Open access peer-reviewed chapter

Lysosomal Degradation of Junctional Proteins

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Catalina Flores-Maldonado, Odette Verdejo-Torres, Jessica Campos- Blázquez, Agustín Ruiz Cabrera, Vicky García-Hernández, Ruth Rincón-Heredia and Rubén G. Contreras

Submitted: October 11th, 2016 Reviewed: April 21st, 2017 Published: August 30th, 2017

DOI: 10.5772/intechopen.69370

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Epithelial cells develop tight junctions (TJs) and cell polarity. Both properties are sensitive to environmental signals such as the epidermal growth factor (EGF) and the cardiotonic steroid ouabain. EGF is regarded as the main protector against injuries in epithelia, and ouabain is a hormone that regulates blood pressure, natriuresis, cell survival, and cell adhesion. After treatment with epidermal growth factor or ouabain, epithelial dog kidney MDCK cells undergo a drastic remodeling that includes changes in the transcription, translation, localization, and degradation of cell junction proteins. Degradation of these proteins involves selective and nonselective autophagy as well as endocytic lysosomal and proteasomal routes. The remodeling mechanism of tight junction’s proteins includes the activation of Src and ERK1/ERK2 kinases, the phosphorylation and translocation into the nucleus of the transcription factor STAT3, the activation of PKC to induce the endocytosis of claudin-2, and the delivery of this protein to the lysosomes. Whole communicating junctions and desmosomes are internalized by one cell and sent to degradation by nonselective autophagy. Nonselective and selective autophagies in epithelial cells are very context dependent; nevertheless, it is clear that, together with endocytic lysosomal and proteasomal degradation, they play a key role in the remodeling and functioning of cell junctions.


  • claudin-2
  • ouabain
  • EGF
  • MDCK
  • tight junctions

1. Introduction

When multicellular organisms left the sea of constant chemical composition to conquer the land, it became necessary to develop mechanisms to maintain a constant internal milieu similar to the sea that was left behind. Epitheliawere fundamental in terrestrial conquest because these tissues avoid the loss of water, transport vectorially substances to introduce nutrients into the body, eliminate waste substances, and protect the individual from pathogens and their toxins [1]. These functions of epithelia depend on two basic characteristics of the cells that constitute them; the first one is plasma membrane polarity, which consists on the division of the plasma membrane in an apical, a lateral, and a basal domains, each one with characteristic structure, composition, and function [2]. Due to its continuity, the lateral and the basal domains are often referred as the basolateral domain. The second characteristic of epithelia is cellular junctionsthat bind epithelial cells to each other and to the substrate (Figure 1) [1, 3]. Adhesion between neighboring cells depends on the junctional complex, which is a group of intercellular junctions that includes the tight junctions (TJs), adherens junctions (AJs), desmosomes (Des), and communicating or GAP junctions (GAPJs) [4]. Attachment to the substrate requires the formation of two additional cellular junctions: focal adhesions (FAs) and hemidesmosomes (HDes) (Figure 1) [5, 6]. Cellular junctionsand plasma membrane polarityare crucial requirements for epithelial function and influence each other in complex ways, for example, the formation of a specific cell junction is the result of the polarized expression of many proteins and lipids; conversely, the development of cell polarity requires cell adhesion, initially to the extracellular matrix and then to the neighboring cells [710], and the development of AJs is, in general, a prerequisite for the formation of the other intercellular junctions [11, 12].

Figure 1.

Cell junctions and polarity in epithelial cells. Substances cross the epithelial layer through the transcellular or paracellular routes or by transcytosis. Each scheme depicts the main molecular components of cellular junctions and its organization in the membrane. MAGI are inverted membrane-associated guanylate kinase-like proteins, ZO-1, -2 and -3 are Zonula Occludens-1, -2 and 3, Src is the protein homologous to the Rous sarcoma virus kinase, FAK is the focal adhesion kinase, LSR stands for the lipolysis-stimulated lipoprotein receptor, JAM is the junctional adhesion protein.

The complex organization and regulation of cell junctions and cell polarity in epithelial cells are adaptations to perform vectorial transport. A given substance crosses epithelial layers either through the transcellular pathway, using polarized channels, transporters, and pumps [13, 14], the paracellular pathway, constituted by TJs and the intercellular space [15], or both (Figure 1). The contribution of each pathway depends on the kind of substance transported, the cell type that executes the transport, and the physiological state of the transporting cell. In bowels and kidneys of mammalians, the transcelllular pathway accounts for most of the glucose transport [13], while flying vertebrates, perhaps because they have a smaller intestine than other vertebrates, relay in a majoritarian paracellular transport [16]; most Ca2+ is reabsorbed at the thick ascending limb of Henle’s loop through the paracellular pathway by a specific type of claudin (CLDN), claudin-16, a TJ protein that forms Ca2+- and Mg2+-selective pores [17], while glucose is transported predominantly through the transcellular pathway by polarized SGTL and GLUT transporters driven by the Na+ concentration gradient [13, 18]. There are also two more processes to cross epithelia and endothelia: whole cells cross epithelia, as shown in spermatogenesis in which immature sperm cells open its way through the junctional complex of epithelial Sertoli cell monolayers [19] and in diapedesis, where neutrophils and leucocytes open transitorily the junctional complex and accede to the internal media [20]. Moreover, some substances and even viruses cross epithelia by transcytosis(Figure 1), a process that consists in the incorporation of the material to a vesicle generated in one plasma membrane domain that is, later on, translocated and fused to the opposite membrane domain where it delivers its content [21].

Recent research demonstrated that cell junctions are signaling stationsthat inform the adhesion status of the cells to cytoplasmic and nuclear mechanisms so they can induce proper responses [22, 23]. Contact inhibitionand healing of a woundillustrate this property of cell junctions: loose AJs, or their absence by the lack of E-cadherin, an essential transmembrane protein of this cellular junction, trigger a period of high cell proliferation and migration through the activation of the Hippo signaling pathway and stop only when the normal size of the organ and a strong adhesion between epithelial cells are reached, a process referred to as contact inhibition[24]; when an epithelium is wounded, the cells at the front of the wound lose their cell junctions, start to proliferate, and migrate to heal the wound and extracellular protective factors, such as the epidermal growth factor (EGF), ouabain (OUA), and marinobufagenin, favor this reparation [25, 26]. Several membrane-associated proteins of cell junctions, notably β-catenin [27], a protein of the AJs, and zonula occludens-2 (ZO-2), a protein of TJs [28], are transcriptional cofactors that regulate the expression of proliferation-related genes, for instance, cyclin D1 and myc; conversely, transcription factors such as Jun, Fos, and ZONAB reside transitorily at the cytoplasmic region of the TJs [29, 30] and, at least ZONAB, downregulates paracellular permeability [30].

Cellular junctions and plasma membrane polarity are highly regulated. For example, a progressive conversion of renal intercalated cells of the collecting tubules from α to β type comprises the inversion of the apical H+-ATPase and a basolateral Cl/HCO3+ exchanger polarity, in response to the increase in the expression of hensin, a protein of the extracellular matrix [31, 32], and cysts embedded in collagen displace their TJs from the vicinity of the lumen toward the proximity of the external surface [33, 34].

Cell junctions and plasma membrane polarity are crucial for the normal physiology of the organism, and its failure in several pathologies has disastrous consequences. To start with, it is common that the genetic elimination of crucial proteins, such as E-cadherin from the AJs or ZO-2, is lethal at embryonic stages [35, 36], but whenever an epithelial adhesion protein is not expressed, epithelia compartmentalization and vectorial transport are lost. For example, in hereditary familial hypomagnesemia with hypercalciuria and nephrocalcinosis, the lack of CLDN-16 and CLDN-19 impairs Ca2+ and Mg2+ reabsorption in the kidney [17, 37]; in cholestatic children’s liver disease, the absence of ZO-2 and TJs provokes the invasion of bile salts into the blood [38]; in pemphigus vulgaris, the depletion of Des by autoantibodies against the desmosomal cadherin desmoglein-3 results in the formation of skin blisters [39], which can also appear if HDes are disassembled by mutations in the integrin β4, an adhesion molecule of this cell junction [40]; loss of adhesion and augmented proliferation in colon cancer are elicited by mutations that increases the cytosolic and nuclear pools of β-catenin [27]; infection and inflammation boost diapedesis[20]; rotaviruses that cause diarrheas open TJs and accede to their basolateral receptors by exposing its VP8 protein, which bear small peptides with sequences identical to some regions of occludin, an integral membrane protein of the TJs [41]; and the exogenous expression of the E6 oncoprotein of the high-risk papilloma virus type 16 induces the translocation of β-catenin into the nucleus to increase proliferation in the skin [42].


2. Cell junctions have a general common layout

All junctions have a similar structural layout: they have transmembrane proteins that are the receptors for adhesion, and a series of membrane-associated proteins that bind the cytoplasmic aspect of transmembrane receptors to the actin, tubulin, or cytokeratin cytoskeleton to provide mechanical strength. Besides cell adhesion, cell junctions are sensors that inform, in and out, the state of extracellular environment to modulate cell’s proliferation, differentiation, and fate. Given that lysosomes are of paramount importance for cell junctions and plasma membrane polarity, it is necessary to briefly review the degradation routes where this organelle intervenes.


3. Lysosome degradation pathways

Lysosomes are major degradative organelles of eukaryotic cells. They were first identified as cell compartments enriched in hydrolases [43], but now they are also recognized as providers of building blocks during starvation and powerful stations to sense nutrients and regulate transcription and cellular homeostasis [44]. Lysosomes have a highly acid lumen (pH 4.5–5.0) produced by a vacuolar H+-ATPase. The acidic pH is necessary for the hydrolysis of waste materials and drives the transport of sugars, amino acids, nucleotides, and lipids, through the single membrane of the organelle for recycling [45]. The lysosomal membrane owes its resistance against the activity of the hydrolases that it contains, to the expression of a prominent glycocalyx in its inner surface, formed by glycosylated transmembrane proteins such as the human LIMP-2 and its homologues in Caenorhabditis elegansSCAV-3 [46].

Many intracellular proteins are ubiquitiated and degraded in the proteasome (Figure 2, 1). There are also several routes to deliver cellular material into the lysosomes: an endosomal-lysosomalroute, similar to the secretory route, that consists in the ubiquitin (Ub) or clathrin mediated endocytosis of the protein to be degraded, the posterior fusion of the vesicles containing this protein to the early endosomes (EE), which in turn may mature to late endosome (LE, 2) or produce multivesicular bodies (MVB, 3) and fuse with the lysosome (Figures 2 and 3) [47, 48]; the microautophagydegrades intracellular proteins engulfed by the lysosome (Figure 2, 4) [49], the nonselective macroautophagydegrades big intracellular waste (Figure 2, 6) [52, 53] (Figure 2,5). Endocytosisrequires the ESCRT machinery and the activity of several Rab GTPases that tether and dock the SNARE fusion machinery [54]. Rab5 participates in the fusion of clathrin-coated vesicles with early endosomes (Figure 2, EE) [55, 56], Rab4 in the recycling of proteins to the plasma membrane [57], and Rab7 in the transport from early to late endosome and lysosome biogenesis and maintenance [58]. Nonselective macroautophagyconsists of the engulfment of intracellular waste material by a phagophore, a double-membrane structure possibly derived from the endoplasmic reticulum [53]. Since its appearance and through all its maturation, the phagophore incorporates the microtubule-associated protein 1A/1B light chain 3 (LC3). LC3 exist in a cytosolic form (LC3I) that, at the onset of autophagy, is conjugated with the lipid phosphatidylethanolamine to form LC3 type II (LC3II). Selective macroautophagyis triggered with the purpose to degrade a specific protein. This protein is first ubiquitinated by ligases such as Beclin1, which is then bound to the ubiquitin-binding sequestosome 1 protein (p62), which in turn binds to LC3II. The protein can also be bound, indirectly, to p62 through the autophagic cargo receptor protein (ACR, Figure 2, 7). Progressive encirclement of the waste material by the developing phagophore conduces to the formation of the organelle called autophagosome, which later on fuses with the lysosome to become an autophagolysosome. Fusion of the autophagosome to the lysosome requires a low acidic pH of the lysosome and is inhibited by drugs that access the lumen of this organelle and neutralize its pH, such as chloroquine and NH4Cl [51], or inhibit the lysosomal V type H+-ATPase, as bafilomycin A1 [59]. The lysosome itself may invaginate small cytoplasmic waste materials, in a process similar to the formation of multivesicular bodies from the late endosome [60]. In the chaperone-mediated autophagy, cytosolic unfolded proteins that contain a KFERQ-like pentapeptide bind to the lysosome receptor Lamp-2 and are directly translocated into the lumen of the lysosome through a multimeric translocation complex [50, 53].

