Regulation of Claudin-3 Expression in Kidney Tubular Epithelial Cells

Shaista Anwer

A thesis submitted in conformity with the requirements for the degree of Master of Science Institute of Medical Science University of Toronto

Shaista Anwer Master of Science Institute of Medical Science University of Toronto 2020

Abstract

The overall objective of my studies was to gain insight into the regulation of the tight junction protein, claudin-3, upon inflammatory stimuli in kidney tubular epithelial cells. Claudins mediate paracellular transport and modulate key cellular events like proliferation, migration and differentiation. The inflammatory cytokine Tumor Necrosis Factor-α (TNFα) is a pathogenic factor in kidney disease and alters epithelial permeability. However, the effect of TNFα on claudin-3 expression in the tubules and the mechanisms are not defined. My studies showed that TNFα elevated claudin-3 expression in kidney tubular cells. This effect was due to increased claudin-3 synthesis and mediated by two signaling pathways: extracellular signal regulated kinase- dependent activation of NFκB and protein kinase A-induced CREB1 activation. Claudin-3 overexpression elevated transepithelial resistance in tubular cells and it may play a role in regulating tubular epithelial permeability. Claudin-3 downregulation also affected cell cycle proteins; thus, claudin-3 may also affect cell proliferation.

Acknowledgements

I would like to dedicate this thesis to two important people in my life: my mother, Tahniyat Anwer and my supervisor, Dr. Katalin Szászi. My mom has been a great source of inspiration and a pillar of strength in my life and has always supported me in my endeavours. I will be forever grateful for all her hard work and the sacrifices she has made throughout her life. I would also like to acknowledge my supervisor, Dr. Katalin Szászi, for her constant encouragement and guidance throughout my Master’s. She has played a major role in my academic and personal growth and she has definitely inspired me to be a well-rounded researcher. I am extremely grateful that I had this opportunity to work with her. It would have been impossible to write this thesis without her encouragement and assistance. I would also like to thank my dad and my sisters for always supporting me and for understanding my dedication and passion for research. I would also like to thank my grandparents, uncles, aunts and cousins for always wishing me the best and congratulating me for all my achievements. I would also like to thank my close friends for cheering me up and constantly supporting me. I would also like to express my gratitude to my program advisory committee members, Dr. András Kapus and Dr. Mauricio Terebiznik, for their valuable advice and guidance throughout my Master’s. I really appreciate their input and their passion for science which has fueled my curiosity for many aspects of my project. I would also like to thank all the current and previous lab members, including Qinghong Dan, Shruthi Venugopal, Emily Branchard, Iris Huang, Jenny Xiao and Vida Maksimoska, for helping me with my project. I have had a very enjoyable experience of working with all of them. I have especially learnt a lot from Qinghong Dan and Shruthi Venugopal and I am thankful that I had the opportunity to work with such excellent researchers. I would also like to thank members of the Kapus lab, Pam Speight, Michael Kofler and Zena Miranda, for their kindness, positivity and for providing a different perspective of life. I am also thankful for Muskan Gupta, Zahra Khan, Misha Ditmans and Samantha Mahabir for creating a positive, lively and friendly atmosphere in our office. They were definitely a great addition to our office, and I had a great time having endless conversations with them. Finally, I am also really appreciative of the continuous support of all the people on the 6th floor as well as the core facility of the Li Ka Shing Knowledge Institute.

iii

Statement of Contributions

I am the primary author of this thesis. I have participated in the planning, execution and analysis of research experimentation. I have received funding from UofT Open Fellowship and St. Michael’s Hospital Research Training Centre Scholarship.

I would also like to acknowledge the contributions of my supervisor Dr. Katalin Szászi and my program advisory committee (PAC) members, Dr. András Kapus and Dr. Mauricio Terebiznik. My supervisor and PAC members provided tremendous mentorship and guidance in planning, executing, analysing and interpreting my experiments and experimental findings.

I would also like to acknowledge the contributions of the following individuals:

Qinghong Dan, senior lab technician – provided assistance with some of my experiments.

Emily Branchard, past lab member – made initial observations for this project.

Caterina Di Ciano-Oliveira, bioimaging specialist – provided microscopy training and assisting with acquisition of confocal images and 3D analysis of confocal images.

Vida Maksimoska, volunteer – provided assistance with some of my experiments.

Table of Contents

Abstract...... ii Acknowledgements...... iii Statement of Contributions...... iv List of Abbreviations...... vii List of Figures...... ix List of Tables...... ix List of Publications...... x

CHAPTER 1 Literature Review...... 1 1.1 Epithelial cells...... 1 1.2 Intercellular junctional complexes...... 1 1.3 Tight junctions: structure and function...... 4 1.3.1 Tight junction structure...... 4 1.3.2 Overview of tight junction functions: gate, fence and signaling…...... 4 1.4 Tight junction proteins...... 5 1.4.1 Claudin proteins...... 5 a. Claudin structure…...... 5 b. Functions of claudins...... 8 c. Claudin localization and expression...... 9 d. Claudin and tight junction regulation...... 9 e. Role of claudins in diseases...... 11 1.4.1.1 Claudin-3 a. Claudin-3 structure, expression and interactions...... 12 b. Functions of claudin-3...... 14 c. Claudin-3 expression regulation...... 18 d. Role of claudin-3 in diseases...... 24

1.5 Inflammatory cytokines and TJs...... 27 1.5.1 TNFα structure and synthesis...... 27 1.5.2 TNFα receptor activation and signaling pathways...... 28 1.5.3 Role of TNFα in kidney diseases...... 32

CHAPTER 2 Research Aims/Objectives and Hypotheses...... 33 CHAPTER 3 Materials and Methods...... 36

CHAPTER 4 TNFα-induced claudin-3 expression regulation in kidney tubular epithelial cells...... 41 Results...... 41 4.1 TNFα induced an increase in Cldn-3 protein expression...... 41 4.2 TNFα did not alter Cldn-3 degradation and increased Cldn-3 mRNA levels...... 44 4.3 TNFα-induced increase in Cldn-3 requires NFκB and CREB1 and not Slug...... 46 4.4 TNFα-induced increase in Cldn-3 requires ERK-mediated NFκB activation...... 51 4.5 TNFα-induced elevation in Cldn-3 requires PKA-dependent CREB1 activation...... 54 4.6 Sp1 regulates TNFα-induced Cldn-3 upregulation...... 57 4.7 Cldn-3 overexpression increased transepithelial resistance...... 59 4.8 Cldn-3 silencing upregulates the cell cycle protein p27kip1...... 62

CHAPTER 5 General discussion, conclusions and future directions...... 64 5.1 General discussion...... 64 5.2 Future directions…...... 73 5.3 Conclusions………...... 74

References...... 75

List of Abbreviations

ADAM Α disintegrin and metalloprotease AEC Alveolar epithelial cells AJ Adherens junction AP Activating protein BBB Blood-brain barrier BSA Bovine serum albumin BTB Blood-testis barrier cAMP Cyclic adenosine monophosphate CKD Chronic kidney disease Cldn Claudin CPE Clostridium perfringens enterotoxin CRE cAMP response element CREB cAMP response element binding protein DD Death-domain ECIS Electric cell-substrate impedance sensing ECL Extracellular loop ECM Extracellular matrix EGF Epidermal growth factor EGFR Epidermal growth factor receptor EMT Epidermal-mesenchymal transition ERK Extracellular signal regulated kinase FADD Fas-associated death domain GAPDH glyceraldehyde-3-phosphate dehydrogenase IBD Inflammatory bowel disease IL Interleukin IMCD Inner medullary collecting duct JAM Junctional adhesion molecule JNK c-Jun N-terminal kinase

vii

KD Knockdown KO Knock-out LLCPK Lilly Laboratories Cell Porcine Kidney LPS Lipopolysaccharide MAPK Mitogen-activated protein kinase MLCK Myosin light chain kinase MMP Matrix metalloproteinase NFκB Nuclear factor-Kappa-B NR Non-related PKA Protein kinase A RIP Receptor interacting protein RPE Retinal pigment epithelium RVP Rat ventral prostate siRNA Small interfering RNA Sp1 Specificity protein 1 TACE Tumour necrosis factor-α-converting enzyme TAMP Tight junction-associated Marvel proteins TER Transepithelial resistance TGFβ Transforming growth factor β TI Tubulointerstitial inflammation TJ Tight junctions TNBC Triple negative breast cancer TNFR TNFα receptor TNFα Tumour necrosis factor-α TRADD TNFR-associated via death domain TRAF TNFR activating factors ZO Zonula occludens

viii

List of Figures

Fig 1-1 Intercellular junctional complexes in epithelial cells...... 3 Fig 1-2 General claudin structure and modification sites...... 7 Fig 1-3 Simplified TNFα receptor activation of classical pathways...... 30 Fig 1-4 Simplified TNFα receptor crosstalk overview in epithelial cells...... 31 Fig 4-1 TNFα increased Cldn-3 protein expression in tubular cells...... 42 Fig 4-2 TNFα did not alter Cldn-3 degradation and increased Cldn-3 mRNA levels...... 45 Fig 4-3 TNFα-induced increase in Cldn-3 required NFκB and CREB1...... 48 Fig 4-4 TNFα-induced increase in Cldn-3 is mediated by ERK and NFκB...... 52 Fig 4-5 TNFα-induced increase in Cldn-3 required PKA and CREB1...... 55 Fig 4-6 Sp1 regulates TNFα-induced increase in Cldn-3...... 58 Fig 4-7 Cldn-3 overexpression increased TER...... 60 Fig 4-8 Cldn-3 regulates cell cycle protein p27kip1...... 63 Fig 5-1 Proposed mechanisms involved in regulating Cldn-3 expression...... 72

List of Tables

Table 3-1. List of inhibitor and activator reagents...... 40

List of publications

Manuscript under preparation: 1. Anwer S, Branchard E, Dan Q, Szaszi K. Tumour-necrosis factor-α induces Claudin-3 upregulation in kidney tubular cells. To be submitted to Am J Physiol Cell Physiol.

Published papers 1. Venugopal S, Anwer S, Szaszi K. Claudin-2: beyond permeability functions. Review. Int J Mol Sci. 2019 Nov; 20(22):5655 2. Dan Q, Shi Y, Rabani R, Venugopal S, Xiao J, Anwer S, Ding M, Speight P, Pan W, Alexander RT, Kapus A, Szaszi K. Claudin-2 suppresses GEF-H1, RhoA, and MRTF thereby impacting proliferation and profibrotic phenotype of tubular cells. J Biol Chem. 2019 Sep; 294(42):15446- 15465. 3. Anwer S, Amoozadeh Y, Dan Q, Venugopal, S, Shi Y, Branchard E, Liedtke, E, Ailenberg, M, Rotstein O.D, Kapus A and Szászi K. Cell confluence regulates Claudin-2 expression: possible role for ZO-1 and Rac. Am J Physiol Cell Physiol. 2018 Mar; 314(3):C366-C378. 4. Amoozadeh Y, Dan Q, Anwer S, Huang HH, Barbieri V, Waheed F and Szászi K. Tumor Necrosis Factor-α increases claudin-1, 4 and 7 expression in tubular cells: role in permeability changes. J. Cellular Physiol. 2017 Aug; 232(8):2210-2220.

CHAPTER 1: LITERATURE REVIEW

1.1 Epithelial Cells The vital organs of the human body are lined with epithelial cells which are specialized for absorption, secretion and barrier formation. For example, the tubular epithelium in the kidney nephron facilitates efficient transport of ions, water and other substances, thereby, maintaining ion and fluid homeostasis. The layer generates a permeability barrier that allows highly regulated transcellular and paracellular transport (Tsukita & Furuse, 2006). This is due to the apicobasal polarity of cells generated by the tight junctions, which is vital for directional transport processes. A distinct apical side of the cells faces the lumen, while a basal side faces the renal tissue and is anchored to the extracellular matrix (ECM). The lateral side faces the neighbouring cell and forms a paracellular space (Campanale et al., 2017). The basal and lateral membrane can freely mix, and therefore, are referred to as the basolateral membranes. The protein and lipid compositions of the apical and basolateral membranes determine the overall permeability of an epithelial layer (Szaszi & Amoozadeh, 2014).

1.2 Intercellular Junctional Complexes There are four distinct intercellular junctional complexes which connect neighbouring cells, including tight junctions (TJ), adherens junctions (AJ), desmosomes and gap junctions (Figure 1- 1) (Blaskewicz et al., 2011; Fleming et al., 1992; Raviola & Raviola, 1978). Each junctional complex is composed of multiple proteins which define distinct roles in barrier formation, transport, cell adhesion and signaling.

The tubular epithelium controls water and solute exchange via paracellular and transcellular transport pathways. The TJs play a major role in mediating paracellular transport and are key for the transcellular pathway as well (Anderson, 2001; Dahlgren & Lennernäs, 2019; Karasov, 2017). The paracellular pathway allows passive transport of small solutes and water through the TJs and is driven by the electrochemical gradient of these molecules. Specific transport proteins present on the apical and basolateral membranes are also vital for the active transcellular transport. The

1 distinct localization of the transport proteins is due to the apico-basal polarity of the cells which is maintained by the tight junctions.

AJs and desmosomes hold cells together and are formed by distinct members of the cadherin family of adhesion proteins and nectins. AJs are located below the TJs and form adhesion belts which encircle the cells, whereas, desmosomes connect cells to the intermediate filaments (Hartsock & Nelson, 2008; Kowalczyk & Green, 2013). Together, these junctions provide tensile strength to the cell structure. Gap junctions are composed of connexins which connect the cytoplasm of adjacent cells and allow intercellular passage of ions and small molecules (Blaskewicz et al., 2011).

For my thesis project, I have studied the regulation of a specific TJ protein in kidney tubular cells, therefore, the subsequent chapters will focus on the TJ protein complex.

Transcellular Paracellular Apical transport transport membrane

Tight junctions Adherens junctions Desmosomes Gap junctions

Focal Hemi-

Basolateral adhesion desmosome membrane

JAMs Regulatory proteins Claudins Signaling

proteins

r Acto- o

myosin ring protein Adapt TAMP s Transcription factors

Figure 1-1. Intercellular junctional complexes in epithelial cells. Simplified scheme of the types of intercellular junctional complexes present in epithelial cells, including tight junctions (TJ), adherens junction, desmosomes and gap junctions. The figure also demonstrates the distinction between apical and basolateral membranes, which generates polarity in epithelial cells. Transcellular transport creates a chemical and electrical gradient which is the driving force for the paracellular transport pathway. The insert provides an overview of the types of TJ proteins, including various transmembrane proteins, and their association with cytoplasmic plaque proteins. Together, they are involved in regulating TJ dynamics and mediate the effects on various cellular events.

