The Role of WW-PLEKHAs and PDZD11 in the Trafficking of the Copper Pump ATP7A and in Copper Homeostasis

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Thesis

Reference

The Role of WW-PLEKHAs and PDZD11 in the Trafficking of the Copper Pump ATP7A and in Copper Homeostasis

SLUYSMANS, Sophie

Abstract

PLEKHA5 and PLEKHA6 are newly identified proteins from the PLEKHA family which harbor a N-terminal tandem of WW domains that bind to PDZD11, similarly to the junctional protein PLEKHA7. PLEKHA5 and PLEKHA6 have a tissue distribution and subcellular localization distinct from PLEKHA7, since they are present at lateral cell-cell contacts, and additionally along microtubules in the case of PLEKHA5 and at junctions concerning PLEKHA6. Key structural domains, i. e. WW, PH and coiled-coil motifs, direct the distinct subcellular distributions of WW-PLEKHAs (PLEKHA5, PLEKHA6 and PLEKHA7), defining different subcellular pools of PDZD11. By promoting the binding of PDZD11 with the C-terminal PDZ-binding domain of the Menkes copper pump ATP7A, WW-PLEKHAs together with PDZD11 mediate the copper-induced distribution of ATP7A from the trans-Goli network to the cell periphery in polarized kidney epithelial cells, and are required to limit the increase of intracellular copper levels when cells are exposed to elevated copper conditions.

SLUYSMANS, Sophie. The Role of WW-PLEKHAs and PDZD11 in the Trafficking of the Copper Pump ATP7A and in Copper Homeostasis. Thèse de doctorat : Univ. Genève, 2021, no. Sc. Vie 111

DOI : 10.13097/archive-ouverte/unige:155198 URN : urn:nbn:ch:unige-1551988

Available at:

http://archive-ouverte.unige.ch/unige:155198

Disclaimer: layout of this document may differ from the published version.

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UNIVERSITÉ DE GENÈVE FACULTÉ DES SCIENCES Section de Biologie

Département de Biologie Cellulaire Professeure Sandra Citi

The Role Of WW-PLEKHAs And PDZD11 In The Trafficking Of The Copper Pump ATP7A And In Copper Homeostasis

THÈSE

présentée aux Facultés de médecine et des sciences de l’Université de Genève pour obtenir le grade de Docteur ès sciences en sciences de la vie,

mention Biosciences moléculaires

par

Sophie SLUYSMANS

de

Tervuren (Belgique)

Thèse N^o 111

GENÈVE

Centre d’Impression de l’Université de Genève 2021

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Acknowledgments

My PhD time highlighted to me that research is of course about sciences, but also that people are extremely important for doing sciences; many thus deserve my acknowledgments.

I would like to thank Professor Sandra Citi, for giving me the opportunity to carry out my thesis in her lab, where I learned so much. I appreciated her curiosity and enthusiasm about science.

I am grateful Sandra encouraged me to explore other questions that arose from my initial Ph.D.

project, and to collaborate on other projects in the lab, opening my research topic. I further would like to acknowledge her for giving me the opportunity to attend national and international conferences and to actively participate in the writing and reviewing of publications and grants, developing my scientific communication skills.

I would then like to thank the jury members of my Ph.D. thesis defense, Professor Svetlana Lutsenko, Professor Antonis Kourtidis and Professor Jean-Claude Martinou, for their reading of my manuscript and their critical feedback and judgment of my work.

The Department of Cell Biology also deserves my gratitude. I thank my scientific colleagues for their active participation during my presentations at our weekly department meetings, and for the subsequent constructive discussions. I am also grateful for sharing some useful reagents. I am thankful to the “non-scientific” staff too, i. e. the secretaries, Carol and Francesca, and Jacques from the crew of the cleaning facility. Altogether, I thank all the BICEL members for the friendly and positive working atmosphere.

Of course, I will not forget the past and current members of Citi laboratory, who participated to my scientific progress, but who were most importantly amazing colleagues. I am so grateful to have done my thesis in a lab with such a friendly, cohesive and helpful team, formed along my thesis by Diego, Domenica, Jimit, Katya, Arielle, Florian, Isa, Lionel, Marine, Thomas, Vladimir and Annick. I thank Diego, Dome, Jimit, Katya, Lionel and Isa for rapidly welcoming me in the team at my arrival, and for the great time we had also outside the lab (I will not forget our

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barbecues in #Morges). I especially thank Isa for becoming my friend and being my pair in the lab. Working with her in team was so straight as we were “connected”. I am also grateful to Lionel, not only for his precious technical help in the lab, but also for the friendly discussions (about the right way to speak French (Belgian or Swiss…), among others). I smile on the weather forecast advice from Florian and Lionel, and I am wondering how I would have determined the best ski weekend without. Finally, I thank Arielle for being a great desk neighbor and for our essential coffee breaks, Marine for her energy in the lab, and Thomas for being a fantastic lab mate.

I would like to extend my gratitude to the persons who improved my pedagogic skills during my thesis, another essential part of my doctoral training. I gave classes in the course of my Ph.D. and I thank the lecturers for their support in the preparations of these lessons, especially Estella (Sim) Poloni for her kindness. For their patience and interest in my explanations and teaching, I also thank the students in class, and the two students I trained in the lab, Amina Boukhatemi and Flavio Ferreira, who contributed to my research too.

My beloved partner in life, Quentin, deserves all my thankfulness for setting up with me this project of leaving our native country, Belgium, to start our post-master life in Switzerland.

I thank you for having supported me in this journey, and for joining me in our future adventures.

I then express my heartfelt thank you to my parents, les Croûtons Carine and Olivier, for their unconditional love, as well as for their encouragement all along my studies and their support when we decided to settle in Switzerland.

Finally, I would like to thank my family, my brother Antoine, my aunt Annick and my “heart godmother” Véro, for their kindness and the happiness they bring in my life. This thanking has to be extended to my old close friends, Loulou, Jérémy and Mathilde, and to the new friends we have made during these years in our new country, Diego, Sonia, Abhinav, Nastaran…

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Résumé

Le cuivre représente une épée à double tranchant pour la physiologie cellulaire : il est le cofacteur de protéines essentielles impliquées dans des processus tels que la respiration cellulaire, mais il est aussi toxique en raison de son pouvoir oxydant. Par conséquent, une carence en cuivre tout comme un excès sont préjudiciables, comme le montrent les maladies de Menkes et de Wilson. Le cerveau en particulier a un besoin élevé en ce métal, nécessaire pour les nombreuses cuproenzymes qui participent à son développement et fonctionnement.

L’homéostasie du cuivre est d’ailleurs dérégulée dans de nombreuses maladies neurologiques. La teneur intracellulaire en cuivre est dès lors finement contrôlée par des transporteurs qui l’importent ou l’exportent, dont la pompe ATP7A (aussi nommée Menkes).

En conditions basales de teneur en cuivre, la pompe ATP7A se trouve au niveau du trans- Golgi et transporte le cuivre vers son lumen. Elle se déplace vers la périphérie de la cellule pour évacuer le cuivre lorsque de hautes concentrations de ce dernier sont atteintes, afin d’en éviter la toxicité. La sévérité clinique de la maladie de Menkes, due à des mutations de l’ATP7A, est liée au degré de perturbation de ce mouvement intracellulaire de la pompe en réponse à l’excès de cuivre (Skjørringe et al., 2017). Cependant, les mécanismes régulant ce déplacement de l’ATP7A ne sont que partiellement compris. Un motif d’interaction avec les domaines PDZ présent à l’extrémité C-terminale de l’ATP7A a été identifié comme primordial pour sa relocalisation induite par le cuivre vers la membrane basolatérale des cellules épithéliales polarisées (Greenough et al., 2004). Ce motif se lie à PDZD11, une protéine contenant un unique domaine PDZ (Stephenson et al., 2005). Dès lors, PDZD11 pourrait jouer un rôle dans le déplacement de l’ATP7A en réponse à des niveaux élevés de cuivre.