Figure 2.

Protein Degradation pathways. (1)Proteasome pathway, used for degradation of the ubiquitin (Ub) bound cytosolic proteins. (2) Endocytic pathway, involving endocytic vesicles that fuse to the early endosome (EE), progress to the late endosome (LE), multivesicular bodies (MVB, (3) and finally to the lysosome. (4)Microautophagy, the lysosome engulfes some cytosolic proteins. (5)Chaperon mediated autophagy, allows the passage of cytosolic proteins via translocons. (6)Nonselective macro autophagy, large intracellular waste material is engulfed by the phagophores that becomes, later on, the autophagosome (AP). This organelle then fuses with the lysosome to form the autophagolysosome (APL). (7)Selective macro autophagy, intracellular waste material is ubiquitinated, bound to autophagic cargo receptors (ACR), LC3 or p62, and then engulfed by the phagophores to continue the autophagy route. RE is the recycling endosome.

A central regulator of lysosomal activity, particularly autophagy, is the target of rapamycin (mTOR), a multi-protein complex that includes the kinase mTOR itself, inhibited by rapamycin, the raptor adaptor, two intrinsic inhibitors of mTOR activity, DEPTOR and PRAS40, and a G-protein. The mTOR complex senses energy and nutrient availability, growth factors, and stress conditions to modify cell growth and proliferation. In normal conditions, mTOR localizes in the cytosol and triggers anabolic programs, like mRNA translation. Under starvation, mTOR is translocated to the cytosolic side of the lysosome membrane, where it initiates catabolic processes like autophagy [44].

A growing body of evidence suggests that lysosomes can function as Ca2+ stores and contact intimately to the endoplasmic reticulum, the peroxisome, and the mitochondria to deliver necessary lipids [61, 62] and that lysosomes can fuse to the plasma membrane to pour hydrolytic enzymes in the extracellular media that modify the extracellular matrix and induce differentiation [59]. Lysosomes induce cell death when its membrane is permeabilized and hydrolases such as cathepsin B, a Ca2+-sensitive protease, are released in the cytoplasm. Cell death induced by lysosomal damage is observed in tissue remodeling, elimination of excessive intracellular waste or metals, and the immune response to intracellular pathogens and neurodegenerative diseases [63, 64].

To maintain compartmentalization and vectorial transport in epithelial cells, the synthesis and degradation of adhesion proteins must be closely coordinated. Nevertheless, epithelial cells must have certain degree of plasticity to modify cell junctions in response to the variable environment. Lysosomal activity is crucial in both situations.


4. Role of lysosomes in the degradation of tight junction proteins

4.1. Biology of tight junctions

TJs, also known as zonula occludens(ZO), are formed at the most apical region of the lateral plasma membrane domain [4], constitute the frontier between the apical and the basolateral domains, and obliterate the intercellular space at this region, converting the epithelia in an effective barrier against the free diffusion of ions and other substances [3]. This cellular junction has two main functions: it is a gatethat transports ions and other substances through the paracellular pathway [6567] and a fencethat impairs the diffusion of integral membrane proteins and lipids from the apical to the basolateral domain and the other way around [68, 69], a property that helps to maintain cell polarity [8]. In the case of lipids, its impairment is restricted to those from the exoplasmic leaflet of the plasma membrane. In some experimental conditions, TJs can be a bridgethat allows the diffusion, in the plane of the membrane, of membrane liposoluble probes and of phosphatidylinositol bisphosphate (PIP2), from one epithelial cell to its neighbors [7074]. TJs are composed of three main types of integral proteins that perform the task of adhesion receptors: the four-transmembrane-domain family of CLDNs [75] and TAMP proteins, that includes occludin and tricellulin [76], the single transmembrane domain proteins of the junctional adhesion molecule (JAM) family and the lipolysis-stimulated lipoprotein receptor (LSR) (Figure 1) [7782]. CLDNs constitute the paracellular ion and water channels and barriers [17, 65, 83] and confer to the TJs and the epithelia its specific paracellular permeability and selectivity, depending on the type of CLDNs expressed [17, 65, 8385]. The integral membrane proteins of the TJs bind to a cluster of three cytoplasmic proteins of the ZO, named ZO-1, ZO-2, and ZO-3 (Figure 1, green). These proteins belong to the MAGUK family and are platforms for protein-protein interaction [8688]. ZO’s plaque proteins in turn bind transmembrane once to the actin cytoskeleton [89]. There are a number of other integral and associated proteins that are nonessential or reside transitorily at TJs but, nevertheless, are important for vesicular trafficking and signaling [78]. In insects and mollusks, the paracellular pathway is controlled by the septate junctions, structures that have different molecular compositions and structures and are located bellow the AJs [90]. Based on the crystal structure of the euglena tetraspan protein IP39, another member of the group of proteins to which CLDNs belong, the PMP22/EMP/MP20/Claudin superfamily, it has been suggested that CLDNs may be originated from an ancestral four-transmembrane-domain protein, similar to IP39, which may had played a role in the peculiar tortuous movement of this photosynthetic protist [91].

Epithelia adjust the permeability of their paracellular route in response to physiological requirements, pathological conditions, and pharmacological challenges. One simple way to gaze epithelial permeability is to measure the transepithelial electrical resistance (TER) [92]: the higher the value of TER, the lower the paracellular permeability. On this regard, the renal system is very illustrative. Human kidneys filtrate 170 l of plasma but secrete only 1.7 l of urine. Water, proteins, sugars, and ions from the glomerular filtrate are reabsorbed, and the filtrate is steadily concentrated along the nephron. The epithelia that line this tubular surface in vertebrate species gradually increase their TER from approximately 10 Ω cm2 at the proximal convoluted tubule [93, 94] to several thousands of Ω cm2 at the collecting duct [95, 96] and up to hundreds of thousands of Ω cm2 at the bladder [97, 98]. A number of epithelial cell adaptations account for by this TER gradient: increments in cell size, reduction of the junctional membrane tortuosity, a progressive increase in the structural complexity of TJ strands, and the expression of a specific set of CLDNs in each nephron segment [15, 65]. CLDN-2 induces a low TER phenotype in renal MDCK cells [99], from cation and water-selective channels [83, 100, 101], and it is expressed in proximal tubules [102104], where it is necessary for the uptake of Na+, water and, likely, Ca2+ [105]. CLDN-4 induces a high-resistance phenotype upon the epithelial cells that express it [106108], including those at the distal nephron segment epithelium [102104].

4.2. Autophagy of CLDN-2 participates in the differentiation of epithelia induced by the epidermal growth factor and exerts a protective effect

The fluids that bathe apical membranes, such as urine, semen, and milk, are radically different from each other, but the interstitial milieu that contacts the basolateral membranes has a constant composition maintained by homeostatic mechanisms. This difference suggests that substances in the apical media might regulate specific epithelial properties. Several substances in the extracellular milieu induce TER changes in canine MDCK cells [109]. One of them is EGF [110], a substance previously known to increase the TER of epithelial kidney pig LL-CPK1 cells [111]. Urinary EGF reduces the cellular CLDN-1 and CLDN-2 protein level and increases CLDN-4 one [110]. EGF decreases the cellular level of CLDN-2 [112] through the simultaneous activation of Src kinase, extracellular regulated kinases 1/2 (ERK1/2) [113, 114], and the transcription factor STAT3 [114] that, in turn, may accelerate clathrin-mediated endocytosis and lysosomal degradation of CLDN-2 [113], block CLDN-2 [115], and trigger CLDN-4 [116] transcription in MDCK cells. In lung cancer cells though, EGF increases CLDN-2 through the activation of the EGF/EGFR/MEK and cFos pathway [117]. It would be interesting to find out the molecular mechanisms that fail in cancer and provoke the opposite response.

The response elicited by EGF is transient, reaches a maximal value of TER at 15 h, and slowly decreases to control values at 24 h. This downregulation is provoked by the induction of the synthesis of prostaglandin E2 by the EGF itself that increases AmpC production, which in turn blocks the activation of ERK1/2 [118].

It is not clear which vesicular compartment participates in the CLDN-2 degradation induced by EGF. The observation that the knockdown of Rab14 induces the lysosomal degradation of CLDN-2 in MDCK cells [119] opens the possibility that EGF somehow be able to inactivate this Rab protein.

The induction of CLDN-2 downregulation by EGF is blocked by bafilomycin A1 and chloroquine, indicating that it may be performed by autophagy (Figure 3). A schematic representation of the mechanisms of EGF effect on CLDNs is shown in Figure 4. The induction of selective autophagy by EGF can be seen as a differentiation or protective effect. In this respect, autophagy has been observed in Caco-2 cancer colon cells deprived of nutrients, where selective autophagy of CLDN-2 is activated, resulting in an increase in TER [120]. Moreover, in porcine gut IPEC-1 epithelial cells, the deprivation of nonessential amino acids induces an apoptotic process that degrades CLDN-1 and ZO-1, but if autophagy is inhibited with 3MA, degradation of adhesion proteins and apoptosis is potentiated, indicating that autophagy has a protective role in these cells [121]. Finally, the injured spinal cord in rats induces the degradation of p120 and β-catenins, as well as CLDN-5 and occludin, in blood vessels of endothelia. This degradation is performed through selective autophagy, considering that these proteins associate to LC3II and p62. The administration of retinoic acid potencies autophagy and improves movement of the injured rats [122].

Figure 3.

Epidermal growth factor (EGF) induces CLDN-2 degradation in a bafilomycin 1A-sensitive manner. Epithelial dog kidney cells (MDCK) confluent monolayer grown on filters were incubated 15 h in control condition, EGF, bafilomycin A1 (Baf), or EGF plus Baf. (A) Transepithelial electrical resistance measurements. (B) Densitometric analysis of the cellular content of CLDN-2 measured by immunoblot. (C) CLDN-2 Immunofluorescence of cells incubated in the indicated conditions.

Figure 4.

EGF triggers a Src-ERK1/ERK2-STAT3 cascade to induce the degradation of CLDN-2 in the lysosomes. Occupancy of EGFR by its ligand induces the simultaneous phosphorylation of the kinases Src, ERK1/ERK2, as well as the phosphorylation of the transcription factor STAT3. The phosphorylated STAT3 is translocated into the nucleus, where it upregulates the transcription of CLDN-4 at the same time downregulates that of CLDN-2. In the cytoplasm, the same cascade plays a role in the induction of CLDN-2 endocytosis and CLDN-4 exocytic fusion, events that result in the lysosomal degradation of CLDN-2, an insertion of CLDN-4 at the TJs, and an increment of the degree of sealing of the TJs.

4.3. Ouabain induces degradation of CLDN-2

In the eighteenth century, William Withering used extracts of the herb foxglove (Digitalis purpurea) to treat successfully what is now called congestive heart failure [123]. The active principle involved in this therapy is digoxin, a member of the family of compounds named cardiotonic steroids, which bind and inhibit the Na+,K+-ATPase [124] and of which ouabainis the prototypic compound. This enzyme is expressed in the plasma membrane of most animal cells, where it transports intracellular Na+ out of the cells, in exchange of extracellular K+ toward the cell interior, against their concentration gradients, in a reaction driven by ATP hydrolysis. The inhibition by OUA became the main criterion in determining whether a given flux depended upon an active transport. Animals, including humans, produce OUA and other similar substances such as marinobufagenin and proscillaridin that are synthesized in the hypothalamus [125] and the adrenal glands [126]. Cardiotonic steroids are now considered hormones [127, 128] that regulate salt-sensitive blood pressure [129], salt handling in the kidney [130], and sodium homeostasis [131]. OUA effects require binding to the Na+,K+-ATPase and the activation of intracellular pathways and genes [132], including ERK1/2 [133] and the inositol (3,4,5)-tris-phosphate receptor [134, 135], indicating that Na+,K+-ATPase is a receptor that transduces cardiotonic steroid occupancy into intracellular mechanism. Interestingly, signaling proceeds even when transport activity is completely inhibited [136138].