1.3 Tight Junctions: Structure and Function 1.3.1 Tight junction Structure Tight junctions (TJ), or zona occludens (ZO), are the most apical junctional complexes. This structure forms a belt-like structure that surrounds epithelial cells and forms a continuous seal (Furuse, 2010). Decades ago, electron microscopy identified TJs as focal attachments which eliminate the gap between adjacent epithelial cells in the paracellular space consisting of cross- linked linear fibrils (TJ strands) (Claude, 1973; Farquhar, 1963; Hirsch & Noske, 1993). This structure forms a permeability barrier and allows for selective passive transport of substances across the paracellular pathway. It also separates the apical and basolateral membranes. Subsequent work revealed that TJs are composed of several transmembrane and cytosolic proteins, as discussed in Chapter 1.4.

1.3.2 Overview of Tight Junction Functions: Gate, Fence and Signaling As mentioned above, TJs have two classic functions: generating paracellular permeability (gate function) and maintaining polarity of the epithelia (fence function) (Matter & Balda, 2003). The gate function means that TJs seal off the intercellular space and at the same time generate a selective pathway for specific ions (Shen et al., 2011). Two distinct paracellular pathways have been identified in mediating the movement of molecules of different sizes. Firstly, a pore pathway permits transport of solutes smaller than 4 Angstrom in radius and displays ionic charge selectivity. Second, the leak pathway allows transport of other larger solutes (Van Itallie et al., 2008; Watson et al., 2005). The former is mediated by the claudin family of proteins, which will be further described below. The identity of the leak pathway remains unknown. The fence function maintains the apico-basal polarity of epithelial cells by preventing mixing of lipids and proteins via free diffusion in the distinct membranes (Schlüter & Margolis, 2012).

In addition to these classic functions, the TJs also act as signaling platforms which engage in bi- directional signaling (Matter & Balda, 2003; Zihni et al., 2014). TJs can transduce extracellular information to regulate cell behaviour, including cell proliferation, differentiation and migration, and on the other hand, they can also be targets of different signaling mechanisms (Bhat et al.,

2019; Takano et al., 2014). This is mediated by the vast majority of molecules present in the cytoplasmic side, including cytoskeletal elements, transcription factors and other signaling and regulatory proteins (Rodgers & Fanning, 2011; Zihni et al., 2016). These functions will be further discussed in Chapter 1.4.1.

1.4 Tight Junction Proteins TJs contain transmembrane and cytosolic proteins. The transmembrane proteins can be classified into three groups: claudins, tight-junction-associated Marvel proteins (TAMPs) and single span proteins. TAMPs are a family of tetra-span membrane proteins which includes occludin, tricellulin and MarvelD3 proteins. The main single span membrane proteins are the junctional adhesion molecules (JAM) (Anderson & Van Itallie, 2009; González-Mariscal et al., 2003). Signaling proteins, adaptor proteins, transcription factors and cytoskeletal elements constitute the cytosolic proteins, which collectively form the cytoplasmic plaque (see insert in Figure 1-1) (Furuse, 2010; Günzel & Yu, 2013).

I will be discussing the claudin family of transmembrane TJ proteins in the next sections, followed by specific emphasis on my protein of interest, claudin-3.

1.4.1 Claudin proteins a) Claudin structure The claudin family of proteins consist of 27 small molecular weight (21-28 kDa) tetraspan membrane proteins which form the backbone of the TJs and are vital for maintaining the TJ structural integrity. Claudins have 4 intramembranous portions, connected by two extracellular loops (ECL1 and ECL2) and a small intracellular loop connecting the second and third transmembrane sections (Baumgartner et al., 2017; Gerd Krause et al., 2008) (Figure 1-2). The extracellular loops span the paracellular space (Günzel & Yu, 2013). ECL1 is responsible for paracellular permeability and charge selectivity (Colegio et al., 2003). ECL1 also contains two conserved cysteines which form disulfide bonds to stabilize the pore (Wen et al., 2004), and negative and positive charges that determine charge selectivity (Amasheh, 2002; Krause et al.,

2008). Claudins can form homotypic and heterotypic interactions with each other or other TJ proteins. The extracellular loops, specifically ECL1, engage in trans (intercellular) interactions (Lim et al., 2008).

Claudins also contain short intracellular N- and C-terminal portions (Furuse et al., 1998). Although the C-terminal has variable lengths and sequences, the last amino acids at the C-terminal of most claudins corresponds to a highly conserved PDZ domain-binding motif, of which the last two C- terminal amino acids (YV) exhibits 100% conservation (Nomme et al., 2015). The PDZ-binding motif was shown to associate with the PDZ domains of a variety of TJ plaque scaffolding proteins including the ZO-1-3, MUPP1 and PATJ, which connect claudins to the actin cytoskeleton (Günzel & Yu, 2013; Itoh et al., 1999). The shorter N-terminus region currently has undefined functions. Claudins also engage in cis (side-to-side) interactions, which is mediated by the transmembrane domains (Van Itallie & Anderson, 2013).

The claudin family proteins are structurally conserved, however, they are highly divergent at the sequence level (Tikiyani & Babu, 2019). Claudins 1-10, 14, 15, 17 and 19 are referred to as classic claudins due to a high degree of structural homology and functional similarity. (Krause et al., 2008). These classifications, however, are not discrete and are still debated.

ECL1

ECL2

Paracellular space

TM2

TM1 TM3 TM4 Cytoplasm H N 2 Y-V-COOH

Palmitoylation site

Phosphorylation site

CPE binding region for claudins 3, 4, 6, 7

PDZ domain binding motif

Figure 1-2. General claudin structure and modification sites. Claudins contain four transmembrane (TM) domains connected by two extracellular loops (ECL) and short intracellular N- and C-terminal ends. ECL2 contains a Clostridium perfringens enterotoxin (CPE) binding region in some claudins, including claudin-3, which represents a promising target for monoclonal antibody therapy. The C-terminal of most claudins contains a highly conserved PDZ domain binding motif (YV) which interacts with the PDZ domain of various adaptor proteins. These proteins connect claudins to cytoskeletal elements and have been implicated in signal transduction. Claudins also contain post-translational modification sites which are vital for regulating its stability and interaction with other claudins.

7 b) Functions of claudins In addition to providing TJ structural integrity, claudins also mediate paracellular permeability. (Balkovetz, 2006; Szaszi & Amoozadeh, 2014). As such, claudins can be classified as either pore- forming, that form specific paracellular channels, or as barrier-forming (claudins 1, 3, 5, 11, 14, 19), that seal the paracellular space and reduce permeability (G. Krause et al., 2015; Van Itallie & Anderson, 2004; Fromm et al., 2017). This classification was determined by various overexpression and knockdown studies conducted in epithelial cells. Pore-forming claudins have unique cation or anion selectivity (permselectivity) and can be further classified as cation- selective (claudins 2, 10b, 15) or anion-selective (claudins 10a and 17). Moreover, the function of some claudins remains unclear and these are classified as either inconsistent claudins (claudins 4 ,7, 8 and 16); or claudins with unknown functions (claudins 6, 9, 12, 13, 18, 20-27) (Anderson & Van Itallie, 2009; Colegio et al., 2003; Schulzke et al., 2012). Due to the complex, tissue-specific expression pattern of claudins (discussed further in the next section), it is imperative that functional studies of each claudin are conducted in the context of a particular cell or tissue type. This is especially important due to the dependence of epithelial permselectivity on the overall claudin expression profile. Generally, knockout (KO) or knockdown (KD) of a specific sealing claudin results in increased epithelial leakiness.

In addition to these classic permeability functions, certain claudins have been shown to regulate non-classic functions which affect signal transduction, and influence cell proliferation, differentiation and migration (Hagen, 2017; Hichino et al., 2017; Li et al., 2019; Shi et al., 2018). These functions are most likely mediated via cytoplasmic interactions of claudins with the cytoplasmic plaque proteins, such as adaptor and scaffolding proteins. These proteins connect claudins to various effectors and cytoskeletal elements and this interaction is key for mediating effects on signaling pathways which alters cell behaviour and regulates the above-mentioned non-classic functions. These non-classic functions will be discussed further in the context of claudin-3 Chapter 1.5.1.1.

8 c) Claudin localization and expression Each claudin has a unique tissue-specific expression profile, and various cells/tissues, have a characteristic claudin profile. This differential expression of claudins has been shown in many mammalian tissues, including the kidney, intestine, lungs, retina, brain and prostate glands. In the kidney, the paracellular permeability generally decreases from the proximal tubules to the collecting ducts, which is important for altering the transport properties and composition of the filtrate (Szaszi & Amoozadeh, 2014). Immunohistochemistry and Northern blot analysis revealed the localization of claudin expression across the tubules. The general trend shows abundance of channel-forming claudins in the proximal tubule and descending limb of the loop of Henle; whereas, sealing claudins were highly expressed in the ascending limb, distal tubule and collecting duct (Kirk et al., 2010; Kiuchi-Saishin et al., 2002; Krug et al., 2012; Lee et al., 2006; Reyes et al., 2002). This localization correlates well with the defined functions of these segments, as there is increased ion reabsorption in the initial segments of the nephron, followed by reduced solute transport near the end.

Taken together, these segmental expression profiles of claudins play a major role in regulating both paracellular permeability and overall tubular transport. However, localizing claudin proteins has some challenges. Potential cross-reaction of antibodies between claudin isoforms and species differences pose difficulties (Gonzalez-Mariscal et al., 2006). In addition, there is evidence that certain claudins can affect the expression of other claudins (see Chapter 1.5.1.1). d) Claudin and tight junction regulation TJs are highly dynamic structures which are continuously assembled and disassembled in response to various cellular and environmental factors. Although the molecular mechanisms involved in regulating TJs are incompletely understood, it is known that various signaling pathways are involved in TJ modifications which alter the TJ composition. TJ proteins are dynamically regulated by a multitude of factors which play a vital role in TJ assembly and remodeling. These factors, through various mechanisms, alter both protein synthesis and degradation. TJ proteins also undergo post-translational modifications which affect its TJ

9 localization, trafficking and interactions with other proteins. In addition, TJ protein turnover is also highly regulated. When TJ proteins are inserted into the membrane, they are protected from degradation, hence, endocytosis and recycling processes are key for regulating protein levels as well. These processes are mediated by various cytoskeletal elements and are also vital for maintaining TJ integrity (Matsuda et al., 2004; Rodgers & Fanning, 2011). TJ proteins degrade in late endosomes, followed by lysosomal-mediated degradation (Bruewer et al., 2003; Capaldo et al., 2014; Weber et al., 2017), and via the ubiquitin-proteasome pathway (Asaka et al., 2011; Mandel et al., 2012; Schmidt et al., 2019).

In general, many aspects of the above-mentioned TJ protein regulation reflect claudin regulation as well. As mentioned previously, the intracellular portions of most claudins are post- translationally modified and these modifications include phosphorylation, palmitoylation, ubiquitination, nitration and SUMOylation (Cummins, 2012; Dörfel & Huber, 2012; González- Mariscal et al., 2008; Murakami et al., 2009; T. Suzuki et al., 2009; Van Itallie et al., 2005). These modifications allow complex protein-protein interactions to occur and allow incorporation of claudins into the TJs (Hou & Goodenough, 2010). Almost all claudins are palmitoylated at the C- X-X-C motifs in the intracellular regions of the second and fourth transmembrane domains. In claudin-14, palmitoylation at both of these sites was required for its efficient localization to the TJs, without affecting its stability or TJ assembly (Furuse, 2010; Van Itallie et al., 2005). In addition, the serine or threonine residues in the C-terminus region of several claudins, including claudin- 3, are highly phosphorylated and this phosphorylation has been shown to regulate TJ localization and barrier function (D’Souza et al., 2005; Furuse, 2010; Ikari et al., 2006).

There is increasing evidence of various stimuli regulating claudin transcription as several transcription factors have been shown to mediate both basal and stimulated claudin expression in various cell lines. For instance, the Forkhead box (foxO) gene family bind to a conserved motif on the claudin-5 promoter, and together with β-catenin, they act to suppress its transcription (Taddei et al., 2008). Several other transcriptional repressors, including Snail, ZEB, Klf8 and Twist, were also shown to downregulate claudins (Ikenouchi et al., 2003; Lin et al., 2013; Wang et al.,

2019). Caudal-related homeobox (Cdx) proteins were shown to activate the claudin-2 promoter, thereby, regulating intestinal differentiation and promoting carcinogenesis (Jung et al., 2011; Sakaguchi et al., 2002; Suzuki et al., 2011; Zhang et al., 2015). The transcription factors NFκB and CREB have also been shown to regulate the expression of some claudins, and this will be further discussed in the context of claudin-3 in Chapters 1.4.1.1 and 1.5. All in all, transcription factor- mediated regulation of claudin synthesis alters permeability properties and signaling functions which leads to modulation of other cellular functions, including differentiation and proliferation (Khan & Asif, 2015). e) Role of claudins in diseases Many studies have provided evidence of claudin expression, functional and localization dysregulation to be associated with various pathological conditions. Generally, overactive signaling pathways can alter claudin expression in various cancers and inflammatory diseases. The crucial role of the tubular epithelium in regulating tubular transport implies that alterations of the TJ proteins can inevitably alter kidney ion and fluid homeostasis, thereby contributing to pathological conditions. In this section, I will provide a brief overview of claudins known to play a role in specific kidney diseases and claudins that are targets of various conditions.

A loss-of-function mutation in the claudin-16 or -19 genes has been shown to lead to hypercalciuria, a renal tubular disorder causing kidney stones, urinary tract infection and polyuria, and eventually leading to early onset chronic kidney disease (CKD) (Claverie-Martin, 2015; Hou & Goodenough, 2010). In addition to genetic mutations, altered claudin expression levels and post-translational modifications are also associated with certain diseases. A study has shown that claudin-2 is post-translationally modified in response to oxidative stress in type 1 diabetic nephropathy. These modifications, including phosphorylation, SUMOylation and nitration of specific residues, led to lowered claudin-2 expression. Ultimately, this decrease in claudin-2 was shown to be associated with increased sodium excretion in the urine, a condition known as natriuresis (Molina-Jijón et al., 2014). In addition, claudin-7 was found to be increasingly phosphorylated in pseudohypoaldosteronism type II (PHAII), a condition caused by

11 mutation in WNK protein kinase (Tatum et al., 2007). This resulted in increased paracellular permeability of Cl–, which may play a role in hypertension (see section 1.4.1.1 for role of claudin- 3 in diseases).

The focus of the next chapter will be specifically on claudin-3 expression, function, regulation and role in various pathologies.

1.4.1.1 Claudin-3 a) Claudin-3 structure, expression and interactions Claudin-3 (Cldn-3), initially named Rat Ventral Prostate.1 Protein Homolog (RVP-1), is a transmembrane protein with three isoforms (a, b and c). The longer isoform is 220 amino acids in length with a molecular weight of 23 kDa (Li et al., 2010; Tipsmark et al., 2008; Tipsmark & Madsen, 2012). Cldn-3 has a high degree of homology with Cldn-5, -15 and -19 (Irudayanathan et al., 2017; Piontek et al., 2017). Cldn-3 is a tetraspan protein, with a structure that is similar to other members of the family described in Chapter 1.4.1. The ECL2 of Cldn-3 has been shown to interact with the bacterial toxin Clostridium perfingens enterotoxin (CPE), leading to membrane pore complex formation and rapid cell death (Pahle et al., 2017; Sonoda et al., 1999).