PDZD11 interagit avec le tandem N-terminal de domaines WW de PLEKHA7, une protéine cytoplasmique de la zonula adherens (ZA) des cellules épithéliales polarisées (Guerrera et al., 2016; Pulimeno et al., 2010). PLEKHA7 recrute PDZD11 aux contacts intercellulaires, ce qui permet l’accumulation des protéines transmembranaires nectines aux jonctions adhérentes et

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la concentration du récepteur ADAM10 à la ZA (Guerrera et al., 2016; Shah et al., 2018). Ces mécanismes indiquent que PDZD11, en association avec PLEKHA7, permet la bonne distribution de protéines transmembranaires à la membrane plasmique. Cependant, PLEKHA7 n’est détecté qu’à la ZA et non le long des contacts latéraux entre les cellules épithéliales polarisées. C’est pourtant là que se déplace l’ATP7A dans les cellules polarisées stimulées au cuivre. Cette observation suggère que PDZD11 pourrait interagir avec d’autres partenaires pour réguler la localisation basolatérale de l’ATP7A induite par le cuivre.

Cette thèse identifie et caractérise deux nouveaux partenaires de PDZD11 de la famille PLEKHA, PLEKHA5 et PLEKHA6. Leur tandem N-terminal de domaines WW se lie à la partie N-terminale de PDZD11, à l’instar de PLEKHA7. Puisque ce sont les seules protéines PLEKHA, avec PLEKHA7, à avoir des isoformes présentant un tandem de domaines WW, il est proposé de les nommer WW-PLEKHAs. PLEKHA5 et PLEKHA6 ont une distribution dans les tissus et cellules différente de celle de PLEKHA7. Elles sont en effet présentes aux contacts latéraux entre les cellules, ainsi que le long des microtubules pour PLEKHA5 et à la ZA pour PLEKHA6. Certains domaines structuraux, à savoir WW, PH et les motifs coiled-coil, sont responsables des distributions subcellulaires distinctes des WW-PLEKHAs, qui définissent des pools différents de PDZD11 dans la cellule. Par ailleurs, en promouvant l’interaction de PDZD11 avec le motif d’interaction avec les domaines PDZ présent à l’extrémité C-terminale de l’ATP7A, les WW-PLEKHAs, avec PDZD11, participent au déplacement de la pompe vers la périphérie des cellules épithéliales polarisées du rein lorsque ces dernières sont exposées à des niveaux élevés de cuivre. Les WW-PLEKHAs et PDZD11 régulent aussi l’homéostasie du cuivre en limitant l’augmentation de la forme labile dans les cellules exposées au cuivre.

En définitive, cette thèse montre que les modules WW-PLEKHAs-PDZD11 coopèrent dans le déplacement de l’ATP7A induit par le cuivre vers la périphérie des cellules, afin de réguler l’homéostasie de ce métal. Cela confirme que les protéines cytoplasmiques associées aux contacts intercellulaires sont fondamentales pour la distribution des composants transmembranaires à la membrane plasmique ainsi que pour la régulation de leurs fonctions.

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Abstract

Copper is a double-edged sword for cellular physiology: it is the cofactor of essential proteins involved in fundamental processes including cellular respiration, but it is also toxic due to its oxidative capacity. As a consequence, both its deficiency and excess are deleterious, as illustrated by the Menkes and Wilson diseases. Especially, brain has a high demand for copper, since its development and function rely on several cuproenzymes, and copper homeostasis is dysregulated in several neurological disorders. Intracellular copper is thus tightly controlled by copper influx and efflux transporters, including the Menkes copper pump ATP7A. Under basal copper conditions, ATP7A localizes at the trans-Golgi network and loads copper into the Golgi lumen. It translocates to the cell periphery to extrude copper out of the cell when facing high copper levels, preventing toxicity. The clinical severity of the Menkes disease, which is caused by mutations of ATP7A, is linked to the degree of disruption of this copper-induced intracellular trafficking (Skjørringe et al., 2017). Nevertheless, the mechanisms of ATP7A trafficking are only partially understood. A PDZ-binding domain present at the C-terminus of ATP7A is essential for its copper-dependent relocalization to basolateral plasma membrane in polarized epithelial cells (Greenough et al., 2004), and binds to the single PDZ domain-containing protein PDZD11 (Stephenson et al., 2005). Thus, PDZD11 could play a role in ATP7A translocation in response to elevated copper.

PDZD11 interacts with the N-terminal tandem of WW domains of the Pleckstrin homology- domain containing family A member 7 (PLEKHA7), which is a cytoplasmic component of the zonula adherens (ZA) from polarized epithelial cells (Guerrera et al., 2016; Pulimeno et al., 2010). PLEKHA7 recruits PDZD11 to cell-cell contacts, which allows the accumulation of the transmembrane proteins nectins at adherens junctions and the clustering of ADAM10 receptor at ZA (Guerrera et al., 2016; Shah et al., 2018). These uncovered mechanisms indicate that PDZD11, in association with the scaffolding protein PLEKHA7, serves as an adaptor to correctly target transmembrane components to the plasma membrane. However, PLEKHA7 is

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restricted to the ZA and is not detected along lateral cell-cell contacts of polarized epithelial cells, where ATP7A translocates in copper-stimulated polarized cells. This observation suggests that PDZD11 may interact with additional partners to mediate the copper-dependent basolateral targeting of ATP7A.

This thesis identifies and characterizes two new PDZD11 binding-partners from the PLEKHA family, PLEKHA5 and PLEKHA6. Their N-terminal tandem of WW domains binds to the N-terminal sequence of PDZD11, similarly to PLEKHA7. Since, with PLEKHA7, PLEKHA5 and PLEKHA6 are the only PLEKHA proteins with isoforms containing N-terminal WW domains, it is proposed to name them WW-PLEKHAs. PLEKHA5 and PLEKHA6 have a tissue distribution and subcellular localization distinct from PLEKHA7, since they are present at lateral cell-cell contacts, and additionally along microtubules in the case of PLEKHA5 and at the ZA concerning PLEKHA6. Key structural domains, i. e. WW, PH and coiled-coil motifs, direct the distinct subcellular distributions of WW-PLEKHAs, defining different subcellular pools of PDZD11. In addition, by promoting the binding of PDZD11 with the C-terminal PDZ-binding domain of ATP7A, WW-PLEKHAs in association with PDZD11 mediate the copper-dependent translocation of ATP7A to cell periphery in polarized kidney epithelial cells. Finally, they regulate copper homeostasis by limiting the increase of intracellular labile copper in cells exposed to high copper concentrations. This thesis thus proposes that WW-PLEKHAs- PDZD11 modules cooperatively regulate the copper-induced translocation of ATP7A to cell periphery to modulate copper homeostasis. This evidence confirms that cell-cell contacts- associated cytoplasmic proteins are fundamental to properly localize transmembrane components and regulate their function.

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List of Figures

Figure 1. Schematic representation of the basic architecture of cellular junctions.

Figure 2. Cellular junctions of vertebrate epithelial cells.

Figure 3. Structure and molecular composition of TJs.

Figure 4. Domain organization and interactors of ZO proteins.

Figure 5. AJs and their molecular organization in polarized epithelial cells.

Figure 6. F-actin linkage and mechanotransduction at cadherin/catenins complex of epithelial cells.

Figure 7. Schematic representation of the molecular structure and interactors of human PLEKHA7.

Figure 8. Schematic representation of the molecular structure and interactors of human PDZD11.

Figure 9. PLEKHA7/PDZD11 complex at AJs and its functions.

Figure 10. Schematic representation of molecular structure and subcellular localization of PLEKHA proteins.

Figure 11. Mammalian copper homeostasis.

Figure 12. Scheme illustrating the regulation of dopamine-ß-monooxygenase by ATP7A and ATP7B in neurons of the locus coeruleus.

Figure 13. Topographic structure of the copper ATPases ATP7A/B.

Figure 14. ATP7A trafficking signals and pathways.

Figure 15. Phosphorylation sites in human ATP7A.

Figure 16. Major trafficking steps of transmembrane proteins towards the basolateral plasma membrane in polarized epithelial cells.

Figure 17. Subcellular localization of WW-PLEKHAs in polarized epithelial cells.

Figure 18. PDZD11 subcellular pools defined by WW-PLEKHAs in polarized epithelial cells.