The activation of the Src-EGFR-ERK1/2 cascade by OUA regulates cell adhesion in a concentration-dependent manner: 10 nM OUA, a concentration near the hormonal level, increases the degree of sealing of the TJs, inducing the transcription, translation, and expression at the TJs of CLDNs [139], and 300 nM or higher concentrations of OUA promote cell detachment resulting from TJ, AJ, De, GAPJ, and FA disassembly, endocytosis, and posterior degradation of their cell adhesion molecules [140]. Occludin, CLDN-2, and CLDN-4 endocytoses are clathrin-dependent [141]. 300 nM but not 10 nM OUA increases p62 signal and its colocalization with CLDN-2 in MDCK cells; degradation of CLDNs at 300 nM OUA is inhibited with NH4Cl and bafilomycin A1 [141], suggesting that ouabain activates CLDN-2 degradation through autophagy (Figure 5). OUA increments the size of intracellular structures that bind an antibody against Rab11, a recycling endosome marker, indicating that OUA is not inducing recycling of CLDN-2 (Figure 6). The mechanism of OUA action is shown in Figure 7.

Figure 5.

High ouabain concentrations increase autophagy. Control MDCK cells have their CLDN-2 localized at the TJs, in a normal quantity, and in the cytoplasm in numerous spots (green); p62 shows no colocalization with CLDN-2. Upon incubation in media containing ouabain (OUA) 10 nM images remains unchanged, indicating that low OUA concentration does not activate autophagy. On the contrary, the incubation with OUA 300 nM decreases CLDN-2 all around the cell and increases p62 that colocalizes with internalized CLDN-2 (white arrows). This result supports the observation that 300 nM OUA increases autophagy of CLDN-2. Confluent monolayers of MDCK cells were grown on coverslips overnight and then incubated with control media, ouabain 10 nM or 300 nM for 20 h. Barr corresponds to 10 μm.

Figure 6.

Ouabain does not induce recycling of CLDN-2. MDCK cells monoloyers were plated on glass coverslips overnight and incubated in control conditions (control) or in media with OUA 10 or 300 nM for 6 h. Cells were stained with antibodies against CLDN-2 and Rab11, a small GTP-binding protein of the recycling endosome. At this time, CDLN-2 has not been degraded yet and does not colocalize with Rab11 at any condition, suggesting that CLDN-2 is not internalized through the recycling endosome. Nevertheless, ouabain 10 nM decreases the intensity of the signal and the number of spots observed in the cytoplasm, implying that low OUA concentrations decrease recycling, while 300 nM increases the signal as it corresponds to cells with very active endocytosis. These results indicate that under OUA stimuli there seems to be a very active endocytic pathway, but CLDN-2 is not being recycled nor internalized through it.

Figure 7.

A high concentration of ouabain induces endocytosis and lysosomal degradation of claudins. OUA induces the formation of the signalosome (structure enclosed by the interrupted line), a caveolar complex including some Na+,K+-ATPases, and their associated Src and EGF receptors (EGFR). OUA activates the Src-ERK1/ERK2 pathway, which induces the clathrin- and dynamin-dependent endocytosis of TJ components. Our results indicate that there are two types of endocytic vesicles: one containing a core complex with essential TJ proteins, such as ZO-1, OCLN, and CLDN-4, and a second one consisting of components that confer a differentiated functional characteristic to TJs, such as CLDN-2, that makes TJs permeable to water and Na+. Src-ERK1/ERK2 pathway is also required to reduce CLDN-2 and ZO-1 mRNA levels. Surprisingly, during the OUA-induced aperture of the TJs, the cellular content of CLDN-4 and OCLN mRNAs increases.TJP1is the gene that codes for ZO-1, and EGFR is the receptor of EGF. Ionic imbalance, indicated by the lowering of K+, may also activate ERK1/2.

The final outcome of the treatment with high concentrations of OUA is the detachment and death of OUA-sensitive cells [132, 137, 140, 142]. Ionic imbalance that results from the inhibition of the enzyme has been considered the prime cause of cell death given the fame of the Na+,K+-ATPase as an ion transporter. However, cells do not detach when they are cultured in low K+ medium [137], which mimics the diminished [K+]i content induced by OUA. The cytotoxic action of OUA in humans and rodent cells depends on the features of the type α subunit expressed, rather than by any downstream components of the cell death machinery [142]. In this respect, epithelial cells expressing a OUA-resistant isoform of Na+,K+-ATPase do not detach when they are incubated in low K+- or K+-free medium [137, 143]. Therefore, ionic imbalance by itself is not sufficient to detach cells; OUA and the activation of kinases (p38 tyrosine kinases, Src, and ERK1/2) are necessary (Figure 7), a finding that agrees with the triple role of the Na+,K+-ATPase: transporter, signaling receptor, and cell-cell adhesion molecule [144].

4.4. Autophagy plays a role in claudin-16 degradation in lysosomes

Renal hypomagnesemia with hypercalciuria and nephrocalcinosis is an autosomal recessive disease characterized by abundant renal Mg2+ and Ca2+ wasting that causes renal parenchymal calcification and renal failure. It can only be cured through renal transplantation. The illness results of the lack of stable expression of CLDN-16 and/or CLDN-19 caused by mutations in CLDN-16and CLDN-19genes [145, 146]. Patients with a mutation in CDLN-19 show hypomagnesemia, yet this symptom is accompanied by malfunction of several glands, as well as visual impairment [146, 147]. Claudin-16 protein forms paracellular Ca2+ pores, while CLDN-19 forms anion impermeable TJs [148, 149]. Both CLDNs form stable membrane dimers able to conduce cations which are bound to the dimers of the neighboring cell. Dimer formation depends on the lateral interaction of their third and fourth transmembrane domains of both CLDNs. Mutations in these domains, which impair dimerization, decrease transepithelial permeability [150]. Most known disease-related mutations of CLDN-16 provoke protein misfolding and induce the accumulation of the defective protein in the endoplasmic reticulum and its degradation in the proteasome [151, 152]; other CLDN-16 mutants reach the plasma membrane but are unable to bind the ZO-1 scaffold; consequently, they are efficiently endocyted, delivered to the lysosome, and degraded there [151, 153]. Inhibitors of endocytosis may provide novel therapeutic strategies [152]. CLDN-16 phosphorylation of serine 217 stabilizes it in TJ, but when this phosphorylation is inhibited, CLDN-16 is dissociated from ZO-1 and sent to degradation in lysosome, which decreases TER and increases the transport of Mg2+ from the apical to the basolateral domain [154]. However, phosphorylation can also induce degradation in lysosomes or proteasomes [151]. Thus, in salivary glandular epithelial cell, SMG-C6, the activation of M3 muscarinic receptor with carbachol switches on ERK1/2, which in turn phosphorylates CLDN-4 at the serine 195. The phosphorylated CLDN-4 is endocyted in clathrin-coated vesicles and subsequently degraded in the proteasome, a phenomenon that decreases TER [155].


5. Role of lysosomes in the degradation of adherens junctions

5.1. Biology of adherens junctions

In chordates, AJs are Ca2+-dependent cell-cell adhesions between neighboring epithelial cells at the lateral domain, immediately below the TJs (Figure 1, red). In prechordates, AJs present an inverted localization with respect to the TJs: AJs are the most apical junction of the lateral membrane, placed over the septated junctions. AJs consist of the nectin-afadine and the cadherin-catenin complexes. The nectin’s complex forms a scaffold necessary for the assembly of the AJs [156159], whereas the cadherins serve as homotypic adhesion receptors [160, 161]. The associated plaque proteins catenins and afadins, in turn, bind the receptors to the cytoskeleton of actin [162, 163]. The homotypic adhesion of cadherin plays an important morphogenetic role because it underlays the selection and association of cells of the same type to form specific tissues, a process denominated “cell sorting” [164]. Based on the fact that there are numerous cadherins in the unicellular choanoflagellate Monosiga brevicollis, and that some of these proteins are expressed on the collar, a structure rich in sticking microvilla that traps and endocyte bacteria from the media to get nutrients, it has been suggested that cadherin-based intercellular adhesion may be originated from the co-option of the “trapping food” function of ancestral cadherins of unicellular organisms [165].

5.2. Endocytic route and selective autophagy degrade key proteins of adherens junctions

In a normal epithelium, β-catenin is mostly associated to E-cadherin at the plasma membrane, and the cytosolic pool of β-catenin is kept low by degradation in the proteasome. However, a proliferation signal, triggered by a Wnt ligand, impedes the β-catenin degradation and induces its accumulation in the nucleus to activate proliferation (Figure 8B) [27]. E-cadherin is degraded by lysosomes through an endocytic route [166]. The cytoplasmic domain of E-cadherin has an endocytosis signal that is normally masked by α, βand p120 catenins [167]. The induction of E-cadherin endocytosis and degradation starts with the tyrosine phosphorylation of E-cadherin by the kinase Src that promotes the disassembly of the cadherin-catenin complex. Then, the phosphorylated E-cadherin is ubiquitinated on the cytosolic domain by the ubiquitin ligase Hakai. The adhesion protein is then endocyted and delivered to the lysosomes via early and late endosomes (Figure 8A) [168]. β-Catenin may be degraded through an alternative mechanism. Thus, when there are nutrients available, a low normal level of autophagy results from the fact that most β-catenin is repressing the transcription of p62. Under starvation, β-catenin interacts with LC3II and is itself targeted for autophagic degradation (Figure 8B) [169]. During embryonic development, cadherin-6 restrains autophagy and drives an epithelial-mesenchymal transition (EMT) to allow a mesenchymal migratory phenotype, which is exacerbated in thyroid cancer [170]. As it happens with other junctions, the role of β-catenin is context dependent. The EMT induced by the transforming growth factor β1 in mouse kidney proximal tubular epithelial C1.1 cells is accompanied by induction of autophagy and by the nuclear translocation of β-catenin [171].

Figure 8.

Degradation of E-cadherin and β-catenin involves endosomal lysosomal and autophagyc routes. (A) Normal conditions. When nutrients abound, E-cadherin and β-catenin are degraded through the endocytic-lysosomal and proteasomal routes, respectively. (B) Starvation. Under nutrient shortage, β-catenin switches to a selective macro autophagy for degradation. Wnt represents the WNT signaling cascade, TCF4 is the transcription factor 1.


6. The peculiar nonselective autophagy mechanisms of degradation of large portions of communicating junctions and desmosomes

6.1. Biology of communicating junctions and desmosomes

GAPJs are molecular ducts that communicate the cytoplasm of contiguous cells and allow the epithelium to respond coordinately to various stimuli or extracellular signals (Figure 1, yellow). These junctions are made up of tetraspan proteins: connexins in chordates and innexins in prechordates [172]. Six connexins polymerize to form a hemichannel or connexon in a cell, which attaches to a connexon in the neighboring cell, forming in this manner an intercellular channel that can be opened by diverse stimuli. The dense clustering of tens to thousands of intercellular channels originates a GAPJ [173175]. Connexins are associated with a scaffold of ZO-1 or ZO-3, vinculin, Src, and tubulin [176]. This association is important for the localization of connexons, the formation of the multimolecular clusters of intercellular channels in the plasma membrane, and the regulation of intercellular communication [177].

Des are cell-to-cell adhesion structures that confer mechanical strength to epithelia and cardiomyocytes. These junctions are composed of five main proteins: the desmosomal cadherins, desmogleins, and desmocollins are the receptors for adhesion. Their cytoplasmic tails bind to plaque proteins of the armadillo family, plakoglobin and plakophilin (Figure 1) [178]. The armadillo proteins attach to another plaque protein, desmoplakin, which, in turn, links the protein cluster to the cytoskeleton made of intermediary filaments of cytokeratin [179]. Observations in tissues and cultured cells have shown that Des can adopt a Ca2+-dependent adhesion state that progresses to a Ca2+-independent hyper-adhesion state, a process that requires PKC activation [22, 180182].

