., Ju S., Mu J., Jiang H., et al. (2012). Grhl2 determines the epithelial phenotype of breast cancers and promotes tumor progression. PLoS One 7:e50781. 10.1371/journal.pone.0050781 [DOI] [PMC free article] [PubMed] [Google Scholar]
  129. Xu N., Emelyanov A. V., Fyodorov D. V., Skoultchi A. I. (2014). Drosophila linker histone H1 coordinates STAT-dependent organization of heterochromatin and suppresses tumorigenesis caused by hyperactive JAK-STAT signaling. Epigenet. Chromatin 7:16. 10.1186/1756-8935-7-16 [DOI] [PMC free article] [PubMed] [Google Scholar]
  130. Xue B., Krishnamurthy K., Allred D. C., Muthuswamy S. K. (2013). Loss of Par3 promotes breast cancer metastasis by compromising cell-cell cohesion. Nat. Cell Biol. 15 189–200. 10.1038/ncb2663 [DOI] [PMC free article] [PubMed] [Google Scholar]
  131. Yan R., Small S., Desplan C., Dearolf C. R., Darnell J. E. (1996). Identification of a stat gene that functions in Drosophila development. Cell 84 421–430. 10.1016/s0092-8674(00)81287-8 [DOI] [PubMed] [Google Scholar]
  132. Yang J., Antin P., Berx G., Blanpain C., Brabletz T., Bronner M., et al. (2020). Guidelines and definitions for research on epithelial–mesenchymal transition. Nat. Rev. Mol. Cell Biol. 21 341–352. [DOI] [PMC free article] [PubMed] [Google Scholar]
  133. Yao L., Wang S., Westholm J. O., Dai Q., Matsuda R., Hosono C., et al. (2017). Genome-wide identification of Grainy head targets in Drosophila reveals regulatory interactions with the POU domain transcription factor Vvl. Development 144 3145–3155. [DOI] [PMC free article] [PubMed] [Google Scholar]
  134. Yoshiura K., Kanai Y., Ochiai A., Shimoyama Y., Sugimura T., Hirohashi S. (1995). Silencing of the E-cadherin invasion-suppressor gene by CpG methylation in human carcinomas. PNAS 92 7416–7419. 10.1073/pnas.92.16.7416 [DOI] [PMC free article] [PubMed] [Google Scholar]
  135. Zetter B. R. (1993). Adhesion molecules in tumor metastasis. Semin. Cancer Biol. 4 219–229. 10.3109/15419069409004440 [DOI] [PubMed] [Google Scholar]
  136. Zhang Q., Dong P., Liu X., Sakuragi N., Guo S.-W. (2017). Enhancer of Zeste homolog 2 (EZH2) induces epithelial-mesenchymal transition in endometriosis. Sci. Rep. 7:6804. [DOI] [PMC free article] [PubMed] [Google Scholar]
  137. Zhang Y., Xu L., Li A., Han X. (2019). The roles of ZEB1 in tumorigenic progression and epigenetic modifications. Biomed. Pharmacother. 110 400–408. 10.1016/j.biopha.2018.11.112 [DOI] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Click here for additional data file. (1.7MB, pdf)

Data Availability Statement

The datasets presented in this study can be found in online repositories. The names of the repository/repositories and accession number(s) can be found below: http://data.modencode.org/, 3187; http://data.modencode.org/, 3188; http://data.modencode.org/, 3391; http://data.modencode.org/, 3392.

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