Cldn-3 engages in both homo- and heterotypic adhesions, via cis and trans interactions. Mutagenesis studies have revealed that the hydrogen-bonding potential of Cldn-3, characterized by glutamine 56, glutamine 62 and glutamic acid 158 sites, are vital for mediating the trans interactions and TJ strand formation (Piontek et al., 2017). These residues were found to be conserved among other barrier-forming claudins as well, and therefore, may play a vital role in paracellular barrier formation. Cldn-3 was shown to form Cldn-3 homopolymers via cis interactions (Milatz et al., 2015). Cldn-3 also binds to Cldn-4 in acinar cells of the salivary gland, and formed cis interactions with Cldn-5 in type II alveolar epithelial cells and in brain endothelial cells (Peppi & Ghabriel, 2004; Santos et al., 2007; F. Wang et al., 2003). Cldn-3 also participates in trans interactions by binding to Cldn-1, -2 and -15 to form paired strands on neighbouring cells (Furuse et al., 1999; Milatz et al., 2015; Peppi & Ghabriel, 2004).

The intracellular interactions of Cldn-3 are vital for its signal-modulating functions. Similar to other claudins, its last C-terminal amino acids correspond to the PDZ-binding motif. In Cldn-3, the last three amino acids are D-Y-V (N- to C-term) (Merino-Gracia et al., 2016; Wu et al., 2015). This motif was shown to associate with a variety of TJ plaque scaffold proteins, including the ZO-1-3 adaptors (Søfteland et al., 2019; Yamaga et al., 2018). The PDZ domain of the neuronal nitric- oxide synthase (nNOS) was also shown to have high affinity for Cldn-3, and this interaction might be dependent on the phosphorylation state of the two tyrosine residues of Cldn-3 at the C- terminus (Merino-Gracia et al., 2016). The intracellular portions of Cldn-3 also undergo post- translational modifications including phosphorylation and palmitoylation. The significance of these modifications will be discussed in section c.

Cldn-3 is ubiquitously expressed in epithelial cells of the kidney, digestive, gastrointestinal (GI) and respiratory tracts, and in the prostate and mammary glands (Coyne et al., 2003; Kaarteenaho-Wiik & Soini, 2009; Morita et al., 1999; Sakai et al., 2007). In the kidney, Cldn-3 is highly expressed in the tighter segments of the nephron, such as the thin and thick ascending limbs of Henle, distal tubules and collecting ducts (Balkovetz, 2006; Kiuchi-Saishin et al., 2002). In the digestive tract, Cldn-3 was shown to be expressed in the salivary glands (parotid and submandibular glands), esophagus, liver, pancreas and gallbladder (Hashizume et al., 2004; Michikawa, 2008; Xu et al., 2014). It was also found in the Peyer’s patches of the small intestine and colon (X. Chen et al., 2018; Mees et al., 2009; Tamagawa et al., 2003).

Interestingly, Cldn-3 was also shown to be expressed in some epidermal and endothelial cells, including sweat glands, keratinocytes, and in the blood-brain barrier (BBB) and blood-testis barrier (BBB) (Günzel & Yu, 2013; Tebbe et al., 2002; Tokumasu et al., 2017; Yamaga et al., 2018). Disease-associated changes in Cldn-3 expression will be discussed in section d.

In addition to the TJs, Cldn-3 was also detected in small unidentified vesicles and in intracellular compartments which colocalized with the endoplasmic reticulum (Rossa et al., 2012).

Interestingly, some studies have found nuclear localization of Cldn-3 in the pancreatic and colon cancer tumours (Gurevich et al., 2014; Oleinikova et al., 2017; Tokuhara et al., 2018).

Cldn-3 also seems to be preferentially expressed during specific stages of organ development. Cldn-3 was found to be a constitutive and vital component of uterine epithelium and placental endothelium during early pregnancy (Challier et al., 2005; Nicholson et al., 2010). Cldn-3 was also implicated in liver and lung development. It was shown to be highly expressed in hepatic stem cells (precursors of hepatoblasts), as compared to hepatoblasts itself (Schmelzer et al., 2006). Interestingly, Cldn-3 expression was also shown to decrease with age in the mouse liver (Tanaka et al., 2018). Further, Cldn-3 is expressed in human fetal lung alveolar and bronchial epithelial cells and affects acinus development and differentiation (Daugherty et al., 2004; Kaarteenaho et al., 2010). In the kidney, Cldn-3 was shown to be upregulated in in-vitro cultured isolated ureteric bud and may play a role in epithelial maturation and polarization (Meyer et al., 2004). b) Functions of claudin-3 The exact permeability role of Cldn-3 is unclear due to differences in the experimental findings in various cell lines. In other words, we cannot affirmatively confirm whether it is a channel forming or a sealing claudin. However, the vast majority of overexpression, KO and KD studies determined Cldn-3 to be have barrier-forming properties which seals the paracellular pathway against transport of small charged or uncharged ions. The evidence for both sealing and pore- forming functions will be discussed below.

Evidence for sealing functions The barrier properties of Cldn-3 are well verified in various cell lines and in KO models. Many studies show that Cldn-3 overexpression and silencing in various cell lines increases and decreases transepithelial resistance (TER), respectively, indicating a sealing role. Coyne et al. constructed a stable cell line in IB3.1 (human airway) cells overexpressing Cldn-3 and found increased TER and decreased solute permeability (Coyne et al., 2003). The same group also stably expressed Cldn-3 in NIH/3T3 fibroblasts, which normally do not form TJs, and noticed a slight

14 development of resistance across the cell layer and a reduction in paracellular permeability (Coyne et al., 2003). Interestingly, Cldn-3 seems to be impermeable to both charged and uncharged solutes in the MDCK II distal tubule cells and overexpression of Cldn-3 in these cells elevated TER (Milatz et al., 2010). This suggests that Cldn-3 is a general barrier-forming protein in a kidney epithelium model. TJ integrity was disrupted when Cldn-3 expression was reduced in the parotid acinar and Sertoli cells of the testis (McCabe et al., 2012; Yokoyama et al., 2017). Other studies also indicate an important role of Cldn-3 in maintaining the BBB and BTB, and in preventing solute permeability (Li et al., 2018; Stammler et al., 2016; Wang et al., 2019). In an elegant study by Peng et al. conducted in ARPE-19, a cell line that expresses all TJ proteins except claudins, exogenous Cldn-3 expression decreased permeability of small ions across the TJs and increased TER (Peng et al., 2016). This further confirms the role of Cldn-3 as a barrier-forming claudin.

Cldn-3 KO models also indicated the essential role of Cldn-3 in maintaining the barrier function in various organs. Cldn-3 KO resulted in impaired BBB in mice exhibiting experimental autoimmune encephalomyelitis (Kooij et al., 2014). Another group demonstrated that Cldn-3 KO mice developed gallstone disease and increased paracellular phosphate ion permeability across the hepatic TJs (Tanaka et al., 2018). This shows that in the hepatobiliary system, Cldn-3 plays a major role in maintaining a paracellular barrier for phosphate ions. Surprisingly, Cldn-3 KO mice also demonstrated increased sweat leakage, implicating its role in maintaining a water barrier function in the sweat gland (Yamaga et al., 2018). However, Cldn-3 was not shown to regulate transepithelial water permeability in the distal tubules (Milatz et al., 2010). Cldn-3 is also a key component of the gut TJs, as several studies have demonstrated Cldn-3 to maintain gut barrier integrity (Ahmad et al., 2017; Mees et al., 2009; Sikora et al., 2019).

Evidence for pore-forming functions In contrast to the findings suggesting a sealing role, few studies have demonstrated pore-forming functions of Cldn-3. Overexpression of Cldn-3 in alveolar epithelial cells decreased barrier function, as evident by reduced TER and increased paracellular ion flux (Mitchell et al., 2011). In

15 addition, Cldn-3 was shown to form a complex with Cldn-16 and -19, and together, these claudins were involved in the permeability and reabsorption of divalent cations in the thick ascending limb (Plain & Alexander, 2018). This indicates that Cldn-3 may be partially involved in regulating permeability when present in a complex with other claudins. Another study has shown that TNFα treatment increased TER in the colorectal cancer cell line, Caco-2, however, under the same conditions, decreased Cldn-3 protein expression was also observed (Gusti et al., 2014).

Overall, these studies show that the permeability functions of Cldn-3 are highly context- and cell- type dependent.

Role in proliferation The possible effect of Cldn-3 on cell proliferation is highly contradictory in different cell types as well. In early gastric cancer, Cldn-3 expression was found to be significantly lower at the submucosal invasive front. This was associated with increased proliferation of gastric cancer and may play a vital role in the progression of early gastric cancer (Okugawa et al., 2012). In contrast, stable transfection of Cldn-3 in mouse inner medullary collecting duct cells (mIMCD-3) was associated with a significant increase in nephric tubulogenesis as a result of increased cell proliferation. This also demonstrates a critical role of Cldn-3 in kidney development (Haddad et al., 2011).

Role in migration Various studies have demonstrated Cldn-3 overexpression to be associated with reduced migration in several cancers, although some studies demonstrated contradictory roles. In general, Cldn-3 overexpression reduced cancer cell migration, proliferation, invasion and epithelial-mesenchymal transition (EMT) in ovarian, lung, and colon cancers (Che et al., 2018; Li et al., 2012; Takehara et al., 2009).

In several studies conducted in cancer cell lines, Cldn-3 KD was shown to activate specific signaling pathways. For example, in ovarian cancer cells, Cldn-3 KD increased tumour growth and

EMT. The effect on growth was mediated via the E-cadherin, GSK-3β and β-catenin signaling pathway, while the effect on EMT required activation of the PI3K-Akt pathway (Lin et al., 2013; Shang et al., 2012). Cldn-3 KD was also associated with increased metastasis and apoptosis in esophageal cancer (Takala et al., 2007). It was also shown that TNFα- and TGFβ-induced downregulation of Cldn-3 mRNA expression induced EMT of breast cancer stem cells, further confirming the vital role of Cldn-3 in suppressing EMT of cancer cells (Asiedu et al., 2011). Together, these findings suggest that Cldn-3 plays a vital role in sustaining an epithelial phenotype and that its loss promotes EMT mediated by distinct pathways.

In contrast, some studies have associated Cldn-3 overexpression with increased malignant potential by modulation of EGF-activated MEK/ERK and PI3K-Akt signaling pathways in both lung and colorectal cancers (de Souza et al., 2013; Zhang et al., 2017). Another study has associated Cldn-3 overexpression in ovarian cancer cells with increased cell motility and invasion via increased matrix metalloproteinase-2 (MMP-2) activity (Agarwal et al., 2005). In addition, injecting a lipidoid-Cldn-3 siRNA resulted in suppressed ovarian tumour growth and metastasis in transgenic mice (Huang et al., 2009). This siRNA was also implicated as a promising therapeutic for ovarian cancer.

Overall, these reported differences in the Cldn-3-mediated regulation of cancer cell migration are not yet completely understood.

Role in epithelial maturation and polarity generation Cldn-3 was shown to affect polarity in several cell types. In liver development, during the formation of bile canaliculi, Cldn-3 was one of the first claudins to settle in the TJs. Cldn-3 silencing delayed bile canaliculi formation, suggesting its role in hepatocyte polarity (Grosse et al., 2013). Interestingly, CPE-mediated removal of Cldn-3 from neural tube TJs resulted in altered cell polarity, reduced cell intercalation and Rho/ROCK-mediated acto-myosin contraction, leading to neural tube defects (Baumholtz et al., 2017).

These complex patterns of Cldn-3 expression and functions reveal the importance of studying Cldn-3 regulation. Thus, in the next section, I will provide a detailed insight into the mechanisms involved in regulating Cldn-3 expression. c) Claudin-3 expression regulation Cldn-3 synthesis is regulated through complex signaling mechanisms in a highly context- dependant manner. Although, these pathways ultimately affect transcription factor-mediated regulation of Cldn-3, some pathways also affect localization, phosphorylation and turnover of Cldn-3. I will first discuss general Cldn-3 regulation, followed by an overview of specific stimuli regulating Cldn-3 expression and turnover.

General claudin-3 regulation Claudin-3 transcriptional regulation Cldn-3 transcription can be affected by a number of stimuli. The Cldn-3 promoter contains putative binding sites for the transcription factors NFκB, Sp1, Klf4/6, Snail1, Slug, GATA3/4, CREB, Cdx2, Egr1, ATF-1 and AP1. Many of these have significant roles in various pathologies as well.

Cdx transcription factors have significant functions in gut differentiation and carcinogenesis (Guo et al., 2004). Cdx2, specifically, was shown to upregulate Cldn-3 expression, suggesting a possible role in gastric carcinoma and intestinal metaplasia (Satake et al., 2008). Klf6 was shown to induce Cldn-3 promoter activity, leading to TJ remolding, BTB maintenance and increased proliferation of Sertoli cells (Wang et al., 2019). In Klf4 KO mice, Cldn-3 expression was downregulated, and the associated upregulation of Snail, Slug and Twist transcription factors contributed to increased EMT in corneal epithelial cells (Tiwari et al., 2017). GATA4 activation in MDCK II cells, and GATA3 and PPARy1 overexpression in buccal epithelial cells were both shown to induce Cldn-3 expression (Guillemot et al., 2013; Hustler et al., 2018). The transcription factor Sp1 was also shown to bind to the Cldn-3 promoter and is vital for its activation in ovarian cancer cells (Honda et al., 2007).

Several studies have shown that the transcriptional co-repressor, Snail1, directly regulates the Cldn-3 promoter as well. For example, a transcriptional factor complex comprised of Snail1 and Smad3/4 was found to decrease Cldn-3 promoter activity which promoted TGFβ-induced EMT of breast epithelial cells (Vincent et al., 2009).

The Cldn-3 promoter has also been found to be regulated by epigenetic processes. The methylation of the Cldn-3 promoter was found to reduce Cldn-3 expression and hypermethylation of the promoter was an independent predictor of poor prognosis in gastric adenocarcinoma and ovarian cancer (Honda et al., 2007; Zhang et al., 2018).

Claudin-3 mRNA stability and post-transcriptional regulation Cldn-3 expression was also found to be regulated by microRNAs at the post-transcriptional level. MiR-34b/c and miR200a-associated upregulation of Cldn-3 was shown to mediate EMT, polarity and differentiation in lung and mammary cells (Kim et al., 2019; Nagaoka et al., 2013). Additionally, silencing the long noncoding RNA SPRY4-IT1 decreased Cldn-3 mRNA stability, leading to reduced translation and dysfunctional intestinal epithelial barrier, suggesting its role in protecting the gut barrier (Xiao et al., 2016).