Figure 19. Proposed model for the role of WW-PLEKHAs-PDZD11 modules in the copper- induced translocation of ATP7A to basolateral plasma membrane in polarized epithelial cells.

Figure 20. Structural determinants for the subcellular distribution of WW-PLEKHAs.

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List of Tables

Table 1. Summary of PLEKHA7 interactors, cellular functions and implications in physiology and pathologies.

Table 2. Summary of PDZD11 interactors and cellular functions.

Table 3. Implications of junctional proteins in the trafficking of transmembrane proteins.

Table 4. Implications of PLEKHA proteins in protein trafficking.

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Abbreviations

AJ Adherens junction

AJC Apical junctional complex

ALS Amyotrophic lateral sclerosis

aPKC Atypical protein kinase C

APP Amyloid precursor protein

BCR B-cell antigen receptor

CAMSAP3 Calmodulin-regulated spectrin-associated protein 3

CAR Cocksackie and Adenovirus Receptor

CCC COMMD/CCDC22/CCDC93

CCS Copper chaperone for Cu/Zn superoxide dismutase

CCO Cytochrome c oxidase

CGN Cingulin

CGNL1 Paracingulin

CHO Chinese hamster ovary

CNS Central nervous system

COMMD1 Copper metabolism MURR1 domain–containing 1

CTR1 Copper transporter 1

Cu Copper

CUL3 E3 ubiquitin ligase Cullin-3

DMT1 Divalent Metal Transporter 1

DVL Dishevelled

eNOS Endothelial nitric oxide synthase

ER Endoplasmic reticulum

FAPP Phosphatidylinositol-four-phosphate adaptor protein FRAP Fluorescence recovery after photobleaching

GAP GTPase activating protein

GEF Guanine nucleotide exchange factor

GLTP Glycolipid-transfer-protein

GWAS Genome-wide association studies

hSMVT Human sodium-dependent multivitamin transporter ICP-MS Inductively coupled plasma mass spectrometry

IF Immunofluorescence

Ig Immunoglobulin

IP Immunoprecipitation

JAM Junctional Adhesion Molecule

JIP c-jun NH2-terminal kinase (JNK)-interacting protein

KD Knockdown

KI knock-in

KLHL12 Kelch-like protein 12

KO Knockout

LC Locus coeruleus

LCH Langerhans cell histiocytosis

MAGI Membrane-associated guanylate kinase inverted

MAGUK Membrane-associated guanylate kinase

MAPK Mitogen-activated protein kinase

MBS Metal binding sites

MT Microtubule

PACG Primary angle closure glaucoma

PAR3 Partitioning defective 3

PAR6 Partitioning defective 6

PDZ PsD-95/Disc-large/ZO-1

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PDZD11 PDZ domain-containing protein 11

PEPP Phosphatidylinositol-three-phosphate-binding PH-domain protein

PH Pleckstrin homology

PI3K Phosphatidylinositol 3-kinase

PKA Protein kinase A

PLEKHA Pleckstrin homology-domain containing family A

PMCA Plasma membrane calcium ATPase

PrP Prion protein

PtdIns Phosphatidylinositol

PTPL1 Protein-tyrosine-phosphatase-like protein-1

Rab Ras-related proteins in brain

RISC RNA-induced silencing complex

ROS Reactive oxygen species

SJ Septate junction

SOD1 Superoxide dismutase 1

TAPP Tandem PH-domain-containing protein

TEM Transmission electron microscopy

TF Transcription factor

TGN Trans-Golgi network

TJ Tight junction

Tspan33 Tetraspanin-33

tTJ Tricellular tight junction

WASH Wiskott–Aldrich syndrome protein and SCAR homologue

WT Wild type

XIAP X-linked inhibitor of apoptosis

ZA Zonula adherens

ZIM ZO-1-interaction motif

ZO Zonula occludens

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Introduction

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1. Cellular junctions

The emergence of multicellular organisms from ancestral unicellular entities has required the evolution of structures present at the surface of cells to provide contact and adhesion with neighboring cells and with the extracellular matrix. One refers to these structures as “cellular junctions” (Figure 1). They have been a prerequisite for the establishment of tissues and organs, which allow multicellular beings to achieve their complexity by developing systems specialized in specific biological functions. Cellular junctions are multi-molecular complexes composed of core transmembrane proteins and of cytoplasmic proteins present just below the plasma membrane, at the cytoplasmic plaque, ensuring a connection between the transmembrane proteins and the cytoskeleton (Hartsock and Nelson, 2008) (Figure 1). These complexes connect either two adjacent cells, forming cell-cell junctions, or the cell to its surrounding matrix, establishing cell-matrix junctions (Figure 1).

Figure 1. Schematic representation of the basic architecture of cellular junctions.

The complexes formed by transmembrane proteins and cytoplasmic plaque link either two neighboring cells (cell-cell junctions), or a cell to the extracellular matrix (cell-matrix junctions).

In metazoans, most of the cell types harbor cellular junctions, but epithelial cells exhibit junctional structures that are highly specialized and crucial for their biological functions.

Epithelia cover the external surface of the body, line inner cavities and the lumen of blood and lymphatic vessels, and form glandular tissues. Epithelial cells thus constitute a protective barrier against the surrounding environment, but have also sensorial functions to sense and

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adapt to the situation of the organism, for instance in the sensory organs of the skin or ears.

In addition, epithelia separate different compartments in the body and regulate the exchange of organic and inorganic components between them. For example, cells covering the intestines are crucial for the absorption of nutrients following digestion, whereas epithelial cells present in the kidney are fundamental for the maintenance of salt homeostasis and the re-absorption and secretion of solutes. To exert all these roles, cell-cell junctions of epithelial cells are of fundamental importance.

2. Cell-cell junctions: structure and canonical functions

Cell-cell junctions exert altogether various functions that are fundamental to fulfill the roles of epithelia in the organism (Abedin and King, 2010): (i) they mechanically connect adjacent cells in order to form a tissue; (ii) they contribute to the establishment and maintenance of the apicobasal cell polarity, i. e. the segregation between apical and basolateral membranes, resulting in an asymmetric distribution of plasma membrane components and in the polarized separation of the extracellular compartments (Anderson and Van Itallie, 2009); (iii) they seal the plasma membrane of adjacent cells to establish a semipermeable paracellular barrier between the apical and basal compartments, preventing the passage of undesired entities across the tissues (Marchiando et al., 2010) and mediating physiological gradient between the extracellular compartments (Anderson and Van Itallie, 2009); (iv) they allow the direct exchange of ions and small molecules between neighboring cells (Skerrett and Williams, 2017).

The organization of cell-cell junctions has increased in complexity along with the evolution from invertebrate to vertebrate organisms. In vertebrate epithelial cells (Figure 2), tight junctions (TJs) are localized at the most apical region of cell-cell contacts, where they form a circumferential belt-like structure of close contact between adjacent cells, known as the zonula occludens (ZO) (Farquhar and Palade, 1963; Staehelin, 1973). Vertebrate TJs are the paracellular barrier controlling the passage of entities between cells and act as a fence to

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prevent the mixing of apical and basolateral plasma membrane components, defining the apicobasal cell polarity (Zihni et al., 2016). Immediately below TJs, adherens junctions (AJs) also form a belt, the so-called zonula adherens (ZA), mediating adhesion between cells (Takeichi, 2014) (Figure 2). Both TJs and the ZA are connected to the underlying actomyosin cytoskeleton and to microtubules (MTs) (Sluysmans et al., 2017). Since TJs and the ZA are associated at the apical end of the lateral membrane, they are referred as the apical junctional complex (AJC) (Farquhar and Palade, 1963) (Figure 2). Below the ZA, AJ-associated proteins are distributed along lateral contacts but do not form organized junctions. TJs and ZA used to be thought as discrete complexes, but numerous lines of evidence support that they are functionally interconnected, with resident proteins of one or the other interacting together (Muller et al., 2005; Itoh et al., 1997; Yamamoto et al., 1997). In addition, their biogenesis is tightly interdependent: after an initial contact between two neighboring cells driven by AJs, the adherens spot-like contacts expand and support the formation of TJs, ultimately leading to cell polarization and junction maturation separating TJs and ZA (Rusu and Georgiou, 2020). In endothelial cells, TJs and AJs are molecularly intermixed and cannot be distinguished, even in mature cell-cell contacts, and a ZA distinct from lateral contacts is not described (Vorbrodt and Dobrogowska, 2003; Dejana, 2004; Vasileva et al., 2017). The two last types of cell-cell junctions, distributed laterally, are desmosomes that further anchor adjacent cells together, providing resistance to mechanical stress (Garrod and Chidgey, 2008), and gap junctions which form intercellular channels for the passage of ions and small molecules (Skerrett and Williams, 2017) (Figure 2). Finally, anchoring cells to the extracellular matrix, hemidesmosomes and focal adhesions are present at the basal side of the cells (Figure 2).