6.2. Macromolecular assemblies of communicating junctions and desmosomes are degraded by autophagy

GAPJs are extremely stable junctional structures: as soon as they are formed, they become indestructible [183, 184]. Nevertheless, they are very dynamic due to the fact that connexins have a very short half-life of only 1–5 h [185]. Consequently, there is a permanent turnover that involves the closure of the intercellular conduction by several stimuli, for example, the binding of EGF to its receptor. The central portion of the GAPJ is then internalized, including the bound hemichannels and membrane of the neighboring cell, forming a peculiar structure named annular GAPJ (Figure 9) [186]. In some conditions, annular GAPJ may be recycled back to the plasma membrane [187] although, usually, they are degraded through autophagy; yet, the precise mechanism, the kind of autophagy involved, and the fate of the cells depend on the trigger and/or the cellular context [185, 187189]. A mechanism that stops autophagy implicates the hijacking of components of the initiation of autophagy, for example, Atg16, by the connexins themselves. On nutrient starvation, connexins release Atg16, the blockade is lost and autophagy proceeds [190].

Figure 9.

Big portions of GAPJ and complete desmosomes (Des) are degraded by nonselective autophagy. The central portion of the GAPJs is internalized and degraded by autophagy. Dependent on the cell type and condition, complete Des are internalized and degraded by autophagy, and halves of Des are degraded by autophagy and in the proteasome.

On liver cells of BRL 3A expressing connexin-43, cadmium inhibits GAPJ intercellular communications and induces the degradation by autophagy of connexin-43 as well as apoptosis. Inhibition of autophagy exacerbates Cd2+-induced inhibition of the intercellular communication and apoptotic cell death [188] revealing the protective role that autophagy plays on cell fate.

Des are also very stable structures which can reach a hyper-adhesion state insensitive to Ca2+ depletion [191]. It has been shown that a half of Des is internalized after extracellular Ca2+ depletion in a PKC- and actin-dependent process [182, 192]. Internalized half desmosome is then transported by kinesins and microtubules toward the centrosome and remains there without recycle to the plasma membrane. Degradation proceeds in lysosomes and proteasomes [193]. In mouse epidermis, the complete Des are engulfed and internalized [192]. Nevertheless, the degradation mechanism is different when disassembly is triggered with autoantibodies from pemphigus vulgaris patients; in this case, Des disassemble in smaller complexes made of the autoantibody, desmoglein-3, and plakoglobin that are endocyted and delivered to the lysosomes through the endocytic route [194].


7. Continuous recycling in focal adhesion requires autophagy

7.1. Biology of focal adhesion and hemidesmosomes

FAs, also known as focal contacts, and HDes are the cellular junctions that attach cells to the extracellular matrix. HDes are common in stratified epithelia and bind epithelial cells to the underlying extracellular matrix (Figure 1, blue) [6] . The adhesion receptors of both, FAs and HDes, are transmembrane proteins of the family of integrins, which exist as heterodimers of α and β subunits form. There are 19 α-integrins and 8 β-integrins that combine to form 25 existing heterodimers in mammals [195, 196]. HDes provide stable adhesion and mechanical resistance to epithelial tissues by anchoring the extracellular matrix to the cytokeratin cytoskeleton, through a protein complex that includes the adhesion receptors α6β4 integrin, BP180, and the tetraspanin CD151, and the intracellular adapter proteins plectin and BP230 [6]. The expression of several HDes proteins depends on the transcription factor SOXF [197]. While the extracellular region of integrins of FAs binds the extracellular matrix, the cytosolic portion contacts specific plaque proteins such as focal adhesion kinase (FAK) and paxillin, which are important signaling proteins. Other protein components of the FA plaque, such as talin, vinculin, and α-actinin, bind the adhesion receptors to the actin microfilaments [5, 198, 199].

7.2. Role of lysosomes in the regulation of focal adhesions

FAs are essential in cell migration and, therefore, for embryogenesis, wound healing, immune cell function, cancer progression, and promoting metastasis [200]. Cell migration requires endocytosis and recycling of integrins given by endocytic signals in its cytoplasmic tail. These signals bind either clathrin or caveolin-1 to induce integrin endocytosis. Once inside the cell, integrins anchored to protein complexes are sent to the early endosomes, where they can be sorted either to late endosomes and lysosomes for degradation (Figure 10, 1) or to recycling endosomes and plasmatic membrane for the assembly of new FAs. A short loop for recycling requires Rab4 proteins and is generally activated in response to growth factors (Figure 10, 2); the long loop is Rab11 and Arf6 dependent and delivers integrins to the perinuclear recycling compartments (PNRCs) and, from there, to the cell membrane (Figure 10, 3) [190]. The actin cytoskeleton is essential to the recycling pathway; in fact, depletion of the actin-related protein (Arp) 2/3 or the nucleating-promoting factors such as the members of the Wiskott-Aldrich syndrome protein (WASP) blocks recycling and induces delivery to the lysosomes [5, 198].

FA disassembly is linked to autophagy in two ways: a nonselective autophagy triggered by extreme stress condition, such as starvation or hypoxia (Figure 10, 4), and a selective autophagy for housekeeping and quality control that includes ubiquitin-tagged substrate association of them with an autophagic cargo receptor (ACR) attached to LC3II. This autophagy provokes the disassemby of FA Under starvation, β1 integrin is degraded in autophagosomes in cervix adenocarcinoma epithelial HeLa cells. This autophagy is inhibited by high mTOR activity at the leading edge during migration, which promotes increased motility [201], whereas the activation of selective autophagy promotes FA disassembly in metastatic mammary epithelial cells (4T1) [202]. Thus, the inhibition of autophagosome reduces cancer cell’s malignancy, indicating that selective authophagy is also a cell migration regulator (Figure 10).

Figure 10.

Autophagy is crucial for the recycling of integrins in focal adhesions during migration. Cell migration requires continuous recycling of integrins. (1) Integrin endocytic pathway degradation. (2) Integrin short loop recycling; vesicular transport of integrins from the EE to the Rab4 containing RE, and from there back to the plasma membrane. (3) Integrin long loop recycling; vesicles transport integrins from EE to Rab11 containing RE, later on, to a perinuclear recycling compartment (PNRC) and then to the plasma membrane. (4) During starvation, integrins are endocyted and directed to the AP. (5) Cell migration leading edge. FA´s protein paxillin is recognized by autophagic cargo receptors (ACR) and degraded by selective macroautophagy, which induces FAs disassembly through a mTOR dependen pathway. In the leading edge, FAs must be first formed and then degraded to allow motility. Autophagy plays a crucial role in this process.

Besides migration, autophagy is linked to anoikis, a type of cell death due to detachment from the substrate. Loss of integrin-mediated adhesion initiates autophagy, which delays anoikis and downregulates apoptotic signals. This process affords cells time to reattach; however, in cancer cells, high autophagic activity after detachment provides resistance and promotes malignancy, allowing the cell to support stress condition, increase motility, and resist anoikis [203, 204].

Although there are several illnesses produced by the lack of HDes protein expression, little is known about HDes degradation.


8. Concluding remarks

Lysosomal degradation mechanisms are crucial for the formation, differentiation, and degradation of epithelial cell junctions. Epithelial cells use selective autophagy to degrade claudin-2, in response to the stimulation with the epidermal growth factor. Ouabain, at a concentration close to the hormonal, does not induce autophagy of tight junction proteins and, at high concentrations though, induces lysosomal degradation that can involve autophagy. The precise sequence of events and outcome of each lysosomal degradation mechanism is context dependent; nevertheless, it is clear that the degradation through macroautophagy of large plaque of complete communicating junctions and desmosomes, as well as of desmosomal halves, takes place either in natural tissues or in cultured cells. It is also clear that the desmosomal transition from weak to strong adhesion stages requires lysosomal activity, that β-catenin undergoes selective autophagy in some conditions and that E-cadherin degradation is performed in lysosomes through an endocytic route.