Post-translational modifications of the claudin-3 protein Cldn-3 protein is targeted by several post-translational modifications, such as phosphorylation, palmitoylation and glycosylation, which may play a key role in regulating its trafficking (see below). Cldn-3 undergoes palmitoylation at multiple cysteine sites (C-X-X-C or C-C) present on the intracellular loop and in the fourth transmembrane domain (Butt et al., 2012; Rodenburg et al., 2017). The close proximity of potential palmitoylation and phosphorylation sites may suggest a possible interplay between these types of modifications. Cldn-3 protein was also found to have O-glycosylation sites in the extracellular loop and the cytoplasmic tail (Butt et al., 2012). However, the implications of this modification are not yet known. Cldn-3 was also shown to be phosphorylated by protein kinase A (PKA), the implications of which will be discussed in the next section (D’Souza et al., 2005).

Claudin-3 trafficking and degradation Cldn-3 trafficking and degradation are mainly regulated by post-translational modifications. Several studies have shown phosphorylation of Cldn-3 to be vital for its trafficking and localization regulation. PKA-dependent phosphorylation of Cldn-3 at threonine 192 led to TJ disruption, which was evident by decreased TER and increased paracellular permeability in ovarian cancer cells (D’Souza et al., 2005). The decrease in Cldn-3 was attributed to decreased localization and TJ insertion of Cldn-3 in these cells, leading to its degradation. Calcitonin-induced PKA-dependent phosphorylation of Cldn-3 was also found to contribute to TJ disassembly in prostate cancer cells (Aljameeli et al., 2017). Another study has implicated phosphorylation of the tyrosine 214 residue of Cldn-3 in controlling Cldn-3 delivery to the TJs via vesicular trafficking from the cytosol (Twiss et al., 2013).

Many of the above described findings likely involved Cldn-3 endocytosis, although this was not directly indicated by these studies. However, one particular study has visualized Cldn-3 to be pinching off from the membrane into intracellular granular structures. Moreover, upon wounding, increased cell motility was associated with increased endocytosis of Cldn-3, suggesting Cldn-3 endocytosis is a vital process during TJ remodeling (Matsuda et al., 2004).

There is very little known regarding the mechanisms involved in basal Cldn-3 degradation. A study has shown that in mouse hepatocytes, β- and γ-catenin silencing resulted in decreased Cldn-3 expression, and this was rescued by inhibition of proteasomal degradation. This suggests that Cldn-3 may potentially be degraded by the proteasomes (Pradhan-Sundd et al., 2018). There is currently no evidence of lysosomal degradation of Cldn-3, however, lysosomes are known to be involved in degradation of other claudins (Amoozadeh et al., 2018; Flores-Maldonado et al., 2017).

Specific stimuli regulating claudin-3 Effect of growth factors and cytokines on claudin-3 A variety of growth factors were found to affect Cldn-3 expression. In MDCK II cells, epidermal growth factor receptor (EGFR) activation significantly increased Cldn-3 expression and insertion into the TJs, resulting in elevated TER. In contrast, heparin-binding-epidermal growth factor (HB- EGF) was found to downregulate Cldn-3 expression via upregulation of the transcription factor Snail in endometrial epithelial cells (Liang et al., 2013). Cldn-3 promoter activity was also supressed by Snail1 in a mitogen-activated protein kinase (MAPK) and Egr-1-dependent manner upon hepatocyte-growth factor (HGF) induction. This also led to increased EMT, migration and invasion of liver carcinoma cells (Grotegut et al., 2006). HGF was also shown to reduce Cldn-3 trafficking to the TJs in MDCK cells (Twiss et al., 2013). Among growth factor regulated pathways, Src and the p38 MAPK pathways were shown to downregulate Cldn-3 expression in the parotid gland, resulting in increased paracellular permeability (Fujita-Yoshigaki, 2011; Yokoyama et al., 2017).

Cytokines were also shown to affect Cldn-3 expression. TNFα was shown to decrease paracellular permeability via an ERK1/2/Slug-dependent downregulation of Cldn-3 expression in a submandibular gland cell line (Mei et al., 2015). IL-1β was also shown to downregulate Cldn-3 expression via β-catenin-mediated regulation of the Cldn-3 promoter, in part through non muscle myosin light chain kinase (nmMLCK) activation (Haines et al., 2016) The exact downstream effector of this pathway is not yet known. Combined IFNγ and TNFα treatment also led to decreased Cldn-3 expression, increased inflammation and paracellular permeability in colonic epithelial cells (Li et al., 2014; Prasad et al., 2005). Interleukins affect Cldn-3 via a variety of signaling pathways. IL-1β, IL-6 and IL-18 were all shown to suppress Cldn-3 expression in the amniotic membrane and in breast cancer cells, mediated by the p38 MAPK pathway (Kobayashi et al., 2010; Yang et al., 2015). IL-9 stimulation was also shown to decrease Cldn-3 expression in patients with ulcerative colitis through STAT3 phosphorylation (Tian et al., 2018).

Hormones regulating claudin-3 expression Glucagon-like peptide-2, a growth promoting hormone of the intestine, glucocorticoids and prolactin were shown to increase Cldn-3 expression and improve overall barrier function (Dong et al., 2014; Kobayashi et al., 2016).

Various sex hormones were also shown to augment Cldn-3 expression. Cldn-3 expression was upregulated by estradiol, progesterone, LH, FSH and testosterone (Kobayashi et al. 2011; Meng et al., 2005; Rimon et al., 2004; Someya et al., 2013). In contrast, Cldn-3 was downregulated by estrogen and gonadotrophin (Liang et al., 2013; Tarulli et al., 2008). These regulations had major implications on embryo implantation and BTB maintenance. In addition, dehydroepiandrosterone sulfate (DHEAS), a circulating steroid, was shown to increase Cldn-3 expression through an ERK1/2-mediated activation of the transcription factors ATF-1 and CREB1, resulting in increased TER in Sertoli cells (Papadopoulos et al., 2018).

Effect of drugs on claudin-3 Cldn-3 expression is also altered by several drugs. Dodecylmaltoside (DDM), a surfactant-like permeation enhancer which improves absorption of drugs in the gut, resulted in reduced Cldn-3 expression (Gradauer et al., 2017). In addition, methamphetamine treatment was also shown to decrease Cldn-3 expression in brain endothelial and uterine cells, respectively (Mahajan et al., 2008; Parikh et al., 2015). The Cldn-3 promoter also contains an epidermal growth response 1 (Egr1) binding site. Interestingly, a cell-permeable zinc chelator, TPEN, induced a decrease in the Cldn-3 promoter activity via Egr1 and this was associated with intestinal TJ disruption (Miyoshi et al., 2016). This is especially important because altered zinc levels often lead to pathogenesis of liver disease and inflammatory bowel diseases (Bartholomay et al., 1956; Vagianos et al., 2007).

The administration of a glycogen synthase kinase-3β (GSK-3β) inhibitor, lithium, was shown to increase Cldn-3 expression following intracerebral hemorrhage (ICH), thereby improving BBB integrity (Weishan Li et al., 2018). Artesunate, a popular anti-malaria drug, elevated Cldn-3

22 expression via sphingosine-1-phosphate-mediated PI3K activation in the BBB following subarachnoid hemorrhage, leading to increased BBB integrity (Zuo et al., 2017). Metformin was also shown to promote Cldn-3 expression via AMPK signaling in Caco-2 cells, and this correlated with improved intestinal barrier function (Xue et al., 2016). The cardiac glycoside, ouabain, induced Cldn-3 expression through an ERK1/2 signaling pathway and the functional implications of this effect were impaired TJ barrier function and increased cell viability and proliferation in Caco-2 cells (de Souza et al., 2014). This is contradictory to the previously described barrier- forming function of Cldn-3.

Effects of pathogens on claudin-3 Cldn-3 expression is also affected by various bacterial, fungal and viral pathogens. Importantly, Cldn-3 is one of the main receptors of the Clostridium perfringens enterotoxin (CPE), and many studies have shown that upon CPE binding, Cldn-3 is removed from the TJs, resulting in elevated permeability in MDCK I and ovarian cancer cells (Fujita et al., 2000; Sonoda et al., 1999; Yuan et al., 2009). Several pathogens including Toxoplasma gondii, Trichinella spiralis, Salmonella typhimurium and Citrobacter rodentium were shown to decrease Cldn-3 expression in the GI tract, enabling transmigration of these pathogens across the gut epithelium (Dalton et al., 2006; Fernández-Blanco, Estévez, Shea-Donohue, Martínez, & Vergara, 2015; Xiao Li et al., 2014). In TLR2 KO mice infected with C. rodentium, Cldn-3 was found to be delocalized from the membrane into the cytoplasm, where it was visualized as punctiform aggregates near the apical region of colonic crypts (Gibson et al., 2008; Guttman et al., 2006).

Some mycotoxins also decrease Cldn-3 expression and induce intestinal barrier dysfunction via activation of the MAPK and NFκB signaling pathways (Gao et al., 2017; Romero et al., 2016; Ying et al., 2019). The HIV-1 Tat protein was also shown to downregulate Cldn-3 expression in RPE cells via activation of ERK1/2 and NFκB signaling pathways, leading to increased paracellular permeability (Bai et al., 2008; Tan et al., 2014).

Several studies have also shown downregulation of Cldn-3 by the bacterial endotoxin lipopolysaccharide (LPS). LPS infection in pregnant mice decreased Cldn-3 expression and was associated with dysfunctional amniotic barrier, eventually leading to apoptosis of the amniotic epithelium (Kobayashi et al., 2010). LPS treatment was also shown to decrease Cldn-3 expression in the blood-biliary barrier and in the intestine of mice (Reiling et al., 2017). LPS was also shown to increase Cldn-3 phosphorylation, although the underlying mechanisms are unknown (Jing Li et al., 2015).

In contrast, treatment of cholera toxin was shown to increase Cldn-3 expression in rat colon, leading to increased TER (Markov et al., 2014).

Effect of cell stress on claudin-3 Various sources of physical cell stress were also shown to regulate Cldn-3 expression. Heat stress and hydrogen peroxide treatments were shown to downregulate Cldn-3 expression in RPE, intestinal and gastric epithelial cells (Hashimoto et al., 2008; Y. Li et al., 2010; Pearce et al., 2013). Another study demonstrated that the hydrogen peroxide-induced reduction in Cldn-3 trafficking to the TJs was mediated by an MLCK and Src kinase-dependent pathway in the bile duct epithelium (Guntaka et al., 2011). In contrast, hypoxia and ozone exposure upregulated Cldn-3 expression in the human umbilical cord vein and alveolar epithelial cells (Kim et al., 2018; Scheurer et al., 2004). d) Role of claudin-3 in diseases A growing number of studies have demonstrated dysregulation of Cldn-3 expression, phosphorylation and/or localization in various pathological conditions. These might be due to pathological changes in the activity of various signaling pathways that control Cldn-3 expression and trafficking. In this section, I will provide a brief overview of the evidence linking Cldn-3 to various cancers and inflammatory gastrointestinal diseases and discuss the signaling mechanisms involved in Cldn-3 dysregulation.

Role in cancer Multiple studies have found Cldn-3 to be highly expressed in various cancers including lung, liver, breast, ovarian, prostate, colorectal and gastric cancer (Bartholow et al., 2011; Blanchard et al., 2009; G. Chen et al., 2018; Coutinho-Camillo et al., 2011; Moldvay et al., 2007; Tokuhara et al., 2018; Wang et al., 2015). An increasing amount of literature correlated Cldn-3 overexpression with BRCA mutation-associated triple negative breast cancer (TNBC) as well (Danzinger et al., 2019; Masili-Oku et al., 2017; J. Xu et al., 2017). Generally, this Cldn-3 overexpression was associated with decreased tumour cell proliferation, migration, metastasis and EMT, although some contradictions do exist and this will be discussed below.

Cldn-3 KO mice were found to have a leaky and dedifferentiated colonic epithelium and these mice developed invasive adenocarcinoma. This was regulated by IL6/gp130/Stat3 signaling- mediated hyperactivation of the Wnt/β-catenin signaling pathway (Ahmad et al., 2017). This demonstrates the role of Cldn-3 in regulating colorectal cancer malignancy by suppressing further metastasis of cancer cells. Another study has shown that silencing Cldn-3 increased breast cancer cell migration (Yang et al., 2015). However, this correlation did not prove a causative connection. Interestingly, Cldn-3 is now used as a biomarker in predicting TNBC progression as well. Specifically, elevated levels of cytoplasmic Cldn-3 expression is a predictor of poor survival in TNBC patients (Jääskeläinen et al., 2018).

In contrast, in the colorectal cancer cell line, HT-29, Cldn-3 elevation was shown to contribute to increased cancer cell development and this upregulation was mediated by the SCF/c-kit/JNK signaling pathway. This led to activation of the transcription factor AP-1 which bound to and increased activity of the Cldn-3 promoter (Wang et al., 2017).

Role in gastrointestinal inflammatory diseases Cldn-3 is known to be a key regulator of the intestinal barrier and its dysregulation contributes to various gastrointestinal pathologies. Cldn-3 was found to be downregulated in inflammatory bowel disease (IBD) which includes celiac disease and Crohn’s disease, and in liver cirrhosis and

25 experimental colitis induced by dextran sulfate sodium (DSS) (Garcia-Hernandez et al., 2017; Goswami et al., 2014; Z. Wang et al., 2019; Xia et al., 2011; Yuan et al., 2015). This downregulation was associated with increased intestinal barrier dysfunction and paracellular permeability and enhanced bacterial invasion. In celiac disease, the impaired duodenal epithelial barrier function was attributed to intracellular Cldn-3 localization mediated by dysregulated expression of the cell polarity proteins Par-3 and serine/threonine phosphatase PP-1 (Schumann et al., 2012; Szakál et al., 2010). Cldn-3 expression was also reduced in the small intestine of mice with sickle-cell disease, and this correlated with increased TNFα levels and increased intestinal permeability. Interestingly, antibiotic treatment rescued the decreased Cldn-3 levels, leading to elevated intestinal barrier function (Tavakoli & Xiao, 2019). In addition, HIV infection in a macaque model also reduced Cldn-3 expression in the late stage of HIV infection, and this correlated with severe intestinal diseases (Croteau et al., 2017). Interestingly, cannabinoid-mediated increase in Cldn-3 expression was linked to reduced intestinal inflammation and maintenance of the barrier integrity in the same model (Kumar et al., 2019; Zhang et al., 2014).

Role in other diseases Interestingly, some studies have also demonstrated vascular effects of Cldn-3. Cldn-3 was selectively lost in the choroid plexus of patients with multiple sclerosis and in the brain endothelial cells in response to acute hypertension (Kooij et al., 2014b; Mohammadi & Dehghani, 2014). This led to increased arterial pressure and microvascular permeability in the BBB. Some studies also suggest a role for Cldn-3 in brain disorders. In patients with autism spectrum disorder, increased Cldn-3 expression was observed in the brain. This suggests a potential protective effect imparted by Cldn-3 on maintaining the BBB integrity (Fiorentino et al., 2016). Finally, Cldn-3 was also found to be increased in idiopathic pulmonary fibrosis (Lappi-Blanco et al., 2013).