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Figure 2. Cellular junctions of vertebrate epithelial cells.

The different types of cell-cell (tight junctions, adherens junctions, desmosomes and gap junctions) and cell-matrix (hemidesmosomes and focal adhesions) junctions present in vertebrate epithelial cells.

In invertebrate epithelial cells, the adhesion and barrier functions are carried out respectively by AJs and septate junctions (SJs) which are the functional counterparts of vertebrate TJs and localize basally to AJs at the lateral cell-cell contacts (Rusu and Georgiou, 2020). Unlike vertebrate TJs, SJs do not exert a fence function: the subapical complex/marginal zone, which is apical to the AJs, defines cell polarity in invertebrate epithelia (Citi et al., 2014). Gap junctions are present in invertebrate epithelial cells too, with minor differences with respect to those of vertebrates (Skerrett and Williams, 2017).

The organization and molecular composition of cell-cell junctions are finely tuned to the physiological requirements of each tissue. For instance, endothelial cells of the blood-brain barrier possess TJs of drastically limited permeability in order to protect the brain from the entry of pathogens and undesired components, whereas endothelial cells of non-neural tissues are leakier (Weiss et al., 2009). This characteristic is explained molecularly by a different expression level of the TJ protein occludin (Hirase et al., 1997). Similarly, cultured keratinocytes exhibit discontinuous junctions at confluence but continuous linear junctions when overconfluent and multilayered (Vasileva et al., 2017), consistent with the observation of continuous zonular junctions only in the top layer of stratified epithelia such as the skin (Niessen, 2007). This indicates that the composition and organization of cell-cell contacts

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adapt to various physiological conditions that cells can encounter within a tissue, including mechanical tension. Likewise, when epithelial cells change their position within a tissue, such as during development, they become motile and cell-cell contacts are adapted by changing their composition and organization (Wheelock et al., 2008). Dysregulation of these processes are associated with pathologies such as cancers (Wheelock et al., 2008).

In summary, cell-cell junctions are fundamental structures primarily responsible of cell-cell adhesion, paracellular permeability control and apicobasal cell polarity, but the AJC is also involved in the regulation of cell morphology and migration via its association with the actin and MT cytoskeletons (Bryant and Mostov, 2008; Hartsock and Nelson, 2008). In addition, the AJC appears more and more as a signaling hub regulating pathways controlling gene expression, cell proliferation, differentiation and signaling (Guillemot et al., 2008b; Citi et al., 2014; Meng and Takeichi, 2009; Sluysmans et al., 2017).

3. Tight junctions

The morphological hallmarks of TJs were first observed in 1963 by Farquhar and Palade (Farquhar and Palade, 1963) using transmission electron microscopy (TEM) on ultrathin sections of epithelia (Figure 3A). TJs were found at the most apical region of the junctional complex and appeared as close apposing sites between the plasma membrane of neighboring cells, looking as if the outer leaflets of membranes fused with one another and seal the gap of extracellular space. Freeze fracture electron microscopy (Figure 3A) further revealed TJs as a network of interconnected strands presumably composed of transmembrane components that interact with the fibrils from adjacent cell, linking the cells together (Claude and Goodenough, 1973; Staehelin et al., 1969; Staehelin, 1973). A cytosolic electron-dense area associated to TJs was also observed by TEM (Farquhar and Palade, 1963) (Figure 3A). It corresponds to the cytoplasmic plaque, including scaffolding proteins that link transmembrane TJ proteins to the cytoskeleton, as well as adaptor and signaling proteins.

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In the following decades, the identification and characterization of TJ components (Figure 3B) allowed to understand how TJs exert their canonical functions, i. e. the regulation of paracellular permeability and the maintenance of apicobasal polarity, and led to the discovery of additional roles in gene expression control, cell proliferation, differentiation and migration, and mechanotransduction (Balda and Matter, 2008; Guillemot et al., 2008b; Schneeberger and Lynch, 2004; Spadaro et al., 2017; Sluysmans et al., 2017).

Figure 3. Structure and molecular composition of TJs.

(A) Morphology of TJs observed by TEM of ultrathin sections and freeze fracture replicas of mouse intestinal epithelial cells. Abbreviations: TJ: tight junction; AJ, adherens junction; Ds, desmosome.

Adapted from Otani and Furuse (2020). (B) Overview of the molecular organization of TJs. Claudins (red), occludin (green), and JAM/ESAM/CARs (blue) are the major integral membrane proteins of TJs.

Claudins form TJ strands. Adaptor proteins and cytoskeletal linkers (purple and pink ovals), as well as

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polarity proteins (blue octagons), present at the cytoplasmic plaque of TJs include: the ZO proteins ZO-1, ZO-2 and ZO-3; cingulin (CGN) and paracingulin (CGNL1); membrane-associated guanylate kinase inverted (MAGI); and partitioning defective 3 (PAR3) and 6 (PAR6). Signaling components associated with TJs (yellow rectangles) include atypical protein kinase C (aPKC); the small Rho GTPases Cdc42, Rac1 and RhoA; and their regulators, guanine nucleotide exchange factors (RhoGEFS; orange rectangle). Additionally, the interaction between ZO-1 and the transcription factor ZONAB/DbpA is shown (green octagon).

3.1. Transmembrane proteins of tight junctions

The first transmembrane protein of TJs to be identified was occludin, following the generation of monoclonal antibodies against a liver junctional fraction, by the Tsukita laboratory in 1993 (Furuse et al., 1993). Occludin belongs to the TJ-associated MARVEL domain-containing protein family, which also includes tricellulin. Tricellulin is present at tricellular TJs (tTJs) (Ikenouchi et al., 2005), i. e. specialized structures occurring at the junction between three neighboring cells, but can localize along bicellular TJs when occludin is depleted (Ikenouchi et al., 2008). Spanning the plasma membrane four times, occludin harbors a cytoplasmic C-terminus that is rich in phosphorylatable residues (serine, threonine and tyrosine), and its phosphorylation has been linked to its accumulation at junctions and to the dynamic regulation of TJs (Wong, 1997; Sakakibara et al., 1997; Andreeva et al., 2001; Hirase et al., 2001). The C-terminal tail of occludin interacts directly with the actin cytoskeleton and also indirectly via adapters such as ZO proteins and cingulin (Furuse et al., 1994; Fanning et al., 1998; Itoh et al., 1999b; Haskins et al., 1998; Cordenonsi et al., 1999b).

It was recently shown that an actin-dependent stretching of ZO-1 allows the accumulation of occludin at TJs (Spadaro et al., 2017), possibly related to its partitioning into ZO-1 phase- separated condensates (Beutel et al., 2019). The role of occludin at TJs remains nevertheless elusive. Early studies highlighted its importance in TJ structure and function: occludin exogenously expressed in fibroblasts, which normally lack TJs, localized at cell-cell contacts, conferred them adhesiveness (Van Itallie and Anderson, 1997), and participated in the formation of TJ-like strands (Furuse et al., 1998), whereas the treatment of epithelial cells with synthetic occludin peptides disrupted TJ assembly and barrier function (Lacaz-Vieira et al., 1999; Chung et al., 2001; Nusrat et al., 2005). However, occludin-knockdown (KD) or - knockout (KO) cells were able to form morphologically normal TJ strands and displayed

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unaffected paracellular permeability (Yu et al., 2005; Saitou et al., 1998). In addition, TJs in occludin-KO mice were not disrupted and the intestinal epithelium showed a functional barrier despite the observation of histological abnormalities in the mutant animals (Saitou et al., 2000;

Schulzke et al., 2005). Therefore, although a substitutional redundancy exerted by tricellulin could obscure the roles of occludin (Furuse, 2010; Ikenouchi et al., 2005; Ikenouchi et al., 2008), additional transmembrane components of TJs must be involved in the formation of TJ strands and their barrier function.