  1. 1. Cereijido M, Contreras RG, Shoshani L. Cell adhesion, polarity, and epithelia in the dawn of metazoans. Physiological Reviews. 2004;84:1229-1262. DOI: 10.1152/physrev.00001.2004
  2. 2. Cereijido M, Contreras RG, Shoshani L, García-Villegas MR. Membrane targeting. Progress in Biophysics & Molecular Biology. 2003;81:81-115
  3. 3. Cereijido M, Gonzalez-Mariscal L, Contreras RG. Epithelial tight junctions. American Review of Respiratory Disease. 1988;138:S17-S21. DOI: 10.1164/ajrccm/138.6_Pt_2.S17
  4. 4. Farquhar MG, Palade GE. Junctional complexes in various epithelia. Journal of Cell Biology. 1963;17:375-412
  5. 5. Burridge K, Chrzanowska-Wodnicka M. Focal adhesions, contractility, and signaling. Annual Review of Cell and Developmental Biology. 1996;12:463-518. DOI: 10.1146/annurev.cellbio.12.1.463
  6. 6. de Pereda JM, Ortega E, Alonso-García N, Gómez-Hernández M, Sonnenberg A. Advances and perspectives of the architecture of hemidesmosomes: Lessons from structural biology. Cell Adhesion & Migration. 2009;3:361-364
  7. 7. Cereijido M, Valdés J, Shoshani L, Contreras RG. Role of tight junctions in establishing and maintaining cell polarity. Annual Review of Physiology. 1998;60:161-177. DOI: 10.1146/annurev.physiol.60.1.161
  8. 8. Cereijido M, Contreras RG, González-Mariscal L. Development and alteration of polarity. Annual Review of Physiology. 1989;51:785-795. DOI: 10.1146/
  9. 9. Shoshani L, Contreras RG. Biogenesis of epithelial polarity and the tight junctions. In: Anderson JM, Cereijido M, editors. Tight Junctions. Boston, MA: Springer Science & Business Media. 2007. pp. 165-198
  10. 10. Yeaman C, Grindstaff KK, Nelson WJ. New perspectives on mechanisms involved in generating epithelial cell polarity. Physiological Reviews. 1999;79:73-98
  11. 11. Gumbiner B, Simons K. A functional assay for proteins involved in establishing an epithelial occluding barrier: Identification of a uvomorulin-like polypeptide. Journal of Cell Biology. 1986;102:457-468
  12. 12. Gumbiner B, Simons K. The role of uvomorulin in the formation of epithelial occluding junctions. Ciba Foundation Symposium. 1987;125:168-186
  13. 13. Poulsen SB, Fenton RA, Rieg T. Sodium-glucose cotransport. Current Opinion in Nephrology and Hypertension. 2015;24:463-469. DOI: 10.1097/MNH.0000000000000152
  14. 14. Chen L, Tuo B, Dong H. Regulation of intestinal glucose absorption by ion channels and transporters. Nutrients. 2016;8:43. DOI: 10.3390/nu8010043
  15. 15. Cereijido M, Ruiz O, Gonzalez-Mariscal L, Contreras RG, Balda MS, Garcia-Villegas R. The paracellular pathway: A small version of the kidney nephron. In: Audus KL, Raub TJ, editors. Biological Barriers to Protein Delivery. Boston, MA: Springer Science & Business Media. 1994. pp. 107-125. DOI: 10.1007/978-1-4615-2898-2
  16. 16. Price ER, Rott KH, Caviedes-Vidal E, Karasov WH. Claudin gene expression patterns do not associate with interspecific differences in paracellular nutrient absorption. Comparative Biochemistry and Physiology—Part B: Biochemistry & Molecular Biology. 2015;191:36-45. DOI: 10.1016/j.cbpb.2015.09.003
  17. 17. Simon DB, Lu Y, Choate KA, Velazquez H, Al-Sabban E, Praga M, et al. Paracellin-1, a renal tight junction protein required for paracellular Mg2+ resorption. Science. 1999;285:103-106
  18. 18. Koefoed-Johsen V, Ussing HH. The nature of the frog skin potential. Acta Physiologica Scandinavica. 1958;42:298-308. DOI: 10.1111/j.1748-1716.1958.tb01563.x.
  19. 19. Cheng CY, Mruk DD. Cell junction dynamics in the testis: Sertoli-germ cell interactions and male contraceptive development. Physiological Reviews. 2002;82:825-874. DOI: 10.1152/physrev.00009.2002
  20. 20. Brazil JC, Parkos CA. Pathobiology of neutrophil-epithelial interactions. Immunology Reviews. 2016;273(1):94-111. DOI: 10.1111/imr.12446
  21. 21. Thuenauer R, Müller SK, Römer W. Pathways of protein and lipid receptor-mediated transcytosis in drug delivery. Expert Opinion on Drug Delivery. 2016;14(3):1-11. DOI: 10.1080/17425247.2016.1220364
  22. 22. Broussard JA, Getsios S, Green KJ. Desmosome regulation and signaling in disease. Cell and Tissue Research. 2015;360:1-12. DOI: 10.1007/s00441-015-2136-5
  23. 23. Matter K, Balda MS. Signalling to and from tight junctions. Nature Reviews Molecular Cell Biology. 2003;4:225-237. DOI: 10.1038/nrm1055
  24. 24. Kim NG, Gumbiner BM. Adhesion to fibronectin regulates Hippo signaling via the FAK-Src-PI3K pathway. Journal of Cell Biology. 2015;210:503-515. DOI: 10.1083/jcb.201501025
  25. 25. Scarpa E, Mayor R. Collective cell migration in development. Journal of Cell Biology. 2016;212:143-155. DOI: 10.1083/jcb.201508047
  26. 26. El-Okdi N, Smaili S, Raju V, Shidyak A, Gupta S, Fedorova L, et al. Effects of cardiotonic steroids on dermal collagen synthesis and wound healing. Journal of Applied Physiology. 2008;105:30-36. DOI: 10.1152/japplphysiol.00119.2008
  27. 27. Clevers H. Wnt/beta-catenin signaling in development and disease. Cell. 2006;127:469-480. DOI: 10.1016/j.cell.2006.10.018
  28. 28. Huerta M, Muñoz R, Tapia R, Soto-Reyes E, Ramírez L, Recillas-Targa F, et al. Cyclin D1 is transcriptionally down-regulated by ZO-2 via an E box and the transcription factor c-Myc. Molecular Biology of the Cell. 2007;18:4826-4836. DOI: 10.1091/mbc.E07-02-0109
  29. 29. Betanzos A, Huerta M, Lopez-Bayghen E, Azuara E, Amerena J, Gonzalez-Mariscal L. The tight junction protein ZO-2 associates with Jun, Fos and C/EBP transcription factors in epithelial cells. Experimental Cell Research. 2004;292:51-66
  30. 30. Balda MS, Garrett MD, Matter K. The ZO-1-associated Y-box factor ZONAB regulates epithelial cell proliferation and cell density. Journal of Cell Biology. 2003;160:423-432. DOI: 10.1083/jcb.200210020
  31. 31. Takito J, Hikita C, Al-Awqati Q. Hensin, a new collecting duct protein involved in the in vitro plasticity of intercalated cell polarity. Journal of Clinical Investigation. 1996;98:2324-2331. DOI: 10.1172/JCI119044
  32. 32. Al-Awqati Q, Vijayakumar S, Takito J. Terminal differentiation of epithelia. Biological Chemistry. 2003;384:1255-1258. DOI: 10.1515/BC.2003.138
  33. 33. Barriere H, Chambard M, Mauchamp J, Gabrion J. Polarity reversal of inside-out thyroid follicles cultured within collagen gel: An ultrastructural study. Biology of the Cell. 1986;57:39-51
  34. 34. Barriere H, Chambard M, Selzner JP, Mauchamp J, Gabrion J. Polarity reversal of inside-out thyroid follicles cultured within collagen gel: Structure of the junctions assessed by freeze-fracture and lanthanum permeability. Biology of the Cell. 1988;62:133-144
  35. 35. Larue L, Ohsugi M, Hirchenhain J, Kemler R. E-cadherin null mutant embryos fail to form a trophectoderm epithelium. Proceedings of the National Academy of Sciences. 1994;91:8263-8267
  36. 36. Xu J, Kausalya PJ, Phua DCY, Ali SM, Hossain Z, Hunziker W. Early embryonic lethality of mice lacking ZO-2, but Not ZO-3, reveals critical and nonredundant roles for individual zonula occludens proteins in mammalian development. Molecular and Cellular Biology. 2008;28:1669-1678. DOI: 10.1128/MCB.00891-07
  37. 37. Konrad M, Schaller A, Seelow D, Pandey AV, Waldegger S, Lesslauer A, et al. Mutations in the tight-junction gene claudin 19 (CLDN19) are associated with renal magnesium wasting, renal failure, and severe ocular involvement. The American Journal of Human Genetics. 2006;79:949-957. DOI: 10.1086/508617
  38. 38. Sambrotta M, Strautnieks S, Papouli E, Rushton P, Clark BE, Parry DA, et al. Mutations in TJP2 cause progressive cholestatic liver disease. Nature Genetics. 2014;46:326-328. DOI: 10.1038/ng.2918
  39. 39. Samuelov L, Sprecher E. Inherited desmosomal disorders. Cell and Tissue Research. 2014;360(3):1-19. DOI: 10.1007/s00441-014-2062-y
  40. 40. Koster J, Kuikman I, Kreft M, Sonnenberg A. Two different mutations in the cytoplasmic domain of the integrin beta 4 subunit in nonlethal forms of epidermolysis bullosa prevent interaction of beta 4 with plectin. Journal of Investigative Dermatology. 2001;117:1405-1411. DOI: 10.1046/j.0022-202x.2001.01567.x
  41. 41. Nava P, López S, Arias CF, Islas S, Gonzalez-Mariscal L. The rotavirus surface protein VP8 modulates the gate and fence function of tight junctions in epithelial cells. Journal of Cell Science. 2004;117:5509-5519. DOI: 10.1242/jcs.01425
  42. 42. Bonilla-Delgado J, Bulut G, Liu X, Cortés-Malagón EM, Schlegel R, Flores-Maldonado C, et al. The E6 oncoprotein from HPV16 enhances the canonical Wnt/β-catenin pathway in skin epidermis in vivo. Molecular Cancer Research. 2012;10:250-258. DOI: 10.1158/1541-7786.MCR-11-0287
  43. 43. de Duve C. Lysosomes. Bulletin De l'Academie Royale De Medecine De Belgique. 1958;23:608-618
  44. 44. Lim CY, Zoncu R. The lysosome as a command-and-control center for cellular metabolism. Journal of Cell Biology. 2016;214:653-664. DOI: 10.1083/jcb.201607005
  45. 45. Forgac M. Vacuolar ATPases: Rotary proton pumps in physiology and pathophysiology. Nature Reviews Molecular Cell Biology. 2007;8:917-929. DOI: 10.1038/nrm2272
  46. 46. Li Y, Chen B, Zou W, Wang X, Wu Y, Zhao D, et al. The lysosomal membrane protein SCAV-3 maintains lysosome integrity and adult longevity. Journal of Cell Biology. 2016;215:167-185. DOI: 10.1083/jcb.201602090
  47. 47. Schreij AMA, Fon EA, McPherson PS. Endocytic membrane trafficking and neurodegenerative disease. Cellular and Molecular Life Sciences. 2016;73:1529-1545. DOI: 10.1007/s00018-015-2105-x
  48. 48. Scharaw S, Iskar M, Ori A, Boncompain G, Laketa V, Poser I, et al. The endosomal transcriptional regulator RNF11 integrates degradation and transport of EGFR. Journal of Cell Biology. 2016;261:543-558. DOI: 10.1083/jcb.201601090
  49. 49. Marzella L, Ahlberg J, Glaumann H. Autophagy, heterophagy, microautophagy and crinophagy as the means for intracellular degradation. Virchows Arch B Cell Pathol Incl Mol Pathol. 1981;36(2-3):219-34
  50. 50. Orenstein SJ, Cuervo AM. Chaperone-mediated autophagy: Molecular mechanisms and physiological relevance. Seminars in Cell and Developmental Biology. 2010;21:719-726. DOI: 10.1016/j.semcdb.2010.02.005
  51. 51. Mizushima N, Komatsu M. Autophagy: Renovation of cells and tissues. Cell. 2011;147:728-741. DOI: 10.1016/j.cell.2011.10.026
  52. 52. Rabinowitz JD, White E. Autophagy and metabolism. Science. 2010;330:1344-1348. DOI: 10.1126/science.1193497
  53. 53. Singh R, Cuervo AM. Autophagy in the cellular energetic balance. Cell Metabolism. 2011;13:495-504. DOI: 10.1016/j.cmet.2011.04.004
  54. 54. Schmidt MR, Haucke V. Recycling endosomes in neuronal membrane traffic. Biology of the Cell. 2007;99:333-342. DOI: 10.1042/BC20070007
  55. 55. Bucci C, Parton RG, Mather IH, Stunnenberg H, Simons K, Hoflack B, et al. The small GTPase rab5 functions as a regulatory factor in the early endocytic pathway. Cell. 1992;70:715-728