Interestingly, some animal models also show that Cldn-3 might localize to the cytoplasm instead of the TJs in certain pathological conditions. During sepsis and intestinal ischemia-reperfusion

26 injury, Cldn-3 protein was delocalized from the TJs and was present diffusely within cells, leading to breakdown of TJ barrier integrity (Li et al., 2009).

1.5 Inflammatory Cytokines and TJs It is well-known that inflammation has a major effect on epithelial permeability. During inflammation, both immune and epithelial cells produce pro-inflammatory cytokines which are secreted into the circulation (Uluçkan & Wagner, 2017). While initially this facilitates the repair process, in the long-term, this may initiate a vicious cycle as these cytokines in turn target the epithelial cells and disrupt the TJs, ultimately leading to altered epithelial permeability. In kidney pathology, tubulointerstitial inflammation (TI) is an important initiator of these processes which eventually results in tubular destruction. One of the major inflammatory cytokines affecting the tubular epithelium is Tumour Necrosis Factor-α (TNFα).

In the following section, I will provide a brief overview of TNFα synthesis and its activation of various signaling pathways.

1.5.1 TNFα structure and synthesis TNFα is a pro-inflammatory cytokine synthesized as a 26 kDa transmembrane protein in response to inflammation, injury or infection. It is produced by immune cells (e.g. macrophages) and non- immune cells (epithelial cells) (Idriss & Naismith, 2000). Upon synthesis, the membrane-bound TNFα is cleaved by a metalloprotease, TNFα-converting enzyme (TACE), into a soluble TNF (sTNF) (Kalliolias & Ivashkiv, 2016). This resulting 17 kDa sTNF to circulate through the body and bind to receptors that are distant from its site of synthesis. (Van Zee et al., 1992).

TNFα induces a signaling cascade by binding to either TNF receptor 1 (TNFR1) or TNF receptor 2 (TNFR2) (Figure 1-3). TNFR1 is ubiquitously and constitutively expressed at low levels in most tissues, whereas, TNFR2 is an inducible and highly regulated receptor (Brenner et al. 2015). In the cytoplasm, the TNFα receptors are associated with various TNFα adaptor proteins, activating factors (TRAF) and receptor interacting proteins (RIP). These components activate different

27 signaling pathways which lead to diverse cellular responses including gene transcription, inflammation and cell death. Interestingly, TNFα was also shown to have pro-survival effects in several epithelia, which shows that the effect of TNFα varies according to cell types, leading to differential cell fate (Roy et al., 2017; Suominen et al., 2004). TNFR1 is comprised of a conserved death-domain (DD) motif which induces apoptosis, on the other hand, the lack of DD in TNFR2 prevents it from inducing programmed cell death directly (Grell et al., 1999).

1.5.2 TNF receptor activation and signaling pathways TNFα homotrimers bind to homotrimeric TNFR and initiate a signaling cascade. Activation of TNFR1 results in binding of the adaptor protein TNFα-receptor associated via death domain (TRADD) and recruitment of a protein complex including complexes I, IIa, IIb and IIc (Jixi et al., 2013). Binding of each complex leads to a different functional outcome. TNFα activates three classical pathways: Nuclear Factor Kappa B (NFκB) family, mitogen-activated protein kinase (MAPK) and apoptotic signaling (Leong & Karsan, 2000) (see Figure 1-3). The TNFR-associated adaptor proteins, TRAFs and RIPs play a crucial role in activating these pathways. It should be noted that TNFα signaling is quite complex as evidenced by crosstalk that occurs between these classical pathways and with various other signaling pathways.

For my thesis project, I studied the TNFα-induced activation of Cldn-3 expression mediated by ERK-dependent NFκB activation and PKA-dependent CREB1 activation. I will now provide a brief overview of the literature discussing TNFα-induced activation of these kinases and transcription factors.

As mentioned above, it is well-known that TNFα activates the MAPK signaling pathway, leading to activation of extracellular-signal-regulated kinase (ERK), JUN N-terminal kinase (JNK) and p38 MAPK. Specifically, the ERK MAPK are known to be regulated by TNFα-induced Raf/MEK activation, and affect various cellular processes including cell proliferation, inflammation and cell survival (Sabio & Davis, 2014; Winston et al., 1995). In search for the upstream mechanisms of ERK activation, our lab has previously found that TNFα induces transactivation of the epidermal

28 growth factor receptor (EGFR) through ADAM17 (TACE), which then activates the MEK/ERK pathway in the tubular epithelium (Kakiashvili et al., 2011). TNFα also activates the canonical NFκB signaling pathway by promoting dissociation of the NFκB subunits from its inhibitory complex (Hayden & Ghosh, 2014). TNFα-induced NFκB activation has been shown to regulate transcription of genes involved in inflammation, proliferation and cell survival (Kempe et al. 2005; Schütze et al., 1995).

Several studies have revealed potential crosstalk between the ERK and NFκB pathways, induced by either TNFα or other stimuli. One particular study has shown TNFα-induced ERK-dependent NFκB p65 phosphorylation at the serine 536 residue in epidermal cells (Hu et al., 2004). Other studies have also shown activation of NFκB through ERK, however, this was in response to other stimuli, such as LPS and IL-8 (Diomede et al., 2017; Wang et al., 2010).

It is well-known that cyclic AMP (cAMP) induces gene transcription through activation of cAMP- dependent protein kinase A (PKA), which then phosphorylates the transcription factor, cAMP response element-binding protein (CREB) on Ser133 (Delghandi et al., 2005). TNFα was shown to stimulate adenylyl cyclase activity and increase cAMP levels in human myometrial and rat mesangial cells, respectively (Baud et al., 1988; Gogarten et al., 2003). Interestingly, TNFα was also shown to activate CREB via the p38 signaling pathway in endothelial cells (Gustin et al., 2004). These findings imply that TNFα indeed activates PKA, although this has not been experimentally verified.

These studies reveal the complex crosstalk that exists between TNFα and several other signaling pathways.

TNFα TNFα

TNFR1 TNFR2

RIP1

TRAF2

TRADD TRADD

FADD

RIP1 TRAF2

- Pro

8 caspase IKK p38 JNK ERK complex (MAPK) Apoptosis

Proteasomal p65 degradation p50

p50 p65 AP -1 NFκB

Figure 1-3. Simplified TNFα receptor activation of classical pathways. Overview of classical TNFα signaling pathways induced by TNF receptor type 1 (TNFR1) and type 2 (TNFR2). TNFR1- associated death domain (DD) induces apoptosis through recruitment of TNFα receptor- associated death domain (TRADD), Fas-associated death domain (FADD) and pro-caspase 8. TNFR1 and TNFR2 also transduce inflammatory signals in association with the receptor interacting protein (RIP) and TNF receptor associated factor (TRAF). These receptors induce extracellular signal regulated kinase (ERK), p38 and NFκB activation pathways, which regulate various cellular responses. TNFR2 activates the c-Jun N-terminal kinase (JNK) pathway, leading to activator-protein 1 (AP1) activation.

TNFα

EGF

EGFR TNFR

MEK IKK complex cAMP

ERK

ERK IκB α

p65 p50 ? PKA ? p p65 p50

p p p50 p65 CREB

Figure 1-4. Simplified TNFα receptor crosstalk overview in epithelial cells. TNFR crosstalk with the epidermal growth factor receptor (EGFR) and potentially with the adenylyl cyclase (AC) pathway. TNFR transactivates the EGFR pathway leading to MEK and ERK activation in tubular epithelial cells. Whether ERK activates p65 NFκB in epithelial cells is unknown. It is also not clear whether TNFR activates AC in epithelial cells. AC activation is known to induce phosphorylation of cyclic AMP response element-binding protein (CREB) in a cAMP-dependent protein kinase A (PKA) manner. It is not known whether ERK activates CREB in endothelial cells. Dotted arrows indicate pathways that are currently unknown in epithelial cells.

1.5.3 Role of TNFα in kidney diseases TNFα was shown to play a key role during both acute and chronic kidney disease. In the tubules, many factors were shown to stimulate TNFα production, including reperfusion-injury, hypoxia, unilateral ureteral obstruction and LPS treatment (Donnahoo et al., 1999; Xuan Li et al., 2005; Misseri et al., 2004; Nozaki et al., 2017). In addition, increased synthesis and release of TNFα by infiltrating macrophages also contributes to tubulointerstitial inflammation. TNFα also plays a major role in diabetic nephropathy, ischemic-reperfusion injury, obstructive uropathy and glomerulonephritis (Durán, 2008; Navarro & Morafernandez, 2006; Stokes et al., 2005; Truong et al., 2011). Interestingly, inhibition of TNFα production by administration of TNFα-neutralizing antibodies or by knocking-out TNFR in mice resulted in alleviated kidney injury (S. B. Khan et al., 2005; Singh et al. 2013). TNFα also has a dual role in kidney injury and repair. In addition to its role in promoting kidney injury, it also contributes to kidney regeneration and acts as a pro- survival factor (Brilli et al., 2019; Lech et al., 2014; Yamashita & Passegué, 2019).

Although, the role of TNFα in mediating kidney injury has been characterized, the cellular mechanisms through which it affects kidney functions have not been fully defined. Therefore, my work aims to gain mechanistic insights into the TNFα-induced effects on the tubular epithelium.

CHAPTER 2: RESEARCH AIMS/OBJECTIVES AND HYPOTHESES

Rationale: It is well-known that inflammatory cytokines alter epithelial permeability and that they play a vital role in the dynamic regulation of TJs in several organs. Specifically, the pro-inflammatory cytokine, TNFα, has emerged as a key mediator and a pathogenic factor in various kidney diseases. Although it is known that the kidney tubules respond to inflammatory mediators such as TNFα, the cellular mechanisms through which TNFα affects various kidney functions remain incompletely understood.

Claudin-3 is a ubiquitously expressed TJ protein with important roles as a regulator of paracellular permeability, cell migration and proliferation. Claudin-3 is a highly regulated TJ protein which has also been shown to be altered in various pathologies. Our previous studies have demonstrated that TNFα caused major remodelling of the TJs and altered the expression of several claudins in tubular cells, with important functional consequences. However, the effect of TNFα on claudin-3 in the tubules remained unknown, and possible underlying mechanisms remained undefined.

Overall objective: The overall objective of my project was to gain an in-depth insight into the mechanisms that control claudin-3 expression in tubular epithelial cells. Specifically, I wished to define the effect of TNFα on claudin-3 expression and defined the signaling mechanisms involved in mediating this effect. I also investigated the functional role of claudin-3 expression in regulating epithelial permeability and cell proliferation.

Hypothesis: Stimulation by TNFα elevates Cldn-3 expression in tubular epithelial cells. This effect may occur due to altered Cldn-3 synthesis promoted by specific transcription factors and/or by reduced Cldn-3 degradation. I also hypothesize that this may have functional implications on tubular epithelial permeability and cell proliferation.

In order to test these hypotheses, the following specific aims and questions were defined.

Specific Aims: 1. Explore the effect on TNFα on claudin-3 expression in epithelial cells. 2. Define whether TNFα alters claudin-3 synthesis and/or degradation. 3. Explore the signaling mechanisms through which TNFα alters claudin-3 expression. 4. Define the functional role of claudin-3 expression on epithelial permeability and cell proliferation.

Aim 1: Explore the effect of TNFα on claudin-3 expression in epithelial cells. Questions: 1. How does short- or long-term TNFα treatment alter claudin-3 expression in kidney tubular epithelial cells? 2. Does TNFα treatment alter claudin-3 expression in other epithelial cell lines?

Aim 2: Define whether TNFα alters claudin-3 synthesis and/or degradation. Questions: 1. Does TNFα alter claudin-3 mRNA synthesis? 2. Is claudin-3 degradation affected by TNFα treatment?

Aim 3: Explore the signaling mechanisms through which TNFα alters Cldn-3 expression. Questions: 1. Do ERK and NFκB mediate the TNFα-induced increase in claudin-3 mRNA and protein expression? 2. Do PKA and CREB1 mediate the TNFα-induced increase in claudin-3 mRNA and protein expression? 3. Is there any crosstalk between the two pathways mediating claudin-3 expression?

Aim 4: Define the functional role of claudin-3 expression on epithelial permeability and cell proliferation. Questions: 1. Does claudin-3 upregulation have an effect on transepithelial resistance and epithelial permeability? 2. Does claudin-3 expression have an effect on cell cycle proteins?

CHAPTER 3: MATERIALS AND METHODS

Materials and antibodies: TNFα was obtained from Sigma-Aldrich Chemical Co. (St Louis, MO) and PeproTech (Montreal, QC, Canada). The effects of TNFα from both sources were identical. All experiments were conducted using TNFα from Sigma-Aldrich, unless otherwise indicated. Complete Mini Protease Inhibitor tablet was from Roche Diagnostics (Laval, QC, Canada). Bovine serum albumin (BSA) was from BioShop Canada (Burlington, ON, Canada). Further details on these reagents are presented in Table 1. The following inhibitors were used: GDC0094 (#21107) Bay11-7082 (#10010266), KT5720 (#10011011), and cycloheximide (#14126) from Cayman Chemical (Ann Harbor, MI), H-89 dihydrochloride (#371962) from Sigma-Aldrich (St Louis, MO). All inhibitors were prepared as a stock using DMSO and used at a 1:1000 dilution. This dilution of DMSO has no measurable effects. Forskolin (#11018) was from Cayman Chemical (Ann Harbor, MI) and 8-bromo-cAMP (#1140) was from Tocris Bioscience (Oakville, ON). All other chemicals were from Sigma or Bioshop. The following antibodies were used: for claudin-3, a polyclonal rabbit antibody (#341700) from Invitrogen (Thermo Fisher) was used for Western blotting and a polyclonal rabbit antibody (#AB52231) from Abcam was used for immunofluorescent staining. Antibodies against NFκB p65 phosphoserine 536 (#3033S), p65 (#4764), CREB (#9197S), phospho- CREB (#9198S) were from Cell Signaling Technologies (Danvers, MA); Snai2/Slug (#C353781), from LifeSpan Biosciences. NFκB p65 phosphoserine 276 (#194726) was from Abcam (Cambridge, MA). Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) (#sc-47724) and HA-probe antibodies were (#sc-805) from Santa Cruz Biotechnology (Dallas, TX). Peroxidase-coupled secondary antibodies were from Jackson ImmunoResearch (West Grove, PA). Alexa Fluor 488- labelled secondary antibody and 4',6-diamidino-2-phenylindole (Dapi) were from Thermo Fisher.