Subsequent work from the Tsukita laboratory identified claudins as the main components of TJ strands (Furuse et al., 1998). Claudins are a protein family which encompasses 26 members in human and 27 in mouse, expressed tissue-specifically (Gunzel and Yu, 2013;

Anderson and Van Itallie, 2009; Furuse, 2010). They are the core structural components of TJ fibrils, and can generate TJ strands by themselves when overexpressed in fibroblasts lacking endogenous TJs (Furuse et al., 1998; Kubota et al., 1999). They span the membrane four times, and their C-terminal cytoplasmic tail can be modified by phosphorylation and palmitoylation, which regulate their accumulation at junctions (Van Itallie et al., 2005; D'Souza et al., 2005; Aono and Hirai, 2008; Ishizaki et al., 2003; Ikari et al., 2006). Furthermore, most of the claudins contain at their C-terminus a PsD-95/Disc-large/ZO-1 (PDZ)-binding motif, which allows them to bind to PDZ domain-containing proteins of the TJ cytoplasmic plaque such as ZO proteins (Itoh et al., 1999a). In addition to bridging claudins to the actin cytoskeleton, this interaction with the PDZ domains of ZO-1 and ZO-2 plays a fundamental role in the assembly of TJ fibrils (Umeda et al., 2006). The extracellular loops and transmembrane spans of claudins mediate their homo- and heterophilic interactions, both in cis, i. e. in the same plasma membrane, and in trans, i. e. with claudins from the neighboring cell (Furuse et al., 1999; Krause et al., 2008; Blasig et al., 2006; Rossa et al., 2014; Piontek et al., 2008; Zhao et al., 2018). These assemblies are critical for TJ formation and permeability barrier function. Indeed, by forming paracellular pores with their extracellular loops and controlling the passage of ions and small molecules, claudins are the major determinant of

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paracellular permeability (Gunzel and Yu, 2013). Since TJ strands are a “mosaic” of multiple claudin types, it allows the modulation of the permeability function (water, size and charge selectivity), depending on the specialization of the tissue, by combining different claudin types (Gunzel and Yu, 2013; Furuse et al., 1999). Physiologically, two pathways of paracellular permeability have been observed, one that forms charge-selective small-pores (estimated diameter of ~4Å), and another that is size-selective and allows the permeation of molecules up to ~60Å (Otani and Furuse, 2020; Zihni et al., 2016; Shen et al., 2011; Watson et al., 2005).

It was reported that claudin-5-KO mouse endothelial cells and claudin quintuple-KO MDCK cells have an altered permeability barrier to ions and small molecules but retain their barrier to macromolecules intact (Nitta et al., 2003; Otani et al., 2019), suggesting the involvement of additional proteins in the function of gate to larger molecules. The role of claudins in the fence function of TJs is also not clear: epithelial polarity and lipid apicobasal distribution were not disorganized in claudin quintuple-KO MDCK cells (Otani et al., 2019), suggesting that TJ strands per se are not essential for membrane fence formation.

In addition to tetraspan proteins, other transmembrane components of TJs include single-span transmembrane immunoglobin-like adhesion molecules, such as JAMs (Junctional Adhesion Molecules) (Martin-Padura et al., 1998; Luissint et al., 2014), CAR (Cocksackie and Adenovirus Receptor) (Cohen et al., 2001b), ESAM (Endothelial cell Selective Adhesion Molecule) (Nasdala et al., 2002; Hirata et al., 2001) and the tTJs-specific angulins (Masuda et al., 2011). ESAM is specifically present in endothelial cells, whereas CAR is detected in epithelial cells (Bazzoni, 2003). JAM-A and -C have a wide distribution, whereas JAM-B is restricted to the endothelial cells of specific blood vessels (Bazzoni, 2003). JAMs bind in cis and trans in an homo- and heterophilic manner, but not in all combinations (Bazzoni, 2003). They do not constitute TJ strands, but are involved in the membrane apposition observed at TJs since claudin quintuple-KO MDCK cells lack TJ strands but keep the close juxtaposition of adjacent membranes, and a widening of the intercellular space is observed when JAM-A is additionally removed (Otani et al., 2019). JAM-A appears to stabilize the

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paracellular barrier (Bazzoni, 2003), since the intestinal epithelium of JAM-A-KO mice is easily disrupted when facing inflammation (Laukoetter et al., 2007; Vetrano et al., 2008). In support of this idea, JAM-A-KD cultured epithelial monolayers are leakier to large molecules (Liu et al., 2000). Furthermore, claudin quintuple-KO MDCK monolayers have a disrupted permeability to small molecules and ions but keep their barrier role to macromolecules until JAM-A is also removed, indicating the critical role of JAMs in macromolecule paracellular permeability control in coordination with claudins (Ebnet et al., 2000; Laukoetter et al., 2007; Otani et al., 2019).

The cytoplasmic C-terminal tail of JAMs interact with several proteins of the cytoplasmic plaque of junctions, including ZO-1 via its PDZ domain (Ebnet et al., 2000; Bazzoni et al., 2000) and CGN (Bazzoni et al., 2000). These bindings connect JAMs to the cytoskeletons and support its role in TJ assembly (Bazzoni et al., 2000; Bazzoni and Dejana, 2001; Martinez-Estrada et al., 2001; Bazzoni, 2003). JAMs also bind and recruit polarity complex proteins (Par3, Par6 and aPKC) and have thus been implicated in the fence function of TJs and the establishment of epithelial polarity (Ebnet et al., 2001; Itoh et al., 2001; Rehder et al., 2006; Tuncay et al., 2015; Otani et al., 2019; Otani and Furuse, 2020).

3.2. Tight junction cytoplasmic proteins

Transmembrane components of TJs are directly linked to proteins underlying the membrane and forming the TJ cytoplasmic plaque (Figure 3). These proteins serve as interface between transmembrane components and the actin and MT cytoskeletons, and participate in TJ canonical fence and gate functions (Balda and Matter, 2008). They also confer additional roles by transmitting signals from the junction in order to regulate several cellular processes including cell migration, proliferation and differentiation, gene expression and cytoskeletal organization (Balda and Matter, 2008; Guillemot et al., 2008b; Schneeberger and Lynch, 2004). The TJ plaque is thus composed of a complex network of scaffolding, polarity, adaptor, cytoskeletal and signaling proteins (Guillemot et al., 2008b; Balda and Matter, 2008).

ZO proteins (Figure 4) comprise three structurally related proteins, ZO-1, -2 and -3, belong to the membrane-associated guanylate kinase (MAGUK) family and are at the core of the TJ

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cytoplasmic plaque. ZO-1 was the first cytoplasmic TJ-associated component to be identified, following the generation of monoclonal antibodies against a TJ-enriched fraction from mouse liver (Stevenson et al., 1986). ZO-2 and ZO-3 were subsequently identified as proteins that co-immunoprecipitated with ZO-1 in cultured epithelial cell lysates (Gumbiner et al., 1991;

Haskins et al., 1998). ZO-1 and ZO-2 are essential, since mice KO for one or the other die at early embryonic stage (Katsuno et al., 2008; Xu et al., 2008), whereas ZO-3-KO mice are viable and do not show any TJ disruption phenotype (Xu et al., 2008; Adachi et al., 2006). ZO proteins contain three PDZ motifs and several additional structural domains, which mediate multiple interactions and allow them to be the pillars of TJs (Figure 4).

Figure 4. Domain organization and interactors of ZO proteins.

Domains are color-coded. Interacting partners are indicated with a line that delineates the approximate localization of the binding sites. SH: Src-homology; GUK: guanylate kinase. Adapted from Rouaud et al. (2020a).