  56. 56. Morrison HA, Dionne H, Rusten TE, Brech A, Fisher WW, Pfeiffer BD, et al. Regulation of early endosomal entry by the Drosophila tumor suppressors Rabenosyn and Vps45. Molecular Biology of the Cell. 2008;19:4167-4176. DOI: 10.1091/mbc.E08-07-0716
  57. 57. Sönnichsen B, De Renzis S, Nielsen E, Rietdorf J, Zerial M. Distinct membrane domains on endosomes in the recycling pathway visualized by multicolor imaging of Rab4. Rab5, and Rab11. Journal of Cell Biology. 2000;149:901-914
  58. 58. Bucci C, Thomsen P, Nicoziani P, McCarthy J, van Deurs B. Rab7: A key to lysosome biogenesis. Molecular Biology of the Cell. 2000;11:467-480
  59. 59. Yoshimori T, Yamamoto A, Moriyama Y, Futai M, Tashiro Y. Bafilomycin A1, a specific inhibitor of vacuolar-type H(+)-ATPase, inhibits acidification and protein degradation in lysosomes of cultured cells. Journal of Biological Chemistry. 1991;266:17707-17712
  60. 60. Sahu R, Kaushik S, Clement CC, Cannizzo ES, Scharf B, Follenzi A, et al. Microautophagy of cytosolic proteins by late endosomes. Developmental Cell. 2011;20:131-139. DOI: 10.1016/j.devcel.2010.12.003
  61. 61. Hua R, Cheng D, Coyaud É, Freeman S, Di Pietro E, Wang Y, et al. VAPs and ACBD5 tether peroxisomes to the ER for peroxisome maintenance and lipid homeostasis. Journal of Cell Biology. 2017;16(2):367-377. DOI: 10.1083/jcb.201608128
  62. 62. Costello JL, Castro IG, Hacker C, Schrader TA, Metz J, Zeuschner D, et al. ACBD5 and VAPB mediate membrane associations between peroxisomes and the ER. Journal of Cell Biology. 2017;931:331-342. DOI: 10.1083/jcb.201607055
  63. 63. Gómez-Sintes R, Ledesma MD, Boya P. Lysosomal cell death mechanisms in aging. Ageing Research Reviews. 2016;32:150-168. DOI: 10.1016/j.arr.2016.02.009
  64. 64. Toops KA, Lakkaraju A. Let's play a game of chutes and ladders: Lysosome fusion with the epithelial plasma membrane. Communicative & Integrative Biology. 2013;6:e24474. DOI: 10.4161/cib.24474
  65. 65. Claude P, Goodenough DA. Goodenough. Fracture faces of zonulae occludentes from “tight” and “leaky” epithelia. Journal of Cell Biology. 1973;58:390-400
  66. 66. Diamond JM. Twenty-first Bowditch lecture. The epithelial junction: Bridge, gate, and fence. Physiologist. 1977;20:10-18
  67. 67. Mandel LJ, Bacallao R, Zampighi G. Uncoupling of the molecular ‘fence’ and paracellular “gate” functions in epithelial tight junctions. Nature. 1993;361:552-555. DOI: 10.1038/361552a0
  68. 68. Dragsten PR, Blumenthal R, Handler JS. Membrane asymmetry in epithelia: Is the tight junction a barrier to diffusion in the plasma membrane? Nature. 1981;294:718-722
  69. 69. van Meer G, Simons K. The function of tight junctions in maintaining differences in lipid composition between the apical and the basolateral cell surface domains of MDCK cells. The EMBO Journal. 1986;5:1455-1464
  70. 70. Cereijido M, Contreras RG, Shoshani L, Flores-Benitez D, Larre I. Tight junction and polarity interaction in the transporting epithelial phenotype. Biochimica et Biophysica Acta. 2008;1778:770-793. DOI: 10.1016/j.bbamem.2007.09.001
  71. 71. Laffafian I, Hallett MB. Lipid-protein cargo transfer: A mode of direct cell-to-cell communication for lipids and their associated proteins. Journal of Cellular Physiology. 2007;210:336-342. DOI: 10.1002/jcp.20851
  72. 72. Sneyd J, Wetton BT, Charles AC, Sanderson MJ. Intercellular calcium waves mediated by diffusion of inositol trisphosphate: A two-dimensional model. American Journal of Physiology. 1995;268:C1537-C1545
  73. 73. Turin L, Béhé P, Plonsky I, Dunina-Barkovskaya A. Hydrophobic ion transfer between membranes of adjacent hepatocytes: A possible probe of tight junction structure. Proceedings of the National Academy of Sciences. 1991;88:9365-9369
  74. 74. Grebenkämper K, Galla HJ. Translational diffusion measurements of a fluorescent phospholipid between MDCK-I cells support the lipid model of the tight junctions. Chemistry and Physics of Lipids. 1994;71:133-143
  75. 75. Furuse M, Fujita K, Hiiragi T, Fujimoto K, Tsukita S. Claudin-1 and -2: Novel integral membrane proteins localizing at tight junctions with no sequence similarity to occludin. Journal of Cell Biology. 1998;141:1539-1550
  76. 76. Haseloff RF, Dithmer S, Winkler L, Wolburg H, Blasig IE. Transmembrane proteins of the tight junctions at the blood-brain barrier: Structural and functional aspects. Seminars in Cell and Developmental Biology. 2015;38:16-25. DOI: 10.1016/j.semcdb.2014.11.004
  77. 77. Cereijido M, Contreras RG, Flores-Benítez D, Flores-Maldonado C, Larre I, Ruiz A, et al. New diseases derived or associated with the tight junction. Archives of Medical Research. 2007;38:465-478. DOI: 10.1016/j.arcmed.2007.02.003
  78. 78. González-Mariscal L, Betanzos A, Nava P, Jaramillo BE. Tight junction proteins. Progress in Biophysics & Molecular Biology. 2003;81:1-44
  79. 79. Tsukita S, Furuse M, Itoh M. Multifunctional strands in tight junctions. Nature Reviews Molecular Cell Biology. 2001;2:285-293. DOI: 10.1038/35067088
  80. 80. Anderson JM. Molecular structure of tight junctions and their role in epithelial transport. Physiology. 2001;16:126-130. DOI: 10.1007/BF01870332
  81. 81. Cereijido M, Anderson JM. Tight Junctions. CRC Press; 2001. USA
  82. 82. Gonzalez-Mariscal L. Tight Junctions. Boston, MA: Springer Science & Business Media; 2007. DOI: 10.1007/0-387-36673-3
  83. 83. Weber CR, Liang GH, Wang Y, Das S, Shen L, Yu ASL, et al. Claudin-2-dependent paracellular channels are dynamically gated. eLife Sciences. 2016;4:e09906. DOI: 10.7554/eLife.09906
  84. 84. Kottra G, Frömter E. Functional properties of the paracellular pathway in some leaky epithelia. Journal of Experimental Biology. 1983;106:217-229
  85. 85. Günzel D, Yu ASL. Claudins and the modulation of tight junction permeability. Physiological Reviews. 2013;93:525-569. DOI: 10.1152/physrev.00019.2012
  86. 86. Anderson JM, Stevenson BR, Jesaitis LA, Goodenough DA, Mooseker MS. Characterization of ZO-1, a protein component of the tight junction from mouse liver and Madin-Darby canine kidney cells. Journal of Cell Biology. 1988;106:1141-1149
  87. 87. Gonzalez-Mariscal L, Betanzos A, Avila-Flores A. MAGUK proteins: Structure and role in the tight junction. Seminars in Cell and Developmental Biology. 2000;11:315-324. DOI: 10.1006/scdb.2000.0178
  88. 88. Gonzalez-Mariscal L. Molecular Characterization of the Tight Junction Protein ZO-1 in MDCK Cells*1, *2. Experimental Cell Research. 1999;248:97-109. DOI: 10.1006/excr.1999.4392.
  89. 89. Fanning AS, Ma TY, Anderson JM. Isolation and functional characterization of the actin binding region in the tight junction protein ZO-1. FASEB Journal. 2002;16:1835-1837. DOI: 10.1096/fj.02-0121fje
  90. 90. Molecular organization and function of invertebrate occluding junctions. Seminaries in Cell & Developmental Biology. 2014;36:186-193. DOI: 10.1016/j.semcdb.2014.09.009
  91. 91. Suzuki H, Ito Y, Yamazaki Y, Mineta K, Uji M, Abe K, et al. The four-transmembrane protein IP39 of Euglena forms strands by a trimeric unit repeat. Nature Communications. 2013;4:1766. DOI: 10.1038/ncomms2731
  92. 92. Cereijido M, Robbins ES, Dolan WJ, Rotunno CA, Sabatini DD. Polarized monolayers formed by epithelial cells on a permeable and translucent support. Journal of Cell Biology. 1978;77:853-880
  93. 93. Boulpaep EL, Seely JF. Electrophysiology of proximal and distal tubules in the autoperfused dog kidney. American Journal of Physiology. 1971;221:1084-1096
  94. 94. Lutz MD, Cardinal JE, Burg MB. Electrical resistance of renal proximal tubule perfused in vitro. American Journal of Physiology. 1973;225:729-734
  95. 95. Helman SI, Grantham JJ, Burg MB. Effect of vasopressin on electrical resistance of renal cortical collecting tubules. American Journal of Physiology. 1971;220:1825-1832
  96. 96. Rau WS, Frömter E. Electrical properties of the medullary collecting ducts of the golden hamster kidney. II. The transepithelial resistance. Pflügers Archiv. 1974;351:113-131
  97. 97. Lavelle JP, Meyers SA, Ruiz WG, Buffington CT, Zeidel ML, Apodaca G. Urothelial pathophysiological changes in feline interstitial cystitis: A human model. American Journal of Physiology. Renal Physiology. 2000;278:F540-F553
  98. 98. Lewis SA, Eaton DC, Clausen CH, Diamond JM. Nystatin as a probe for investigating the electrical properties of a tight epithelium. The Journal of General Physiology. 1977;70:427-440
  99. 99. Furuse M, Furuse K, Sasaki H, Tsukita S. Conversion of zonulae occludentes from tight to leaky strand type by introducing claudin-2 into Madin-Darby canine kidney I cells. Journal of Cell Biology. 2001;153:263-272
  100. 100. Amasheh S, Meiri N, Gitter AH, Schöneberg T, Mankertz J, Schulzke JD, et al. Claudin-2 expression induces cation-selective channels in tight junctions of epithelial cells. Journal of Cell Science. 2002;115:4969-4976
  101. 101. Rosenthal R, Milatz S, Krug SM, Oelrich B, Schulzke JD, Amasheh S, et al., Claudin-2, a component of the tight junction, forms a paracellular water channel. Journal of Cell Science. 2010;123:1913-1921. DOI: 10.1242/jcs.060665
  102. 102. Gonzalez-Mariscal L, Del CarmenNamorado M, Martin D, Sierra G, Reyes JL. The tight junction proteins claudin-7 and -8 display a different subcellular localization at Henle's loops and collecting ducts of rabbit kidney. Nephrology, Dialysis, Transplantation. 2006;21:2391-2398. DOI: 10.1093/ndt/gfl255
  103. 103. Kiuchi-Saishin Y, Gotoh S, Furuse M, Takasuga A, Tano Y, Tsukita S. Differential expression patterns of claudins, tight junction membrane proteins, in mouse nephron segments. Journal of the American Society of Nephrology. 2002;13:875-886
  104. 104. Reyes JL, Lamas M, Martin D, Namorado MD, Islas S, Luna J, et al. The renal segmental distribution of claudins changes with development. Kidney International. 2002;62:476-487. DOI: 10.1046/j.1523-1755.2002.00479.x
  105. 105. Muto S, Hata M, Taniguchi J, Tsuruoka S, Moriwaki K, Saitou M, et al., Claudin-2-deficient mice are defective in the leaky and cation-selective paracellular permeability properties of renal proximal tubules. Proceedings of the National academy of Sciences of the United States of America. 2010;107:8011-8016. DOI: 10.1073/pnas.0912901107
  106. 106. Colegio OR, Van Itallie C, Rahner C, Anderson JM. Claudin extracellular domains determine paracellular charge selectivity and resistance but not tight junction fibril architecture. American Journal of Physiology. Cell Physiology. 2003;284:C1346-C1354. DOI: 10.1152/ajpcell.00547.2002
  107. 107. Colegio OR, Van Itallie CM, McCrea HJ, Rahner C, Anderson JM. Claudins create charge-selective channels in the paracellular pathway between epithelial cells. American Journal of Physiology. Cell Physiology. 2002;283:C142-C147. DOI: 10.1152/ajpcell.00038.2002