Cells: LLC-PK1, a kidney tubule epithelial cell line (male) with mostly proximal tubule characteristics from the European Collection of Animal Cell Cultures (Wiltshire, UK) was used as in our earlier studies (Amoozadeh et al., 2015). IMCD3, a mouse inner medullary collecting duct cell line, and Caco-2 (male), a human colorectal adenocarcinoma cell line, were from American Type Culture Collection (ATCC). All cell lines were maintained in DMEM medium supplemented with 10% fetal bovine serum and 1% penicillin/streptomycin in an atmosphere containing 5% 36

CO2. Tissue culture media and reagents were from Thermo Fisher/Life Technologies. pReceiver- M06-3xHA (#EX-M0354-M06) and pReceiver-M56-mCherry (#EX-M0354-M56) referred to as HA- Cldn-3 and Cherry-Cldn-3, respectively, were from GeneCopoeia. The HA-tag is fused to the N- terminus region of Cldn-3, whereas, the mCherry-tag is fused to the C-terminus of Cldn-3. Stable cell lines expressing HA-Cldn-3 or Cherry-Cldn-3 were generated by transfecting the HA- or

Cherry-Cldn-3 vectors into LLC-PK1 cells using FuGENE 6 (Promega Corporation, Madison, WI), according to the manufacturer’s instructions. Transfected cells expressing HA- or Cherry-Cldn-3 were selected by growing cells in 2mM G418 for 7 days.

Short interfering RNA (siRNA). Oligonucleotides were purchased from Thermo Fisher/Dharmacon. The following porcine sequences were targeted to silence various proteins: Cldn-3 siRNA: CCUACGACCGCAAGGACUAUUUU; CREB: siRNA#1: GGAUAAAGAUAGAUGGGAA, and siRNA#2: CAUACUAGCUGGUUGGAUU. Data obtained using the two CREB siRNAs were similar and were pooled. Cells were transfected with 100 nM siRNA oligonucleotide using the Lipofectamine™ RNAiMAX Transfection Reagent (Thermo Fisher/Invitrogen) according to the manufacturer’s instructions. Control cells were transfected with 100 nM Silencer siRNA negative control (non-related (NR) siRNA) (Applied Biosystems/Ambion). Unless otherwise indicated, experiments were performed 48 hours after transfection. Downregulation of silenced proteins was routinely verified using Western blotting.

Western blotting: Following treatment, cells were lysed on ice using cold lysis buffer containing 100 mM NaCl, 30mM HEPES pH 7.5, 20 mM NaF, 1 mM EGTA, 1% Triton X-100, supplemented with 1 mM Na3VO4, 1 mM PMSF, and protease inhibitors (Roche Diagnostics, Laval, QC). For the detection of phosphorylated proteins, the lysis buffer was also supplemented with PhosSTOP phosphatase inhibitor (Roche Diagnostics, Laval, QC). Protein concentration was determined by the bicinchoninic acid assay (Thermo Fisher/ Pierce Biotechnology) with bovine serum albumin used as a standard. Equal amounts of protein were run on SDS-PAGE using 10% or 15% gels at 130V. Proteins were transferred to nitrocellulose membranes using Western blotting (110V for 90 min). The SDS-PAGE and Western blotting was performed using the Bio-Rad

37 system. Membranes were blocked in Tris-buffered saline (TBS) containing 3% BSA and incubated with the primary antibody overnight at 4°C. Unless otherwise indicated, antibodies were used at 1:1000 dilution. The following day, membranes were washed with TBS-T and incubated with the corresponding anti-rabbit or anti-mouse horseradish peroxidase (HRP) conjugated secondary antibody for 1h at room temperature. The membranes were washed with TBS-T again and antibodies were visualized using the enhanced chemiluminescence method (kit from Bio-Rad). GAPDH (1:10000) was used to demonstrate equal loading. Blots were either stripped and redeveloped or were horizontally cut following transfer, and the corresponding parts were simultaneously developed with specific antibodies. Enhanced chemiluminescence (ECL) signal was captured using a BioRad ChemiDoc Imaging system and densitometry was performed using ImageLab (BioRad).

Microscopy: LLC-PK1 cells were grown on coverslips. For immunofluorescence, cells were ﬁxed with ice-cold methanol, washed with phosphate-buffered saline (PBS) and blocked with 3% BSA in PBS. Following fixation, the coverslips were incubated with primary antibody (1:50) for 1h, then washed with PBS and bound antibody was detected using Alexa Fluor488 labelled secondary antibody (1:1000). Nuclei were counterstained with Dapi. Unless otherwise indicated, Z-stacks were obtained using a Zeiss LSM700 confocal microscopy system (60x objective). Original z-stack files were imported into the Imaris 8.0.2 software package to perform 3D modeling and analysis of Cldn-3 staining. Using the Imaris surface creation module, the Cldn-3 surface was defined based on an absolute intensity threshold of 12 and a surface area detail of 0.15 μm. Total surface volumes (μm3) were obtained. All acquisition parameters for image acquisition and 3D analysis were kept constant.

RT-PCR for mRNA analysis: LLC-PK1 cells were treated as indicated. RNA was extracted using the RNeasy kit (Qiagen, Valencia, CA), and cDNA was synthesized from 1 µg total RNA using iScript reverse transcriptase (Bio-Rad Laboratories). SYBR green-based real-time PCR was performed using the Quant Studio 7 real-time PCR system. cDNA was denatured at 95°C for 30 sec, followed by 40 cycles of 95°C for 15 sec and then 60°C for 30 sec. Primer pairs designed

38 against the corresponding porcine sequences were as follows: Cldn-3: 5’- GTCCATGGGCCTGGAGAT-3’ and 5’-GATCTGCGCTGTGATAATGC-3’; and GAPDH: 5’GCAAAGTGGACATGGTCGCCATCA-3’ and 5’-AGCTTCCCATTCTCAGCCTTGACT-3’. Relative gene expression of Cldn-3 was compared using the comparative CT method using GAPDH as the reference standard.

Electric cell-substrate impedance sensing (ECIS): The ECIS Ztheta system (Applied Biophysics, Troy, NY) was used to follow transepithelial resistance during development of a confluent layer (Amoozadeh et al., 2015). LLC-PK1 expressing either HA-tagged-Cldn-3 or mCherry-Cldn-3 were seeded on 8W2LE electrode arrays at 3 x 105cells/well in 400 μl culture medium and resistance (R) and capacitance (C) values were collected continuously for 45h at 4000 Hz and 32000 Hz, respectively.

Statistical analysis. All blots and immunoﬂuorescent pictures are representative of at least three similar experiments. Data are presented as means ± SD of the number of experiments indicated (n). Statistical signiﬁcance was assessed by one-way ANOVA performed using the GraphPad Prism software. Unless otherwise indicated, Tukey’s post-hoc test was used. For comparison of two conditions, Student’s t-test (unpaired, equal variance) was performed.

Table 3-1. List of inhibitor and activator reagents

Targeted Reagent name Mode of action References pathway Protein Inhibition of translation Baskić et al., 2006; Cycloheximide synthesis elongation Obrig et al., 1971 Selective inhibition of GDC0994 ERK Blake et al., 2016 ERK1/2 Selective and irreversible Karin, 1999; Pahl, 1999; Bay11-7082 NFκB inhibition of IKBα Pierce et al., 1997 phosphorylation Selective inhibition of IκB IKK16 NFκB kinase (IKK) and IκB Baumgartner et al., 2017 degradation Davies et al., 2000; De Reversible, ATP-competitive H-89 PKA Rooij et al., 1998; inhibitor of PKA Leemhuis et al., 2002 Selective, ATP-competitive Davies et al., 2000; Kase et KT-5720 PKA inhibitor of PKA al., 1987; Murray, 2008 Adenylyl Direct activator of adenylyl Insel & Ostrom, 2003; Forskolin cyclase cyclase Robbins et al., 1996 Cell-permeable cAMP Carranza et al., 1998; 8-bromo-cAMP PKA analog involved in Chow & Wang, 2002; Hei activating PKA et al., 1991

CHAPTER 4: TNFα-induced claudin-3 expression regulation in kidney tubular epithelial cells

Results

4.1 TNFα induced an increase in Cldn-3 protein expression We have previously shown that TNFα exerted a time-dependent effect on the expression of claudin-1, -2 and -7 in tubular cells (Amoozadeh et al., 2017). Whether TNFα also affected Cldn- 3 remained unknown. To fill this gap, we first explored the effect of TNFα on Cldn-3 expression in kidney tubular epithelial cells. LLC-PK1 tubular epithelial cells were exposed to 20 ng/ml TNFα for 3, 16, 24 or 48h and Cldn-3 protein was detected by Western blotting. Cldn-3 protein levels were quantified and normalized to the housekeeping protein GAPDH. As shown in Fig 4-1A, a significant increase in Cldn-3 protein expression was detectable as early as 3h after the addition of TNFα and Cldn-3 levels continued to increase up to 48h treatment. Surprisingly, the Cldn-3 band consistently appeared lower (around 18 kDa) than the expected molecular weight (around 23 kDa). Therefore, we verified the specificity of the Cldn-3 antibody by using a porcine Cldn-3- specific siRNA. Indeed, this resulted in marked downregulation of the 18 kDa Cldn-3 band (Fig 4- 1B). Since commercially available TNFα shows some variability, we also verified the effect using TNFα from another source. As seen in Fig 4-1A and 1C, the effect of TNFα from two different sources were similar. Based on these introductory findings, throughout the next experiments we used 24h TNFα treatment. The effect of TNFα on elevated Cldn-3 levels was also verified using immunofluorescent staining. In control cells, Cldn-3 showed a cobblestone-like staining typical of TJ localized proteins (Fig 4-1D). In addition, Cldn-3 was also detectable in intracellular vesicular structures. In cells exposed to TNFα, both the membrane-localized and intracellular Cldn-3 staining was significantly stronger. In line with the data obtained using western blotting, quantification of the signal revealed close to two-fold increase in the staining.

Next, we wished to establish that the effect of TNFα on Cldn-3 is not unique to LLC-PK1 cells. As shown in Fig 4-1E, TNFα induced a significant elevation in Cldn-3 protein levels in the collecting duct cell line, IMCD3, and in the colorectal cancer cells, Caco-2. Taken together, our results show that TNFα can upregulate Cldn-3 protein expression at the TJs, and that the effect is detectable in a variety of epithelial cells.

Figure 4-1. TNFα increased Cldn-3 protein expression in tubular epithelial cells

Sigma TNF A 6 ** Con Con 16h 24h 3

3h Con Con 48h - 5

n ) d 20 l **

d l

C 4

o f

d (

Cldn-3

e s

z 3 **

i l

17 e

a * v

e 2

m l

r o GAPDH 35 N 1 0 3h 16h 24h 48h

Cldn-3 TNF

NR siRNA 25 B 20

Cldn-3 17

GAPDH 35

C PeproTech TNF 2.5 p<0.0001

2.0 3

Con TNF -

)

d l

20 l 1.5

(

Cldn-3 e s

z i

e 1.0

17 l

e l

m r

o 0.5 GAPDH 35 N 0.0 Con 24h TNF

10 D Control TNF

10 μm

10 μm 10 μm Figure 4-1. TNFα increased Cldn-3 protein expression in other epithelial cells

IMCD3 cells 4.0 E p<0.0001 3.5

Con TNF 3.0

3 -

) d

d 2.5 l

20 l

( d

2.0 e

Cldn-3 s

i e

l 1.5 v

17 a

l m

r 1.0

o N GAPDH 35 0.5 0.0 Con TNF

Caco-2 cells 3.5 p<0.0001 Con TNF 3.0

3 2.5

20 -

)

d l

l 2.0 o

Cldn-3 C

(

s l

17 z 1.5 i

e l

m 1.0

r o GAPDH 35 N 0.5 0.0 Con TNF

Figure 4-1. (A) Confluent LLC-PK1 cells were treated with 20 ng/ml TNFα from Sigma-Aldrich for the indicated times. Cells were lysed and Cldn-3 and the loading control protein GAPDH were

detected by Western blotting. Normalized Cldn-3 levels were expressed as fold change from their time-matched controls, set as 1. Graph shows mean ±SD, n=4, *p<0.01, **p<0.0001. (B)

LLC-PK1 cells were transfected with nonrelated (NR) siRNA or Cldn-3 specific siRNA. Forty-eight hours later, the cells were lysed and Cldn-3 levels were detected and quantified. The graph

shows mean ±SD, n=4. (C) Confluent LLC-PK1 cells were treated with 40 ng/ml TNFα from PeproTech for 24h. Cells were lysed and Cldn-3 levels were detected and quantified. The graph

shows mean ±SD, n=5. (D) LLC-PK1 cells grown on coverslips were treated with 20 ng/ml TNFα for 24h, then washed and fixed with methanol. Cldn-3 was visualized using an antibody from

Abcam and an Alexa488 labelled secondary antibody. Nuclei were counter-stained with Dapi. Z- stacks were obtained using a Zeiss LSM700 confocal microscope and 3D modeling and analysis

3 was performed using Imaris 8.0.2. The graph shows total Cldn-3 surface volume (μm ) per cell ±SD, n=8. (E) IMCD3 cells and Caco-2 cells were treated with TNFα for 24h, and Cldn-3 levels were detected and quantified as above. The graph shows mean ±SD, n=3. Scale bar represents

10 μm.

4.2 TNFα did not alter Cldn-3 degradation and increased Cldn-3 mRNA levels We have previously shown that short-term (3h) TNFα treatment increased the stability of Cldn- 2, thereby elevating its expression (Amoozadeh et al., 2015). To explore the effects of TNFα on Cldn-3 degradation, we treated cells with the protein synthesis inhibitor cycloheximide (CHX) and detected changes in Cldn-3 levels. Cldn-3 levels significantly dropped after 16h CHX treatment (Fig 4-2A). While TNFα alone elevated Cldn-3 expression, this effect was no longer detected in the presence of CHX, suggesting that TNFα did not rescue Cldn-3 from degradation.

Having demonstrated that TNFα did not alter Cldn-3 degradation, we next explored the effect of

TNFα on mRNA levels. Cldn-3 mRNA levels in untreated LLC-PK1 cells and in cells treated with TNFα were determined using quantitative RT-PCR. Cldn-3 mRNA levels were significantly elevated in TNFα-treated cells (Fig 4-2B), suggesting that this effect occurs at the transcriptional level.

Figure 4-2. TNFα did not affect Cldn-3 degradation

and increased Cldn-3 mRNA levels

p<0.0001

p<0.01 A CHX p<0.001

CHX 2.5

Con TNF Con TNF 3

- 2.0 n

20 )

kDa o 1.5

Cldn-3 f

(

s z

17 l

i l

e 1.0

a v

kDa e

l r

GAPDH o 0.5

35 N

kDa 0.0 Con TNF Con TNF

CHX

B p<0.0001 23

20 19

(∆Ct) levels Cldn-3 mRNA 18 Control TNF

Figure 4-2. (A) Confluent LLC-PK1 cells were treated with 100 μM CHX and 20 ng/ml TNFα for

16h. At the end of the treatment, cells were lysed and Cldn-3 levels were detected and quantified as in Fig 3-1. The graph shows mean ±SD, n=4. (B) LLC-PK1 cells were treated with 20 ng/ml TNFα for 24h and mRNA levels were determined using SYBR green based real-time PCR with GAPDH as the normalizing control. The graph shows mean delta Ct values ±SD, n=9.