ZO proteins associate with themselves by homo- and heterodimerization. The first PDZ domain of ZO-1 mediates its homodimerization, whereas the second is responsible for heterodimerization through binding to the second PDZ domain of either ZO-2 or ZO-3 (Utepbergenov et al., 2006). In addition, PDZ domains of ZO proteins directly bind the C-terminal cytosolic tail of the main TJ transmembrane components (claudins, occludin and JAMs) to promote their junctional recruitment and stabilization, and are indispensable for TJ formation, cell polarity and paracellular barrier functions (Itoh et al., 1999a; Furuse et al., 1994;

Fanning et al., 1998; Haskins et al., 1998; Ebnet et al., 2000; Umeda et al., 2006; Otani and

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Furuse, 2020). It was recently shown that ZO proteins can phase separate and recruit into their condensates several TJ components, including occludin and claudins, driving the clustering of TJ-associated proteins required for TJ assembly (Beutel et al., 2019; Schwayer et al., 2019;

Otani and Furuse, 2020; Rouaud et al., 2020a). ZO proteins also ensure the crosstalk between TJs and the actomyosin cytoskeleton, directly by binding with actin and cortactin and indirectly via their interaction with other actin-binding proteins and regulators of Rho GTPases (Itoh et al., 1997; Fanning et al., 1998; Katsube et al., 1998; Rouaud et al., 2020a; Guillemot et al., 2008b; Citi et al., 2014; Itoh et al., 2012; Ikenouchi et al., 2007; Odenwald et al., 2018). Finally, ZO-1 and ZO-2 regulate gene expression by sequestering at junctions the transcription factor DbpA/ZONAB, mediating TJ mechanotransduction (Spadaro et al., 2014; Spadaro et al., 2017). Indeed, this ability to bind to DbpA, and also occludin, depends on the conformation of ZO-1 and ZO-2, related to their heterodimerization and the contractility of the actin cytoskeleton or the substrate stiffness (Spadaro et al., 2014; Spadaro et al., 2017).

A canonical function of TJs is to mediate the establishment and maintenance of apicobasal cell polarity. The coordinated activity of the apical polarity complexes Par3/Par6/aPKC and Crumbs/PALS1/PATJ/MUPP1, which are localized at TJs, is fundamental for this process (Rodriguez-Boulan and Macara, 2014; Assemat et al., 2008; Suzuki and Ohno, 2006; Riga et al., 2020). Partitioning defective 3 (Par3) interacts with the transmembrane proteins JAMs through its first (out of three) PDZ domain (Itoh et al., 2001). It subsequently recruits the other members of the complex, i. e. partitioning defective 6 (Par6) and the catalytical partner atypical protein kinase C (aPKC) activated by the Rho GTPases Cdc42 or Rac1 (Lin et al., 2000;

Peterson et al., 2004; Johansson et al., 2000). The second polarity complex is composed of the transmembrane component Crumbs bound to the PDZ protein PALS1 that is in turn linked to the multi-PDZ proteins PATJ and MUPP1 (Tepass et al., 1990; Roh et al., 2002b; Roh et al., 2002a; Assemat et al., 2013; Bulgakova and Knust, 2009). PATJ also directly interacts with the transmembrane TJ components claudins, and MUPP1 with claudins, JAMs and CAR, serving thus as scaffolds for both cytoplasmic and transmembrane proteins (Roh et al., 2002a;

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Hamazaki et al., 2002; Jeansonne et al., 2003; Coyne et al., 2004; Rouaud et al., 2020a). All these interactions bring together several proteins and their downstream effectors at the specific localization of TJs and thus drive junction assembly and polarity formation (Assemat et al., 2008; Suzuki and Ohno, 2006; Riga et al., 2020).

Two related proteins, cingulin (CGN) and paracingulin (CGNL1, also known as JACOP), are crucial scaffolds for the crosstalk between TJs and actomyosin and MT cytoskeletons (Rouaud et al., 2020a). CGN was discovered as a protein that co-purified with non-muscle myosin II from chicken intestinal epithelial cells (Citi et al., 1988; Citi et al., 1989), and CGNL1 was detected in a junction-enriched fraction isolated from mouse liver (Ohnishi et al., 2004). Both form parallel homodimers and contain an N-terminal globular head domain, a long α-helical rod domain and a small C-terminal globular tail (Cordenonsi et al., 1999a; Ohnishi et al., 2004;

Citi et al., 2000). They are localized in a distal layer of the TJs, further away from the membrane than ZO proteins (Citi et al., 1988; Citi et al., 1989; Stevenson et al., 1989; Van Itallie and Anderson, 2014; Rouaud et al., 2020a). Depending on the tissue, CGNL1 is also detected at the ZA in addition to TJs (Ohnishi et al., 2004). The junctional recruitment of CGN depends on its interaction with ZO-1 through its N-terminal ZO-1-interaction motif (ZIM) (D'Atri et al., 2002), whereas CGNL1 is brought to junctions either by ZO-1 via the same conserved ZIM domain, or by PLEKHA7, or both, depending on the cell type (Tornavaca et al., 2015; Ohnishi et al., 2004; Pulimeno et al., 2011). In addition to its interaction with ZO-1, CGN binds actin and myosin, and co-pellets and bundles actin filaments in vitro, thus directly tethering the actomyosin cytoskeleton to ZO proteins and TJs (D'Atri and Citi, 2001; Cordenonsi et al., 1999a; Yano et al., 2018; Rouaud et al., 2020a). Moreover, CGN and CGNL1 are adaptors of guanine nucleotide exchange factors (GEFs) and GTPase activating proteins (GAPs), thus fine-tuning the activity of Rho small GTPases and therefore regulating indirectly cytoskeletal organization (Citi et al., 2014). CGN and CNGL1 are also linked to MTs, since CGN binds to MTs in a phosphorylation-dependent manner via its head domain (Yano et al., 2013; Mangan et al., 2016; Yano et al., 2018) and CGNL1 co-pellets with MTs in vitro and is less recruited to

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junctions when MTs are depolymerized (Vasileva and Citi, 2018; Paschoud et al., 2011).

Depletion of either CGN or CGNL1 in cellular models does not result in striking changes of the TJ morphology and functions, neither in cytoskeleton architecture (Guillemot et al., 2008a;

Guillemot and Citi, 2006; Guillemot et al., 2004; Guillemot et al., 2012). CGN-KO mice are viable but have an impaired mucosal repair (Guillemot et al., 2012), and CGN modulates the endothelial barrier both in human cultured cells and in mouse brains (Schossleitner et al., 2016). Angiogenesis in CGNL1-KO endothelial cells is disturbed (Chrifi et al., 2017), and CGNL1-KO and CGN-KO cells exhibit an altered dynamic of junction formation and gene expression due to the impacts on RhoA and Rac1 activation (Aijaz et al., 2005; Guillemot et al., 2008a; Guillemot and Citi, 2006; Guillemot et al., 2004; Guillemot et al., 2013).

Therefore, proteins of the TJ cytoplasmic plaque exert crucial roles in junction assembly and stabilization, and represent a signaling platform regulating several cellular processes such as gene expression, mechanotransduction, cytoskeletal organization, cell migration, proliferation and differentiation. They are thus fundamental players of the canonical and non-canonical functions of TJs.

4. Adherens junctions

In polarized epithelial cells, AJs are found immediately below TJs, in a belt-like structure that forms the ZA (Figure 2 and Figure 5A). The biogenesis of TJs and AJs is tightly coordinated and the two are functionally interconnected. However, in mature polarized epithelia, TJ and AJ proteins are effectively segregated into distinct macromolecular complexes (Rusu and Georgiou, 2020; Rouaud et al., 2020a). AJs mediate intercellular adhesion but are also a tensile network which, through their binding with actin and MTs, physically links cytoskeletons of neighboring cells and mechanically transduce signals between cells to drive cytoskeletal rearrangements. They are therefore at the core of tissue morphogenesis, homeostasis and plasticity. For example, AJ remodeling needs to be tightly regulated during gastrulation and neurulation, and their disruption leads to loosened cell contacts and disorganized tissue

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architecture (Nishimura and Takeichi, 2009; Meng and Takeichi, 2009; Takeichi, 2014). AJs also represent an important signaling hub for intracellular signaling and gene expression. They control the nucleo-cytoplasmic shuttling of the transcriptional co-activator YAP (Silvis et al., 2011; Schlegelmilch et al., 2011; Giampietro et al., 2015), and some AJ cytoplasmic components (ß- and p120-catenins) can translocate to the nucleus to associate with transcription factors (Valenta et al., 2012; Daniel and Reynolds, 1999; Daniel, 2007). In addition, AJs can modulate gene expression through the regulation of RhoA and Rac1 activity (Klezovitch and Vasioukhin, 2015; McCrea et al., 2009). Moreover, a functional RNA interference (RNAi) machinery has been detected at AJs (Kourtidis and Anastasiadis, 2018).