  108. 108. Morita K, Furuse M, Fujimoto K, Tsukita S. Claudin multigene family encoding four-transmembrane domain protein components of tight junction strands. Proceedings of the National Academy of Sciences. 1999;96:511-516
  109. 109. Gallardo JM, Hernández JM, Contreras RG, Flores-Maldonado C, González-Mariscal L, Cereijido M. Tight junctions are sensitive to peptides eliminated in the urine. Journal of Membrane Biology. 2002;188:33-42. DOI: 10.1007/s00232-001-0170-6
  110. 110. Flores-Benitez D, Ruiz-Cabrera A, Flores-Maldonado C, Shoshani L, Cereijido M, Contreras RG. Control of tight junctional sealing: Role of epidermal growth factor. American Journal of Physiology. Renal Physiology. 2007;292:F828-F836. DOI: 10.1152/ajprenal.00369.2006
  111. 111. Mullin JM, Laughlin KV, Ginanni N, Marano CW, Clarke HM, Peralta Soler A. Increased tight junction permeability can result from protein kinase C activation/translocation and act as a tumor promotional event in epithelial cancers. Annals of the New York Academy of Sciences. 2000;915:231-236
  112. 112. Singh AB, Harris RC. Epidermal growth factor receptor activation differentially regulates claudin expression and enhances transepithelial resistance in Madin-Darby canine kidney cells. Journal of Biological Chemistry. 2004;279:3543-3552. DOI: 10.1074/jbc.M308682200
  113. 113. Ikari A, Takiguchi A, Atomi K, Sugatani J. Epidermal growth factor increases clathrin-dependent endocytosis and degradation of claudin-2 protein in MDCK II cells. Journal of Cellular Physiology. 2011;226:2448-2456. DOI: 10.1002/jcp.22590
  114. 114. Garcia-Hernandez V, Flores-Maldonado C, Rincon-Heredia R, Verdejo-Torres O, Bonilla-Delgado J, Meneses-Morales I, et al. EGF regulates claudin-2 and -4 expression through Src and STAT3 in MDCK cells. Journal of Cellular Physiology. 2015;230:105-115. DOI: 10.1002/jcp.24687
  115. 115. Singh AB, Dhawan P. Claudins and cancer: Fall of the soldiers entrusted to protect the gate and keep the barrier intact. Seminars in Cell and Developmental Biology. 2015;42:58-65. DOI: 10.1016/j.semcdb.2015.05.001
  116. 116. Ikari A, Atomi K, Takiguchi A, Yamazaki Y, Miwa M, Sugatani J. Epidermal growth factor increases claudin-4 expression mediated by Sp1 elevation in MDCK cells. Biochemical and Biophysical Research Communications. 2009;384:306-310. DOI: 10.1016/j.bbrc.2009.04.120
  117. 117. Ikari A, Atomi K, Takiguchi A, Yamazaki Y, Hayashi H, Hirakawa J, et al. Enhancement of cell-cell contact by claudin-4 in renal epithelial Madin-Darby canine kidney cells. Journal of Cellular Biochemistry. 2012;113:499-507. DOI: 10.1002/jcb.23373
  118. 118. Flores-Benitez D, Rincon-Heredia R, Razgado LF, Larre I, Cereijido M, Contreras RG. Control of tight junctional sealing: Roles of epidermal growth factor and prostaglandin E2. American Journal of Physiology. Cell Physiology. 2009;297:C611-C620. DOI: 10.1152/ajpcell.00622.2008
  119. 119. Lu R, Johnson DL, Stewart L, Waite K, Elliott D, Wilson JM. Rab14 regulation of claudin-2 trafficking modulates epithelial permeability and lumen morphogenesis. Molecular Biology of the Cell. 2014;25:1744-1754. DOI: 10.1091/mbc.E13-12-0724
  120. 120. Nighot PK, Hu CA, Ma TY. Autophagy enhances intestinal epithelial tight junction barrier function by targeting claudin-2 protein degradation. Journal of Biological Chemistry. 2015;290:7234-7246. DOI: 10.1074/jbc.M114.597492
  121. 121. Yang Y, Li W, Sun Y, Han F, Hu CA, Wu Z. Amino acid deprivation disrupts barrier function and induces protective autophagy in intestinal porcine epithelial cells. Amino Acids. 2015;47:2177-2184. DOI: 10.1007/s00726-014-1844-6
  122. 122. Zhou Y, Zheng B, Ye L, Zhang H, Zhu S, Zheng X, et al. Retinoic acid prevents disruption of blood-spinal cord barrier by inducing autophagic flux after spinal cord injury. Neurochemical Research. 2016;41:813-825. DOI: 10.1007/s11064-015-1756-1
  123. 123. Withering W. An Account of the Foxglove, and Some of Its Medical Uses, 1st ed. Printed by N. Swinney for GGJ and J. Robinson, Paternoster-Row. London; 1785
  124. 124. Schatzmann HJ. Herzglykoside als Hemmstoffe für den aktiven Kalium- und Natriumtransport durch die Erythrocytenmembran. Helvetica Physiologica et Pharmacologica Acta. 1953;11:346-354
  125. 125. Bauer N, Müller-Ehmsen J, Krämer U, Hambarchian N, Zobel C, Schwinger RH, Neu H, Kirch U, Grünbaum EG, Schoner W. Ouabain-like compound changes rapidly on physical exercise in humans and dogs: Effects of beta-blockade and angiotensin-converting enzyme inhibition. Hypertension. 2005;45:1024-1028. DOI: 10.1161/01.HYP.0000165024.47728.f7
  126. 126. Schneider R, Wray V, Nimtz M, Lehmann WD, Kirch U, Antolovic R, et al. Bovine adrenals contain, in addition to ouabain, a second inhibitor of the sodium pump. Journal of Biological Chemistry. 1998;273:784-792
  127. 127. Schoner W, Bauer N, Müller‐Ehmsen J, KrÄmer U, Hambarchian N, Schwinger R, et al. Ouabain as a mammalian hormone. Annals of the New York Academy of Sciences. 2003;986:678-684
  128. 128. Schoner W, Scheiner-Bobis G. Endogenous cardiac glycosides: Hormones using the sodium pump as signal transducer. Seminars in Nephrology. 2005;25:343-351. DOI: 10.1016/j.semnephrol.2005.03.010
  129. 129. Moreth K, Kuske R, Renner D, Schoner W. Blood pressure in essential hypertension correlates with the concentration of a circulating inhibitor of the sodium pump. Klinische Wochenschrift. 1986;64:239-244. DOI: 10.1007/BF01711656
  130. 130. Fedorova OV, Shapiro JI, Bagrov AY. Endogenous cardiotonic steroids and salt-sensitive hypertension. Biochimica Et Biophysica Acta (BBA)—Molecular Basis of Disease. 2010;1802:1230-1236. DOI: 10.1016/j.bbadis.2010.03.011
  131. 131. Nesher M, Shpolansky U, Viola N, Dvela M, Buzaglo N, Ben‐Ami HC, et al. Ouabain attenuates cardiotoxicity induced by other cardiac steroids. British Journal of Pharmacology. 2010;160:346-354. DOI: 10.1111/j.1476-5381.2010.00701.x
  132. 132. Koltsova SV, Trushina Y, Haloui M, Akimova OA, Tremblay J, Hamet P, et al. Ubiquitous [Na+]i/[K+]i-sensitive transcriptome in mammalian cells: Evidence for Ca(2+)i-independent excitation-transcription coupling. PLoS One. 2012;7:e38032. DOI: 10.1371/journal.pone.0038032
  133. 133. Wang H, Haas M, Liang M, Cai T, Tian J, Li S, et al. Ouabain assembles signaling cascades through the caveolar Na+/K+-ATPase. Journal of Biological Chemistry. 2004;279:17250-17259. DOI: 10.1074/jbc.M313239200
  134. 134. Aizman O, Uhlén P, Lal M, Brismar H, Aperia A. Ouabain, a steroid hormone that signals with slow calcium oscillations. Proceedings of the National Academy of Sciences. 2001;98:13420-13424. DOI: 10.1073/pnas.221315298
  135. 135. Zhang S, Malmersjö S, Li J, Ando H, Aizman O, Uhlén P, et al. Distinct role of the N-terminal tail of the Na,K-ATPase catalytic subunit as a signal transducer. Journal of Biological Chemistry. 2006;281:21954-21962. DOI: 10.1074/jbc.M601578200
  136. 136. Akimova OA, Hamet P, Orlov SN. [Na+]i/[K+]i-independent death of ouabain-treated renal epithelial cells is not mediated by Na+,K+-ATPase internalization and de novo gene expression. Pflügers Archiv. 2008;455:711-719. DOI: 10.1007/s00424-007-0283-6
  137. 137. Contreras RG, Flores-Maldonado C, Lazaro A, Shoshani L, Flores-Benitez D, Larre I, et al. Ouabain binding to Na+,K+-ATPase relaxes cell attachment and sends a specific signal (NACos) to the nucleus. Journal of Membrane Biology. 2004;198:147-158. DOI: 10.1007/s00232-004-0670-2
  138. 138. Liang M, Cai T, Tian J, Qu W, Xie ZJ. Functional characterization of Src-interacting Na/K-ATPase using RNA interference assay. Journal of Biological Chemistry. 2006;281:19709-19719. DOI: 10.1074/jbc.M512240200
  139. 139. Larré I, Lazaro A, Contreras RG, Balda MS, Matter K, Flores-Maldonado C, et al. Ouabain modulates epithelial cell tight junction. Proceedings of the National academy of Sciences of the United States of America. 2010;107:11387-11392. DOI: 10.1073/pnas.1000500107
  140. 140. Contreras RG, Shoshani L, Flores-Maldonado C, Lázaro A, Cereijido M. Relationship between Na(+),K(+)-ATPase and cell attachment. Journal of Cell Science. 1999;112(Pt 23):4223-4232
  141. 141. Rincon-Heredia R, Flores-Benitez D, Flores-Maldonado C, Bonilla-Delgado J, García-Hernández V, Verdejo-Torres O, et al. Ouabain induces endocytosis and degradation of tight junction proteins through ERK1/2-dependent pathways. Experimental Cell Research. 2014;320:108-118. DOI: 10.1016/j.yexcr.2013.10.008
  142. 142. Akimova OA, Tverskoi AM, Smolyaninova LV, Mongin AA, Lopina OD, La J, et al. Critical role of the α1-Na+, K+-ATPase subunit in insensitivity of rodent cells to cytotoxic action of ouabain. Apoptosis. 2015;20:1200-1210. DOI: 10.1007/s10495-015-1144-y
  143. 143. Akimova OA, Tremblay J, Van Huysse JW, Hamet P, Orlov SN. Cardiotonic steroid-resistant alpha1-Na+,K+-ATPase rescues renal epithelial cells from the cytotoxic action of ouabain: Evidence for a Nai+,Ki+-independent mechanism. Apoptosis. 2010;15:55-62. DOI: 10.1007/s10495-009-0429-4
  144. 144. Contreras RG, Flores-Beni TD, Flores-Maldonado C, Larre I, Shoshani L, Cereijido M. Na+,K+-ATPase and hormone ouabain:new roles for an old enzyme and an old inhibitor. Cellular and Molecular Biology (Noisy-le-Grand, France). 2006;52:31-40
  145. 145. Hou J, Rajagopal M, Yu ASL. Claudins and the kidney. Annual Review of Physiology. 2013;75:479-501. DOI: 10.1146/annurev-physiol-030212-183705
  146. 146. Arteaga ME, Hunziker W, Teo AS, Hillmer AM, Mutchinick OM. Familial hypomagnesemia with hypercalciuria and nephrocalcinosis: Variable phenotypic expression in three affected sisters from Mexican ancestry. Renal Failure. 2015;37:180-183. DOI: 10.3109/0886022X.2014.977141
  147. 147. Sharma S, Place E, Lord K, Leroy BP, Falk MJ, Pradhan M. Claudin 19-based familial hypomagnesemia with hypercalciuria and nephrocalcinosis in a sibling pair. Clinical Nephrology. 2016;85:346-352. DOI: 10.5414/CN108783
  148. 148. Hou J, Renigunta A, Konrad M, Gomes AS, Schneeberger EE, Paul DL, et al. Claudin-16 and claudin-19 interact and form a cation-selective tight junction complex. Journal of Clinical Investigation. 2008;118:619-628. DOI: 10.1172/JCI33970
  149. 149. Hou J, Paul DL, Goodenough DA. Paracellin-1 and the modulation of ion selectivity of tight junctions. Journal of Cell Science. 2005;118:5109-5118. DOI: 10.1242/jcs.02631
  150. 150. Gong Y, Renigunta V, Zhou Y, Sunq A, Wang J, Yang J, et al. Biochemical and biophysical analyses of tight junction permeability made of claudin-16 and claudin-19 dimerization. Molecular Biology of the Cell. 2015;26(24):4333-4346. DOI: 10.1091/mbc.E15-06-0422
  151. 151. Kausalya PJ, Amasheh S, Günzel D, Wurps H, Müller D, Fromm M, et al. Disease-associated mutations affect intracellular traffic and paracellular Mg2+ transport function of Claudin-16. Journal of Clinical Investigation. 2006;116:878-891. DOI: 10.1172/JCI26323