4.3 TNFα-induced increase in Cldn-3 expression is mediated by NFκB and CREB1 but not Slug Having shown that TNFα induced upregulation of Cldn-3 mRNA, we next wished to determine the transcription factors involved in mediating the effects of TNFα. Interestingly, a previous study conducted in submandibular cells showed that TNFα induced Cldn-3 downregulation through an ERK-dependent upregulation of the inhibitory transcription factor Slug (Mei et al., 2015). This is contradictory to our findings where we found elevated Cldn-3 expression upon TNFα stimulation. Therefore, it was conceivable that in tubular cells, TNFα might regulate Slug differently. To explore this possibility, we first assessed the status of the ERK/Slug pathway in our tubular cell model. Surprisingly, we found that TNFα increased Slug expression, although this effect was not found to be ERK dependent since the ERK inhibitor GDC0994 did not prevent this effect (Fig 4- 3A). Since Slug is an inhibitor of Cldn-3, its upregulation by TNFα is unlikely to account for the observed effect, and in fact, it may mitigate the effect on Cldn-3. Moreover, we also found that ERK inhibition strongly mitigated the increase in Cldn-3 expression induced by TNFα (Fig 4-3A and see below).

In search for alternative mechanisms, we performed an in-silico analysis of the Cldn-3 promoter (Gene ID: 431781) using Transcription Factor Site Scan. Among other binding sites, this analysis revealed the Cldn-3 promoter to have potential NFκB and cAMP Response Element (CRE) binding sites spanning -200 bp to -211 bp and -424 bp to -431 bp, respectively, relative to the transcription start site (Fig 4-3B). Although the role of NFκB in claudin regulation has been previously assessed, the role of CRE in Cldn-3 regulation remains largely unknown. Therefore, in the next series of experiments we sought to define the role of NFκB p65 (RelA) and CREB1 transcription factors in mediating the effects of TNFα on Cldn-3 expression. Firstly, we focused on the possible role of NFκB p65 since TNFα is a strong NFκB activator. We used two different inhibitors of NFκB (Bay11-7082 or IKK16), that have distinct modes of action. Indeed, both inhibitors prevented the TNFα-induced increase in Cldn-3 protein (Fig 4-3C and 4-3D) and mRNA (Fig 4-3E) expression. To assess the role of CREB1, we used a porcine CREB1-specific siRNA. Indeed, this downregulation reduced CREB1 levels by approximately 70%. Importantly, CREB1 silencing also strongly mitigated the TNFα-induced increase in Cldn-3 levels (Fig 4-3F). Together,

46 these data suggest that TNFα induces an upregulation in Cldn-3 expression through the transcription factors NFκB and CREB1.

Figure 4-3. TNFα-induced increase in Cldn-3 required NFκB and CREB1

A GDC0994

Con TNF Con TNF 35 Slug 25 20 Cldn-3 17

GAPDH 35

ns p<0.01 ns p<0.01 4.0 3.0 p<0.01 3.5

3 2.5

n )

u 3.0 d

)

d l

S 2.0

2.5 o

f f

(

z 1.5

2.0 s

l e

a v

v 1.5 m

e 1.0

l r

o 1.0

o N

N 0.5 0.5 0.0 0.0 Con TNF Con TNF Con TNF Con TNF

GDC0994 GDC0994

-431 -424 -390 -381 -211 -200 +1 B CRE Sp1 NFκB

Figure 4-3. (A) Confluent LLC-PK1 cells were treated with 50 nM GDC0994 for 15 min, followed by addition of 20 ng/ml TNFα for 24h in the presence of the inhibitor. At the end of the treatment, Slug and Cldn-3 levels were detected and quantified as above. The graph shows mean ±SD, n=3. (B) Schematic representation of NFκB and CREB1 transcription factor binding sites in the Cldn-3 promoter of porcine. +1, putative transcription start site.

Figure 4-3. TNFα-induced increase in Cldn-3 required NFκB and CREB1 p<0.0001 p<0.05 C Bay11-7082 p<0.0001 3.0

Con TNF Con TNF 3

- 2.5

20 n

)

l d

l 2.0 C

Cldn-3 o

(

e 1.5 s

17 z

a v

e 1.0

l r GAPDH 35 o N 0.5

0.0 Con TNF Con TNF Bay11-7082

p<0.01 IKK16 D p<0.01 Con TNF Con TNF 3.0

20 3 2.5

)

d l

Cldn-3 d 2.0

17 f

d (

e 1.5

l e

a v

e 1.0

m l

GAPDH 35 r

o N 0.5 0.0 p<0.0001 Con TNF Con TNF

E p<0.0001 IKK16 24 p<0.0001

20 18 16

(∆Ct) levels Cldn-3 mRNA 14 Con TNF Con TNF Bay11-7082 p<0.0001 F NR siRNA p<0.0001 CREB siRNA 4.0

Con TNF Con TNF 3.5 3 - 3.0

20 n

)

d l

l 2.5

Cldn-3 o

( d

2.0 e

17 s z

l e

a 1.5 v

48 e

l m r 1.0

CREB1 o

N 35 0.5 0.0 GAPDH 35 Con TNF Con TNF NR siRNA CREB siRNA 49

Figure 4-3. Confluent LLC-PK1 cells were treated with 20 μM Bay11-7082 (C) or 10 μM IKK16

(D) for 30 min, followed by addition of 20 ng/ml TNFα for 16h in the presence of the inhibitor. At the end of the treatment, Cldn-3 levels were detected and quantified as above. The graph shows mean ±SD, n=3. (E) LLC-PK1 cells were treated with TNFα for 24h with or without 20 μM Bay11-708, and Cldn-3 mRNA levels were determined as above. The graph shows mean delta

Ct values ±SD, n=9. (F) LLC-PK1 cells were transfected with nonrelated (NR) siRNA or CREB1 specific siRNA #1 and #2. Forty-eight hours later, the cells were treated with 20 ng/ml TNF for

24h and Cldn-3 levels were detected and quantified. The graph shows mean ±SD, n=3.

4.4 TNFα-induced increase in Cldn-3 requires ERK-mediated NFκB activation We next wanted to define the signaling pathways mediating the TNFα-induced activation of these transcription factors. We have previously shown that TNFα activates ERK through transactivation of the EGFR (Kakiashvili et al., 2011) and this pathway has been implicated in the control of Cldn- 2 expression (Amoozadeh et al., 2015). Moreover, ERK was suggested to control both Cldn-3 expression and activation of NFκB (Hu et al., 2004; Mei et al., 2015). Therefore, we first wanted to establish the role of ERK in TNFα-induced NFκB activation. TNFα induced phosphorylation of p65 on two sites, Ser536 (Fig 4-4A) and Ser276 (Fig 4-4B) that are known to enhance its activity (Mattioli et al., 2004; Okazaki et al., 2003). Phosphorylation on both sites was mitigated by the ERK inhibitor GDC0994, suggesting that the TNFα-induced NFκB activation is mediated by ERK. Next, we asked whether ERK was necessary for Cldn-3 upregulation. In line with data shown in Fig 4-3A, TNFα-induced upregulation of Cldn-3 protein (Fig 4-4C) was prevented by ERK inhibition. Moreover, Cldn-3 mRNA upregulation was also found to be ERK-dependent (Fig 4-4D).

ERK has also been shown to induce CREB1 phosphorylation (Gustin et al., 2004), therefore we next asked if ERK might affect Cldn-3 through this pathway as well. We found that ERK inhibition did not alter CREB1 phosphorylation (Fig 4-4E), suggesting that ERK does not mediate the TNFα- induced CREB1 activation. Together, these data suggest that ERK mediated TNFα-induced Cldn-3 upregulation through NFκB but not CREB1 activation.

Figure 4-4. TNFα-induced increase in Cldn-3 is mediated by ERK and NFκB

p<0.0001 A GDC0994 p<0.001 3.0 Con TNF Con TNF

5 2.5

6 p 75 ) /

p-p65 ) d

l 2.0

o 3

63 f (

(S536) 5

S 1.5

( l

75 e

v 6

e 1.0 p p65 l

63 - h

p 0.5

0.0 Con TNF Con TNF GDC0994

p<0.01 B p<0.01 GDC0994 4.0 3.5 5

Con TNF Con TNF 6 p

) 3.0

/ )

d l 75 6

p-p65 o 2.5

f 2 63 (

s 2.0

(S276) (

e 5

v 1.5

p l

75 - 1.0

p65 h

63 p 0.5 0.0 Con TNF Con TNF GDC0994

C p<0.01 GDC0994 p<0.01 3.0 Con

Con TNF TNF 2.5 3 -

20 n

)

d l d 2.0

Cldn-3 l

f (

17 d

e 1.5

e a

v 1.0

l r

GAPDH 35 o

N 0.5

0.0 Con TNF Con TNF

GDC0994

Figure 4-4. TNFα-induced increase in Cldn-3 is mediated by ERK and NFκB

D p<0.0001 p<0.0001 24 p<0.0001 22 20 18

Cldn-3 mRNA levels (∆Ct) levels Cldn-3 mRNA 12 Con TNF Con TNF

GDC0994

E GDC0994 ns p<0.01 3.0 Con TNF Con TNF 2.5

48 B )

d R

l 2.0 o

p-CREB C

/ f

(

s 1.5 E

35 l e

C -

e 1.0 l

48 h

p CREB1 0.5

35 0.0 Con TNF Con TNF GDC0994

Figure 4-4. Confluent LLC-PK1 cells were treated with 50 nM GDC0994 for 15 min, followed by addition of 20 ng/ml TNFα for 15 min in the presence of the inhibitor. At the end of the treatment, cells were lysed and total p65 and phospho-p65 (S276) (A) and phospho-p65 (S536)

(B) or Cldn-3 (C) were detected and quantified. The graphs show mean ±SD, n=4. The density of the phospho-proteins was normalized using total p65 levels. The graphs show mean ±SD, n=3. (D) LLC-PK1 cells were treated with 50 nM GDC0994 for 15 min, followed by addition of 20 ng/ml TNFα for 24h in the presence of the inhibitor, and Cldn-3 mRNA was determined as above. The graph shows means ±SD, n=9. (E) Confluent LLC-PK1 cells were treated with 50 nM GDC0994 for 15 min, and 20 ng/ml TNFα for 15 min. Cells were lysed and ph-CREB and total

CREB1 were detected and quantified. The graph show mean ±SD, n=3.

4.5 TNFα-induced elevation in Cldn-3 requires PKA-dependent CREB1 activation CRE sites can bind cAMP response element-binding protein (CREB), and cAMP-dependent PKA is a well-established activator of CREB1, which induces its activation by phosphorylation on serine 133 (Delghandi et al., 2005). Therefore, we next explored the role of PKA in mediating the effect of TNFα on Cldn-3 expression. We found that TNFα induced CREB1 phosphorylation and this phosphorylation was mitigated upon PKA inhibition (Fig 4-5A). Furthermore, inhibition of PKA by two different inhibitors, H-89 or KT5720, prevented the increase in Cldn-3 protein levels (Fig 4- 5A and 5B) and mRNA levels (Fig 4-5C) induced by TNFα stimulation. This suggests that PKA mediates the TNFα-induced CREB1 phosphorylation, and that this pathway is involved in regulating Cldn-3 expression.

Activating CREB1 alone is not sufficient to increase Cldn-3 levels Finally, we asked whether activating CREB1 alone could elevate Cldn-3 levels. Therefore, we used pharmacological approaches to elevate cAMP levels and thereby promote CREB1 phosphorylation. Treatment with the phosphodiesterase inhibitor, forskolin, along with the cell permeable cAMP analogue 8-bromo-cAMP is known to induce a strong increase in cAMP levels. Indeed, these agents increased CREB1 phosphorylation (Fig 4-5D). Despite this, they failed to augment Cldn-3 expression. These data suggest that while CREB1 is a necessary mediator of the TNFα-induced Cldn-3 increase, its activation alone is not sufficient to elevate Cldn-3.

We also wanted to explore potential crosstalk between the two signaling pathways shown to regulate Cldn-3 expression. As shown in Fig. 4-5E, PKA inhibition did not alter TNFα-induced NFκB p65 phosphorylation levels, suggesting that PKA does not regulate NFκB activation. Taken together, this shows that there is no crosstalk between the ERK-mediated NFκB activation and the PKA-mediated CREB1 activation pathways, and that these are distinct pathways involved in regulating the TNFα-induced Cldn-3 upregulation. It is also likely that both pathways are needed to mediate this effect.

Figure 4-5. TNFα-induced increase in Cldn-3 required PKA and CREB1

H-89 A Con TNF Con TNF 20 Cldn-3 17

GAPDH 35 48 ph-CREB 35 48 CREB1 35

p<0.01 p<0.0001 p<0.05 p<0.001 2.5 2.0

2.0 3

- 1.5

)

l l

R 1.5 C

(

1.0

l E

l e

e 1.0

m h

r 0.5 p 0.5 o N 0.0 0.0 Con TNF Con TNF Con TNF Con TNF

H-89 H-89 B p<0.001 p<0.01 KT5720 3.0

Con TNF Con TNF 2.5 3

n )

20 d 2.0

l C

Cldn-3 ( d 1.5

17 e

z l

e a

v 1.0

e m l 35 r GAPDH o N 0.5 0.0 Con TNF Con TNF C p<0.0001 KT5720 p<0.0001 24 p<0.0001 22

18 16

(∆Ct) levels Cldn-3 mRNA 14 Con TNF Con TNF H-89 55

Figure 4-5. TNFα-induced increase in Cldn-3 required PKA and CREB

ns D FSK and 8- 2.0

Con br-cAMP

3 1.5

)

d l

20 l

C o

( d

Cldn-3 1.0

i e

17 l

l m

r 0.5 GAPDH o 35 N 0.0 Con FSK + 8-br-cAMP

p<0.05 FSK and 8- 6

Con br-cAMP

B )

E 4 d

48 l

/ (

ph-CREB

s l

e R

35 v

C 2

- l h 48 p CREB 35 0 Con FSK + 8-br-cAMP

E ns H-89 p<0.05 2.5

Con TNF Con TNF 2.0

) p

75 /

d )

p-p65 l 1.5

( 5

63 s

(S536) S

(

e 1.0

6 e

l p 75 - h 0.5 p65 63 p 0.0 Con TNF Con TNF H-89

Figure 4-5. (A) Confluent LLC-PK1 cells were treated with 1 μM H-89 (A) or 1 μM KT-5720 (B) for

30 min, and 20 ng/ml TNFα for 24h. At the end of the treatment, cells were lysed and ph-CREB, CREB and Cldn-3 levels were detected and quantified. The graph shows mean ±SD, n=3-4. (C)

LLC-PK1 cells were treated with 1 μM H-89 for 30 min, and 20 ng/ml TNFα for 24h, and Cldn-3 mRNA was determined as above. The graph shows mean ±SD, n=9. (D) Confluent LLC-PK1 cells were treated with 10 μM forskolin and 100 μM 8-bromo-cAMP for 24h or 30 min. Cells were lysed and ph-CREB, CREB and Cldn-3 levels were detected and quantified. The graphs show means ±SD, n=3. (E) Confluent LLC-PK1 cells were treated with 1 μM H-89 for 30 min, and 20 ng/ml TNFα for 15 min. Cells were lysed and phospho-p65 (S536) levels were detected and quantified. The graph shows mean ±SD, n=3.