Dysregulation of these processes can be associated with pathogeneses such as cancers (Schackmann et al., 2013; Gavert and Ben-Ze'ev, 2007; Daulagala et al., 2019).

The structure of AJs comprises transmembrane components associated with cytoplasmic proteins that are connected to a thick circumferential bundle of actin filaments and to MTs (Figure 5B). Cryo-electron microscopy reveals AJs as apposing membranes from neighboring cells separated by a 15-25 nm intercellular space filled with rod-like structures extending from the cell surfaces and associated with cytoskeletons at the cytoplasmic face (Farquhar and Palade, 1963; Miyaguchi, 2000). Along the lateral contacts of polarized epithelial cells, and also in non-polarized cells such as fibroblasts or endothelial cells, adhesion sites are also present in spot-like or punctate structures for which the classification as AJs remains to be determined (Figure 5A,C) (Yonemura et al., 1995; Dejana, 2004; Takeichi, 2014; Rouaud et al., 2020a). Indeed, lateral contacts differ from the ZA (i) because of the absence of several cytoplasmic proteins that are exclusively present at the circumferential zonula and not laterally (Pulimeno et al., 2010; Meng et al., 2008; Mandai et al., 2015; Rouaud et al., 2020a), indicating thus a different architecture, and (ii) by their association with loosely organized actomyosin filaments terminating perpendicularly to cell-cell contacts and not associated into a circumferential peri-junctional belt such as the one occurring at the ZA (Takeichi, 2014; Citi and Kendrick-Jones, 1991; Yonemura et al., 1995; Rouaud et al., 2020a). Furthermore,

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receptors present all along cell-cell interface are differently stabilized between AJs and lateral contacts (Shah et al., 2018).

Figure 5. AJs and their molecular organization in polarized epithelial cells.

(A) Scheme illustrating the distribution of TJ, ZA and lateral contacts in polarized epithelial cells.

(B) Illustration of the AJ molecular organization highlighting the main transmembrane components, nectins and cadherins, and the associated cytoplasmic proteins. Nectin and cadherin complexes are linked through afadin-ß-catenin (ß) and PDZD11-PLEKHA7-p120-catenin (p120) interactions, further crossing via afadin binding to PLEKHA7. PLEKHA7 also binds to paracingulin (CGNL1) and nezha/CAMSAP3, this last bridging the AJs to MTs. ß-catenin interaction with α-catenin (α) connects AJs to actin cytoskeleton, such as afadin does. PLEKHA7 binds to phosphoinositides (PI) via its PH domain. (C) Confocal images (top view on the left, lateral view on the right) of polarized mouse cortical collecting duct epithelial cells (mCCD) showing the distribution of E-cadherin (red) at the ZA (white arrows) and along lateral contacts (yellow arrows). Blue staining shows nuclei (DAPI).

Nectins and E-cadherins are the two major adhesion transmembrane proteins present at AJs, providing calcium-dependent or -independent intercellular adhesion, respectively (Takai and Nakanishi, 2003; Yoshida and Takeichi, 1982; Yoshida-Noro et al., 1984). They are linked to

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several proteins present just below the membrane and forming the AJ cytoplasmic plaque that connects the transmembrane complexes together and connects AJs to actomyosin and MT cytoskeletons (Figure 5B).

4.1. The nectin/afadin complex

Nectins are immunoglobin (Ig)-like calcium-independent single-pass transmembrane adhesion proteins. The family comprises four members (nectin-1 to -4) and shares sequence homology with a related family of non-canonical components of AJs, the nectin-like molecules (Necls) (Takai and Nakanishi, 2003; Mandai et al., 2015). Nectins are detected at the initial, spot-like cell-cell contacts and subsequently recruit the cadherin complexes, and are thus crucial for junction formation (Asakura et al., 1999; Fukuhara et al., 2002a; Fukuhara et al., 2002b; Hoshino et al., 2004; Mandai et al., 2015; Takai et al., 2008). Nectins form homo-dimers in cis in the membrane of the same cell, and then make either homo- or hetero-trans- interactions with cis-dimers from the opposing cell (Samanta and Almo, 2015; Mandai et al., 2015; Takai et al., 2008). The N-terminal extracellular region of nectins contains three Ig-like loops, the first being required for the trans binding and the second and third for cis-dimers formation (Samanta and Almo, 2015; Mandai et al., 2015). At the extremity of their C-terminal cytoplasmic tail, nectins harbor a conserved four-residue (Glu/Ala-X-Tyr-Val) PDZ-binding motif that mediates the interaction with afadin (Takahashi et al., 1999). Afadin provides the connection of nectins to the actin cytoskeleton (Mandai et al., 1997) and makes the link with cadherin complexes through afadin interaction with α-catenin (Figure 5B) (Miyahara et al., 2000; Pokutta et al., 2002; Tachibana et al., 2000), which allows cadherin recruitment during junction formation. The PDZ-binding motif of nectin-1 and -3 also directly binds PDZD11 (Guerrera et al., 2016). This interaction prevents the proteasome-dependent degradation of nectins and further connects nectins to cadherin complexes and to MTs via the adaptor protein PLEKHA7 and the MT minus end-binding protein nezha/CAMSAP3 (Calmodulin-regulated spectrin-associated protein 3) (Figure 5B), which supports the efficient early assembly of junctions (Guerrera et al., 2016; Meng et al., 2008).

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Afadin is a multi-domain protein of large size (205 kDa) which contains two N-terminal Ras- associated domains, a fork head-associated domain, a dilute domain followed by a central PDZ domain and three proline-rich regions, a coiled-coil region, and an actin-binding C-terminal part (Mandai et al., 1997; Takai et al., 2008). Since the formation of junctions depends on initial nectin-mediated cell-cell adhesion followed by the recruitment of cadherin complexes to form mixed nectin- and cadherin-clusters, afadin plays a fundamental role by connecting the two complexes together, either directly via α-catenin (Miyahara et al., 2000;

Pokutta et al., 2002; Tachibana et al., 2000), or indirectly (Mandai et al., 1999; Ooshio et al., 2004; Asada et al., 2003; Guerrera et al., 2016) (Figure 5B). In addition, the formation of AJs is a prerequisite for the establishment of TJs apically with respect to AJs, and this process requires nectin/afadin-based cell-cell adhesion (Gumbiner et al., 1988; Ooshio et al., 2007;

Fukuhara et al., 2002b; Fukuhara et al., 2002a; Komura et al., 2008). Afadin is also molecularly linked to TJs. Indeed, it directly binds to ZO-1 and allows its recruitment to nascent AJs before its segregation apically at TJs, in cooperation with the binding of ZO-1 to α-catenin-cadherin complexes (Yokoyama et al., 2001; Ooshio et al., 2010; Fukuhara et al., 2002a; Maiers et al., 2013; Ando-Akatsuka et al., 1999; Itoh et al., 1997). Afadin also participates in the recruitment of JAMs, together with ZO proteins (Ebnet et al., 2000; Fukuhara et al., 2002a; Monteiro et al., 2013). Finally, afadin directly interacts with PLEKHA7 through either its N-terminal Ras- associated domains or central PDZ domain (Figure 5B), and recruits it to nectin-based junctions, promoting efficient formation of AJs (Kurita et al., 2013).

Despite their importance in cell-cell adhesion, the single KO of nectins in mice is not embryonic lethal, most probably due to the functional redundancy between different isoforms, but leads to developmental and functional abnormalities such as male infertility (Inagaki et al., 2006;

Mueller et al., 2003; Ozaki-Kuroda et al., 2002). In contrast, afadin-KO mice show embryonic lethality, with early embryonic cells showing an inability to form junctions and establish apicobasal polarity, highlighting the crucial importance of afadin for AJC formation and for cell-

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cell adhesion, cytoskeletal organization and cell polarity (Ikeda et al., 1999; Zhadanov et al., 1999; Mandai et al., 2013).