  152. 152. Müller D, Kausalya PJ, Meij IC, Hunziker W. Familial hypomagnesemia with hypercalciuria and nephrocalcinosis: Blocking endocytosis restores surface expression of a novel Claudin-16 mutant that lacks the entire C-terminal cytosolic tail. Human Molecular Genetics. 2006;15:1049-1058. DOI: 10.1093/hmg/ddl020
  153. 153. Müller D, Kausalya PJ, Claverie-Martin F, Meij IC, Eggert P, Garcia-Nieto V, et al. A novel claudin 16 mutation associated with childhood hypercalciuria abolishes binding to ZO-1 and results in lysosomal mistargeting. The American Journal of Human Genetics. 2003;73:1293-1301. DOI: 10.1086/380418
  154. 154. Ikari A, Matsumoto S, Harada H, Takagi K, Degawa M, Takahashi T, et al. Dysfunction of paracellin-1 by dephosphorylation in Dahl salt-sensitive hypertensive rats. Journal of Physiological Sciences. 2006;56:379-383. DOI: 10.2170/physiolsci.SC008906
  155. 155. Cong X, Zhang Y, Li J, Mei M, Ding C, Xiang RL, et al. Claudin-4 is required for modulation of paracellular permeability by muscarinic acetylcholine receptor in epithelial cells. Journal of Cell Science. 2015;128:2271-2286. DOI: 10.1242/jcs.165878
  156. 156. Mandai K, Nakanishi H, Satoh A, Obaishi H, Wada M, Nishioka H, et al. Afadin: A novel actin filament-binding protein with one PDZ domain localized at cadherin-based cell-to-cell adherens junction. Journal of Cell Biology. 1997;139:517-528
  157. 157. Sakisaka T, Ikeda W, Ogita H, Fujita N, Takai Y. The roles of nectins in cell adhesions: Cooperation with other cell adhesion molecules and growth factor receptors. Current Opinion in Cell Biology. 2007;19:593-602. DOI: 10.1016/
  158. 158. Kemler R, Ozawa M. Uvomorulin-catenin complex: Cytoplasmic anchorage of a Ca2+-dependent cell adhesion molecule. Bioessays. 1989;11:88-91. DOI: 10.1002/bies.950110403
  159. 159. Hirano S, Takeichi M. Cadherins in brain morphogenesis and wiring. Physiological Reviews. 2012;92:597-634. DOI: 10.1152/physrev.00014.2011
  160. 160. Nose A, Nagafuchi A, Takeichi M. Expressed recombinant cadherins mediate cell sorting in model systems. Cell. 1988;54:993-1001
  161. 161. Takeichi M. Cadherin cell adhesion receptors as a morphogenetic regulator. Science. 1991;251:1451-1455
  162. 162. Pokutta S, Weis WI. Structure and mechanism of cadherins and catenins in cell-cell contacts. Annual Review of Cell and Developmental Biology. 2007;23:237-261. DOI: 10.1146/annurev.cellbio.22.010305.104241
  163. 163. Niessen CM, Gottardi CJ. Molecular components of the adherens junction. Biochimica et Biophysica Acta. 2008;1778:562-571. DOI: 10.1016/j.bbamem.2007.12.015
  164. 164. Gumbiner BM. Regulation of cadherin-mediated adhesion in morphogenesis. Nature Reviews Molecular Cell Biology. 2005;6:622-634. DOI: 10.1038/nrm1699
  165. 165. Abedin M, King N. The premetazoan ancestry of cadherins. Science. 2008;319:946-948. DOI: 10.1126/science.1151084
  166. 166. Nanes BA, Chiasson-MacKenzie C, Lowery AM, Ishiyama N, Faundez V, Ikura M, et al. p120-catenin binding masks an endocytic signal conserved in classical cadherins. Journal of Cell Biology. 2012;199:365-380. DOI: 10.1083/jcb.201205029
  167. 167. Anastasiadis PZ, Reynolds AB. The p120 catenin family: Complex roles in adhesion, signaling and cancer. Journal of Cell Science. 2000;113(Pt 8):1319-1334.
  168. 168. Fujita Y, Krause G, Scheffner M, Zechner D, Leddy HEM, Behrens J, et al. Hakai, a c-Cbl-like protein, ubiquitinates and induces endocytosis of the E-cadherin complex. Nature Cell Biology. 2002;4:222-231. DOI: 10.1038/ncb758
  169. 169. Petherick KJ, Williams AC, Lane JD, Ordóñez-Morán P, Huelsken J, Collard TJ, et al. Autolysosomal β-catenin degradation regulates Wnt-autophagy-p62 crosstalk. The EMBO Journal. 2013;32:1903-1916. DOI: 10.1038/emboj.2013.123
  170. 170. Gugnoni M, Sancisi V, Gandolfi G, Manzotti G, Ragazzi M, Giordano D, et al. Cadherin-6 promotes EMT and cancer metastasis by restraining autophagy. Oncogene. 2017;36:667-677. DOI: 10.1038/onc.2016.237
  171. 171. Pang M, Wang H, Rao P, Zhao Y, Xie J, Cao Q, et al. Autophagy links β-catenin and Smad signaling to promote epithelial-mesenchymal transition via upregulation of integrin linked kinase. International Journal of Biochemistry and Cell Biology. 2016;76:123-134. DOI: 10.1016/j.biocel.2016.05.010
  172. 172. Goodenough DA, Goliger JA, Paul DL. Connexins, connexons, and intercellular communication. Annual Review of Biochemistry. 1996;65:475-502. DOI: 10.1146/
  173. 173. Goodenough DA, Paul DL. Beyond the gap: Functions of unpaired connexon channels. Nature Reviews Molecular Cell Biology. 2003;4:285-294. DOI: 10.1038/nrm1072
  174. 174. Hervé JC, Derangeon M. Gap-junction-mediated cell-to-cell communication. Cell and Tissue Research. 2013;352:21-31. DOI: 10.1007/s00441-012-1485-6
  175. 175. Prochnow N, Dermietzel R. Connexons and cell adhesion: A romantic phase. Histochemistry and Cell Biology. 2008;130:71-77. DOI: 10.1007/s00418-008-0434-7
  176. 176. Hervé JC, Bourmeyster N, Sarrouilhe D. Diversity in protein-protein interactions of connexins: Emerging roles. Biochimica et Biophysica Acta. 2004;1662:22-41. DOI: 10.1016/j.bbamem.2003.10.022
  177. 177. Singh D, Solan JL, Taffet SM, Javier R, Lampe PD. Connexin 43 interacts with zona occludens-1 and -2 proteins in a cell cycle stage-specific manner. Journal of Biological Chemistry. 2005;280:30416-30421. DOI: 10.1074/jbc.M506799200
  178. 178. Green KJ, Jones JC. Desmosomes and hemidesmosomes: Structure and function of molecular components. FASEB Journal. 1996;10:871-881
  179. 179. Nekrasova O, Green KJ. Desmosome assembly and dynamics. Trends in Cell Biology. 2013;23:537-546. DOI: 10.1016/j.tcb.2013.06.004
  180. 180. Getsios S, Huen AC, Green KJ. Working out the strength and flexibility of desmosomes. Nature Reviews Molecular Cell Biology. 2004;5:271-281. DOI: 10.1038/nrm1356
  181. 181. Wallis S, Lloyd S, Wise I, Ireland G, Fleming TP, Garrod D. The alpha isoform of protein kinase C is involved in signaling the response of desmosomes to wounding in cultured epithelial cells. Molecular Biology of the Cell. 2000;11:1077-1092
  182. 182. Garrod DR, Berika MY, Bardsley WF, Holmes D, Tabernero L. Hyper-adhesion in desmosomes: Its regulation in wound healing and possible relationship to cadherin crystal structure. Journal of Cell Science. 2005;118:5743-5754. DOI: 10.1242/jcs.02700
  183. 183. Goodenough DA, Gilula NB. The splitting of hepatocyte gap junctions and zonulae occludentes with hypertonic disaccharides. Journal of Cell Biology. 1974;61:575-590
  184. 184. Ghoshroy S, Goodenough DA, Sosinsky GE. Preparation, characterization, and structure of half gap junctional layers split with urea and EGTA. Journal of Membrane Biology. 1995;146:15-28
  185. 185. Berthoud VM, Minogue PJ, Laing JG, Beyer EC. Pathways for degradation of connexins and gap junctions. Cardiovascular Research. 2004;62:256-267. DOI: 10.1016/j.cardiores.2003.12.021
  186. 186. Bjorkman N. A study of the ultrastructure of the granulosa cells of the rat ovary. Acta Anatomica (Basel). 1962;51:125-147
  187. 187. Carette D, Gilleron J, Denizot JP, Grant K, Pointis G, Segretain D. New cellular mechanisms of gap junction degradation and recycling. Biology of the Cell. 2015;107:218-231. DOI: 10.1111/boc.201400048
  188. 188. Zou H, Zhuo L, Han T, Hu D, Yang X, Wang Y, et al. Autophagy and gap junctional intercellular communication inhibition are involved in cadmium-induced apoptosis in rat liver cells. Biochemical and Biophysical Research Communications. 2015;459:713-719. DOI: 10.1016/j.bbrc.2015.03.027
  189. 189. Falk MM, Kells RM, Berthoud VM. Degradation of connexins and gap junctions. FEBS Letters. 2014;588(8):1221-1229. DOI: 10.1016/j.febslet.2014.01.031
  190. 190. Bejarano E, Girao H, Yuste A, Patel B, Marques C, Spray DC, et al. Autophagy modulates dynamics of connexins at the plasma membrane in a ubiquitin-dependent manner. Molecular Biology of the Cell. 2012;23:2156-2169. DOI: 10.1091/mbc.E11-10-0844
  191. 191. Garrod D, Tabernero L. Hyper-adhesion: A unique property of desmosomes. Cell Communication & Adhesion. 2014;21:249-256. DOI: 10.3109/15419061.2014.930133
  192. 192. Allen TD, Potten CS. Desmosomal form, fate, and function in mammalian epidermis. Journal of Ultrastructure Research. 1975;51:94-105
  193. 193. McHarg S, Hopkins G, Lim L, Garrod D. Down-regulation of desmosomes in cultured cells: The roles of PKC, microtubules and lysosomal/proteasomal degradation. PLoS One. 2014;9:e108570. DOI: 10.1371/journal.pone.0108570
  194. 194. Calkins CC, Setzer SV, Jennings JM, Summers S, Tsunoda K, Amagai M, et al. Desmoglein endocytosis and desmosome disassembly are coordinated responses to pemphigus autoantibodies. Journal of Biological Chemistry. 2006;281:7623-7634. DOI: 10.1074/jbc.M512447200
  195. 195. Barczyk M, Carracedo S, Gullberg D. Integrins. Cell and Tissue Research. 2010;339:269-280. DOI: 10.1007/s00441-009-0834-6
  196. 196. Hynes RO. Integrins: Bidirectional, allosteric signaling machines. Cell. 2002;110:673-687
  197. 197. Oommen S, Francois M, Kawasaki M, Murrell M, Kawasaki K, Porntaveetus T, et al. Cytoplasmic plaque formation in hemidesmosome development is dependent on SoxF transcription factor function. PLoS One. 2012;7:e43857. DOI: 10.1371/journal.pone.0043857
  198. 198. Burridge K, Guilluy C. Focal adhesions, stress fibers and mechanical tension. Experimental Cell Research. 2016;343:14-20. DOI: 10.1016/j.yexcr.2015.10.029
  199. 199. Wehrle-Haller B. Structure and function of focal adhesions. Current Opinion in Cell Biology. 2012;24:116-124. DOI: 10.1016/
  200. 200. Paluch EK, Aspalter IM, Sixt M. Focal adhesion-independent cell migration. Annual Review of Cell and Developmental Biology. 2016;32:469-490. DOI: 10.1146/annurev-cellbio-111315-125341
  201. 201. Tuloup-Minguez V, Hamaï A, Greffard A, Nicolas V, Codogno P, Botti J. Autophagy modulates cell migration and β1 integrin membrane recycling. Cell Cycle. 2013;12:3317-3328. DOI: 10.4161/cc.26298
  202. 202. De Franceschi N, Hamidi H, Alanko J, Sahgal P, Ivaska J. Integrin traffic—the update. Journal of Cell Science. 2015;128:839-852. DOI: 10.1242/jcs.161653
  203. 203. Vlahakis A, Jayanta D. The interconnections between autophagy and integrin-mediated cell adhesion. Journal of Molecular Biology. 2016:1-16. DOI: 10.1016/j.jmb.2016.11.027
  204. 204. Kenific CM, Torsen W, Jayanta D. Autophagy in adhesion and migration. Journal of Cell Science. 2016;129:3685-3693. DOI: 10.1242/jcs.188490

Written By

Catalina Flores-Maldonado, Odette Verdejo-Torres, Jessica Campos- Blázquez, Agustín Ruiz Cabrera, Vicky García-Hernández, Ruth Rincón-Heredia and Rubén G. Contreras

Submitted: October 11th, 2016 Reviewed: April 21st, 2017 Published: August 30th, 2017