4.2. The E-cadherin/catenins complex

Cadherins are calcium-dependent adhesion proteins involved in the formation of cell-cell contacts in various cell and tissue types and are part of a large multigene family. The classical cadherins are characterized by the presence of five extracellular cadherin repeats (EC domains), a single transmembrane span and a C-terminal cytoplasmic tail (Rudini and Dejana, 2008). They comprise over 20 isoforms and were originally named according to the tissue of their predominant expression, such as E-cadherin and VE-cadherin in epithelial and vascular endothelial cells, respectively, or N-cadherin in the nervous system (Rudini and Dejana, 2008).

E-cadherin is a central protein in the architecture of epithelial AJs, since all AJs contain cadherin-catenin clusters while only a subpopulation additionally have nectin (Indra et al., 2013; Rusu and Georgiou, 2020). E-cadherin is essential for the assembly but is dispensable for the maintenance of cell-cell contacts, since reducing E-cadherin expression by RNAi does not impair AJs and TJs neither cell polarity but disrupts the re-establishment of junctions after their disassembly (Capaldo and Macara, 2007). This central role of E-cadherin in epithelial morphogenesis is confirmed by the embryonic lethality of E-cadherin-KO mice due to failure in the formation of the trophectoderm (Larue et al., 1994; Ohsugi et al., 1997). Upon formation of a cell-cell contact, after nectin-mediated initial adhesion, cadherins cluster at the AJs and then spread laterally, strengthening the contact (Asakura et al., 1999; Adams et al., 1998). Thus, in polarized epithelial cells, E-cadherin is localized at the ZA, associated with a circumferential peri-junctional belt, and also along lateral cell-cell contacts, connected to a more amorphous actin network (Figure 5A,C) (Takeichi, 2014). Cadherins form dimers in cis in each plasma membrane and interact in trans with dimers from the neighboring cell through their extracellular EC domains, which undergo calcium-dependent conformational changes (Takeda et al., 1999;

Yap et al., 1997; Yoshida and Takeichi, 1982; Yoshida-Noro et al., 1984; Pokutta et al., 1994).

The cytoplasmic tail of E-cadherin binds to proteins which stabilize E-cadherin and connect it

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to the actin and MT cytoskeletons (Figure 5B), acting in mechanotransduction, and which regulate intracellular signaling and gene transcription. This intracellular part of E-cadherin comprises juxtamembrane and catenin-binding domains, where p120-catenin and ß-catenin link, respectively (Yap et al., 1998; Ozawa et al., 1990; Aberle et al., 1994). ß-catenin interacts with the actin-binding protein α-catenin and thus bridges E-cadherin to actin cytoskeleton (Figure 5B) (Aberle et al., 1994; Rimm et al., 1995; Drees et al., 2005; Yamada et al., 2005;

Sakakibara et al., 2020). p120-catenin interaction stabilizes E-cadherin at AJs by masking the residues implicating in clathrin-mediated endocytosis and Hakai-dependent ubiquitination, thus preventing junction disassembly and stabilizing cell-cell adhesion (Yap et al., 1998;

Miyashita and Ozawa, 2007; Fujita et al., 2002; Troyanovsky et al., 2006; Davis et al., 2003;

Ishiyama et al., 2010). In addition, p120-catenin connects cadherins to MTs through PLEKHA7, which is in turn linked to nezha/CAMSAP3 (Figure 5B), further stabilizing E-cadherin at AJs (Meng et al., 2008). The central role of E-cadherin in cell-cell adhesion, and thus the formation and maintenance of epithelial versus mesenchymal phenotype, supports its function as tumor- suppressor. Accordingly, the loss of E-cadherin is relatively common in cancers of epithelial origin (Kourtidis et al., 2017a). However, E-cadherin can support tumor progression in some cancers, suggesting that E-cadherin-linked signaling processes are involved as well as adhesion-mediated collective cell migration (Rodriguez et al., 2012; Daulagala et al., 2019;

Friedl and Gilmour, 2009).

Four major catenins are localized at the ZA and along lateral contacts, similarly to E-cadherin (Figure 5A,C), and coordinate the dynamics and signaling of AJs. α-, ß- and γ-catenins were first identified as proteins present in E-cadherin immunoprecipitates (Ozawa et al., 1989;

Kemler and Ozawa, 1989). p120-catenin was subsequently identified based on sequence homology with ß-catenin (Reynolds et al., 1992). ß-, γ- and p120-catenins contain homologous armadillo repeats and directly bind E-cadherin, whereas α-catenin has a different structure, is indirectly linked to E-cadherin via ß-catenin and is an actin-binding protein (Rudini and Dejana, 2008).

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p120-catenin binds the cytoplasmic juxtamembrane domain of E-cadherin and ensures its stability at AJs by preventing its endocytosis and degradation (Yap et al., 1998; Miyashita and Ozawa, 2007; Fujita et al., 2002; Troyanovsky et al., 2006; Davis et al., 2003; Ishiyama et al., 2010). p120-catenin also bridges E-cadherin to MTs through PLEKHA7 and nezha/CAMSAP3 (Figure 5B), further supporting AJ integrity (Meng et al., 2008). p120-catenin is additionally present in a cytoplasmic pool which interacts directly with MTs (Yanagisawa et al., 2004) and affects cell morphology and motility by regulating the activity of Rho GTPases and thus the organization of actin cytoskeleton (Noren et al., 2000). This regulation of Rho GTPases by p120-catenin also supports AJ assembly by antagonizing Rho and Rac signaling at cell-cell contacts (Wildenberg et al., 2006). In addition, unbound p120-catenin can translocate to the nucleus where it binds the transcription factor Kaiso and releases its repressor activity (Daniel and Reynolds, 1999; Kelly et al., 2004). The fundamental importance of p120-catenin is confirmed by the embryonic lethality of KO mice and strong defects in conditional KO epithelial tissues (Davis and Reynolds, 2006; Perez-Moreno et al., 2006; Smalley-Freed et al., 2010), as well as its dual role in cancers. Indeed, on the one hand, p120-catenin exerts as a tumor suppressor, through the stabilization of E-cadherin at AJs which supports cell-cell adhesion, and through its association with PLEKHA7 at the ZA, which is linked to a RNAi machinery that suppresses the expression of pro-tumorigenic factors (Kourtidis et al., 2013; Kourtidis et al., 2015a; Kourtidis et al., 2017b). On the other hand, a basolateral pool of p120-catenin which is tyrosine-phosphorylated by Src promotes anchorage-independent cell growth and acts thus as a tumor promoter (Kourtidis et al., 2015a; Kourtidis et al., 2015b).

ß-catenin is linked to the C-terminal cytoplasmic catenin-binding domain of E-cadherin via its central armadillo repeats and to the actin-binding protein α-catenin through its N-terminal region (Ozawa et al., 1990; Aberle et al., 1994). ß-catenin promotes cadherin-mediated cell- cell adhesion by participating in E-cadherin trafficking from the endoplasmic reticulum (ER) to the plasma membrane (Chen et al., 1999) and by bridging E-cadherin to the actin cytoskeleton through α-catenin (Figure 5B) (Rimm et al., 1995; Drees et al., 2005). Afadin reinforces this

The Role of WW-PLEKHAs and PDZD11 in the Trafficking of the Copper Pump ATP7A and in Copper Homeostasis

Thesis

Reference

The Role of WW-PLEKHAs and PDZD11 in the Trafficking of the Copper Pump ATP7A and in Copper Homeostasis

The Role Of WW-PLEKHAs And PDZD11 In The Trafficking Of The Copper Pump ATP7A And In Copper Homeostasis

THÈSE

Sophie SLUYSMANS

Acknowledgments

Résumé

Abstract

Table of contents

List of Figures

List of Tables

Abbreviations

Introduction

1. Cellular junctions

2. Cell-cell junctions: structure and canonical functions

3. Tight junctions

3.1. Transmembrane proteins of tight junctions

3.2. Tight junction cytoplasmic proteins

4. Adherens junctions

4.1. The nectin/afadin complex

4.2. The E-cadherin/catenins complex