4.6 Sp1 regulates TNFα-induced Cldn-3 upregulation The Cldn-3 promoter was also predicted to have an Sp1 binding site. In order to assess the role of Sp1 in mediating the TNFα-induced increase in Cldn-3, we used a porcine Sp1-specific siRNA to silence Sp1. Indeed, this downregulation reduced Sp1 levels by approximately 80%. Importantly, Sp1 silencing also strongly mitigated the TNFα-induced increase in Cldn-3 levels (Fig 4-6A). This suggests that Sp1 is also necessary for the TNFα induced upregulation of Cldn-3 expression.

Figure 4-6. Sp1 regulates TNFα-induced increase in Cldn-3

A NR siRNA Sp1 siRNA p<0.05 p<0.01 Con 10 TNF Con TNF

20 3

- 8

n )

Cldn-3 d

l d

17 C

o 6

d (

i l

100 e 4 a

Sp1 v e

75 m l r o 2 N GAPDH 35 0 Con TNF Con TNF NR siRNA SP1 siRNA

Figure 4-6. (A) LLC-PK1 cells were transfected with nonrelated (NR) siRNA or Sp1 specific siRNA.

Forty-eight hours later, the cells were treated with 20 ng/ml TNF for 24h, and Sp1 and Cldn-3 levels were detected and quantified. The graph shows mean ±SD, n=3.

4.7 Cldn-3 overexpression increased transepithelial resistance The exact permeability role of Cldn-3 is unclear due to differences in the experimental findings in various cell lines and this demonstrates that the permeability functions of Cldn-3 are highly context- and cell-type dependent. Moreover, the permeability properties of Cldn-3 in the tubules remains largely unknown. Therefore, we tested the effect of Cldn-3 overexpression in LLC-PK1 cells. We generated cell lines overexpressing Cldn-3 with an N-terminal HA-tag or a C-terminal mCherry-tag (Fig. 4-7A). As expected, immunofluorescence showed that the Cldn-3 protein localized to the TJs which correlates with the localization of endogenous Cldn-3. This also demonstrates that the tags did not interfere with correct localization of Cldn-3. Approximately 80% of the cells expressed the HA-tagged Cldn-3, whereas, approximately 30% of cells expressed the mCherry-tag.

In order to study the effect of Cldn-3 overexpression on epithelial permeability, we used Electrical Cell-Substrate Impedance Sensing (ECIS), as earlier (Amoozadeh et al., 2015). ECIS provides continuous and real-time measurements of specific parameters of a cell layer, including capacitance (C) and resistance (R), upon exposure to alternating currents at different frequencies. R measured at low frequencies indicates the TER of the layer. LLC-PK1 cells overexpressing Cldn-3 with either the HA-tag or the mCherry-tag were grown on filters and monitored for up to 45h. Typically, during the development of a confluent layer, R initially increases for 24h, followed by a slow decline, and eventual stabilization by 30-48h (Amoozadeh et al., 2015). We found that in cells overexpressing HA-tagged Cldn-3 (Fig. 4-7B) or mCherry- tagged Cldn-3 (Fig. 4-7C), the TER reached a higher point compared to WT LLC-PK1 cells. This increase in TER in Cldn-3-overexpressing cells suggests that Cldn-3 indeed seals off the tight junctions and therefore functions as a sealing claudin in these tubular cells.

Figure 4-7. Cldn-3 overexpression increased TER

A WT LLC-PK1 HA-tagged Cldn-3 mCherry-tagged Cldn-3

B 20000

15000

HA-Cldn-3 10000 Control

5000

0 Hz (Ohm) 4000 at Resistance 0 10 20 30 40 50 Time (h)

C 10000 8000 mCherry- 6000 Cldn-3

4000 Control 2000

0 0 10 20 30

Hz (Ohm) 4000 at Resistance Time (h)

Figure 4-7. (A) Immunofluorescent images of stable cell lines expressing N-terminal HA- or C- terminal mCherry-tagged Cldn-3 in LLC-PK1 cells. Cells were grown on coverslips and fixed with methanol. HA-tag was detected using an HA probe antibody from Santa Cruz Biotechnology and an Alexa488 labelled secondary antibody. Nuclei were counter-stained with Dapi. Slides were visualized using an Olympus IX81 microscope coupled to an Evolution QEi Monochrome camera. Scale bar represents 10 μm. Transepithelial resistance (TER) measurements in WT LLC- PK1, HA-tagged-Cldn-3 (B) and mCherry-Cldn-3 (C) cells during the development of a confluent

5 layer. TER of LLC-PK1 cells (3x10 cells/well) grown in wells of 8W2LE array (B) or 8W1E array (C) was followed using electric cell-substrate impedance sensing (ECIS). Capacitance and resistance (R) were monitored for 45h (B) and 25h (C). The graph depicts the normalized R values at 4000 Hz that reflect TER.

4.8 Cldn-3 silencing upregulates the cell cycle protein p27kip1 It has recently started to emerge that various claudins can affect signaling, likely through their cytoplasmic interactions. Accordingly, changes in the expression of various claudins have been shown to affect a variety of vital cell functions, such as proliferation and differentiation. Cell cycle progression is controlled by cyclin-dependent kinases which are regulated by cyclins and cyclin- dependant kinase inhibitor (CDKI) (S. Lim & Kaldis, 2013). We have recently shown that Cldn-2 silencing reduced cell proliferation through the upregulation of p27kip1, a member of the CDKI family of inhibitors which blocks the cell cycle progression from G1 to S phase, resulting in reduced proliferation (Dan et al., 2019). The possible effect of Cldn-3 on cell proliferation is highly contradictory in different cell types. Therefore, in an exploratory study, I silenced Cldn-3 using an siRNA and tested the effects on p27kip1 expression in tubular cells. I found that Cldn-3 silencing elevated p27kip1 levels (Fig 4-8A), suggesting that Cldn-3 may indeed affect cell proliferation in kidney tubular cells.

Figure 4-8. Cldn-3 regulates cell cycle protein p27kip1

A Cldn-3 p<0.001 3.0 NR siRNA 2.5

1 p

25 i

p27-kip1 )

k 2.0 -

(

1.5 d

20 s

i v

Cldn-3 l 1.0

l m

17 r o 0.5 N GAPDH 35 0.0 NR Cldn-3 siRNA

Figure 4-8. (A) LLC-PK1 cells were transfected with nonrelated (NR) siRNA or Cldn-3 specific siRNA. Forty-eight hours later, the cells were treated with 20 ng/ml TNF for 24h, and p27kip1 and Cldn-3 levels were detected and quantified. The graph shows mean ±SD, n=3.

CHAPTER 5: GENERAL DISCUSSION, FUTURE DIRECTIONS AND OVERALL CONCLUSIONS

5.1 General Discussion The overall objective of my thesis was to gain an in-depth understanding of the effect of TNFα on Cldn-3 expression in kidney tubular epithelial cells and the underlying mechanisms involved in this regulation. I was also interested in studying the functional role of Cldn-3 in epithelial permeability and cell proliferation.

In this study, we demonstrated that short- and long-term TNFα treatment increased Cldn-3 expression in tubular epithelial cells. TNFα was shown to upregulate Cldn-3 synthesis and had no effect on Cldn-3 degradation. This effect was mediated by two distinct mechanisms: TNFα- induced ERK-dependent NFκB activation and PKA-dependent CREB1 activation. Additionally, Cldn-3 overexpression increased transepithelial resistance (TER). Further, silencing Cldn-3 affected a cell cycle regulator protein, suggesting that it may affect cell proliferation through the control of cell cycle proteins. These main findings will be further discussed below.

In previous studies, we have shown that TNFα altered the expression of several claudins, including Cldn-1, -2, -4 and -7 (Amoozadeh et al., 2017). However, the effect of TNFα on Cldn-3 expression in the tubules remained unknown. To fill this gap, we first demonstrated TNFα- induced Cldn-3 upregulation in tubular cells. This was detected as early as 3h after TNFα addition, and protein levels continued to rise up to 48h treatment. This effect was not exclusive to tubular cells, as we found similar upregulation of Cldn-3 in other epithelial cell lines, including Caco-2 and IMCD3 cells. This suggests that this effect is a more general response to TNFα in various epithelial cells. However, this effect is contradictory to the findings of Mei et al., where they reported a downregulation of Cldn-3 expression in response to long-term TNFα in SMG-6, a submandibular gland cell line. Since commercially available TNFα shows some variability, we first verified the effect we found using TNFα from two different sources. Although the PeproTech TNFα proved less potent than the Sigma TNFα, the effect on Cldn-3 was similar, resulting in an increase induced

64 by both TNFα preparations. Importantly, the Cldn-3 band appeared much lower than the expected molecular weight of 23 kDa. The identity of the detected band was verified using siRNA- mediated silencing of Cldn-3. Whether the observed Cldn-3 band represents an alternatively spliced variant or a protein degradation product remains to be established. Additionally, we observed increased Cldn-3 mRNA levels upon TNFα treatment, further confirming this effect and suggesting that it is due to increased synthesis of the protein. Thus, a likely explanation for this discrepancy could be largely due to cell type-specific differences. Of note, we verified TNFα- induced upregulation of Cldn-3 in three different cells, suggesting that the effect is not specific to LLC-PK1 cells. The literature contains few reports on the possible effects of TNFα on Cldn-3. In support of our data, recently it was reported that in patients with liver cirrhosis, increased Cldn- 3 levels were positively correlated with increased TNFα levels (Wang et al., 2019). In contrast, combined treatment with TNFα and IFNγ or TNFα and TGFβ decreased Cldn-3 expression in colonic epithelial and breast cancer stem cells (Asiedu et al., 2011; Prasad et al., 2005). These findings are similar to what was reported in the above-mentioned study in submandibular cells.

Of note, TNFα is not the only stimulus that can alter Cldn-3 expression. As detailed in the introduction section, Cldn-3 is a highly regulated protein and various stimuli can upregulate its expression, including pathogens, drugs, amino acids and hormones.

Elevated levels of a protein might be due to an increase in synthesis, or reduction in degradation. We have previously shown that short-term (3h) TNFα treatment increased the stability of Cldn- 2, thereby elevating its expression (Amoozadeh et al., 2015). The Cldn-3 degradation pathways, however, are not well-defined. In order to gain further insight into Cldn-3 degradation, we blocked protein synthesis and measured Cldn-3 levels. We found that Cldn-3 levels started decreasing after 16h cycloheximide treatment. We also found that TNFα did not mitigate the cycloheximide-induced Cldn-3 decrease, suggesting that TNFα does not stabilize the protein, and has no effect on Cldn-3 degradation. This is in contrast to our previous findings, where we showed that TNFα reduced Cldn-2 degradation. However, by the end of the 16h cycloheximide treatment, some cell death was observed, limiting the conclusiveness of the experiment. Indeed,

65 cycloheximide and TNFα are known to induce apoptosis in various cell lines. Therefore, these experiments need to be performed at an earlier time point in order to conclusively rule out the effect of TNFα on degradation.

In addition to membrane-localized Cldn-3, immunofluorescent staining revealed Cldn-3 to be present in some intracellular vesicular structures as well. Some of our preliminary studies indicate that in unstimulated cells, Cldn-3 co-localized with the lysosomal protein, LAMP, (not shown) suggesting that Cldn-3 might be degraded through the lysosomes in tubular cells. However, whether inhibiting lysosomal degradation alters Cldn-3 levels and whether TNFα induces further alterations in its degradation need to be investigated.

Since TNFα did not affect Cldn-3 degradation, we then explored whether TNFα altered Cldn-3 synthesis. Indeed, we found that TNFα elevated Cldn-3 mRNA levels. We have previously shown that TNFα induced Cldn-1, -4 and -7 mRNA expression in tubular cells (Amoozadeh et al., 2017). In addition, others have shown increased Cldn-1 mRNA expression in lung, gastric and colorectal cancer cells in response to TNFα (Bhat et al. 2016; Shiozaki et al., 2012, 2014). In contrast, TNFα was shown to decrease Cldn-1 expression in the parotid gland cell line and in T-84 cells. This shows that the effect of TNFα on claudin expression is highly context- and tissue-dependent.

Since TNFα was shown to upregulate Cldn-3 mRNA levels, we sought to determine the role of transcription factors involved in mediating this effect. Several studies have reported the snail family of zinc-finger transcription factors, including Snail1 (SNAI1) and Slug (SNAI2), to be directly involved in regulating the Cldn-3 promoter. These transcriptional repressors decreased Cldn-3 promoter activity, thereby promoting EMT, migration and invasion of breast epithelial and liver carcinoma cells (Vincent et al., 2009; Grotegut et al., 2006). Mei et al. demonstrated that the TNFα-induced Cldn-3 downregulation in SMG-6 cells was mediated by an ERK1/2-dependent Slug activation (Mei et al., 2015). In our case, it was conceivable that a reduction in Slug might have mediated the TNFα-induced increase in Cldn-3. Hence, we determined the status of the ERK/Slug pathway in tubular cells. We found that TNFα induced an increase in Slug expression, which

66 would have resulted in reduced Cldn-3 expression, however, we observed an upregulation of Cldn-3. This suggests that TNFα may activate other transcription factors which stimulate Cldn-3 synthesis and that these effects are dominant over the repressive effects that Slug may induce. Surprisingly, we also found that TNFα-induced Slug activation was not ERK-dependent, while Cldn-3 upregulation was mediated by ERK. Based on these results, it seems that Slug is unlikely to mediate the TNFα-induced increase in Cldn-3 levels in tubular cells.

In search for other transcription factors that may be involved in regulating Cldn-3 expression, we focused on NFκB and CREB1 which were predicted to have binding sites on the Cldn-3 promoter. We have previously shown that TNFα activates ERK through transactivation of the EGF receptor, and this pathway has been implicated in the control of Cldn-2 expression (Kakiashvili et al., 2011). Also, other studies have shown that ERK mediates NFκB activation in response to TNFα as well as other stimuli such as LPS and IL-8 (Hu et al., 2004; Diomede et al., 2017; Wang et al., 2010). In addition, both ERK and NFκB have been shown to regulate the expression of several claudins (Amasheh et al., 2010; Hichino et al., 2017; Lipschutz et al., 2005). Moreover, the role CREB in regulating claudin expression is largely unknown. Indeed, we found that TNFα-induced ERK- dependent NFκB activation was involved in Cldn-3 upregulation. The effect of ERK-mediated regulation of Cldn-3 is also in disagreement to the findings of Mei et al., where they suggested an ERK1/2-dependent downregulation of Cldn-3 expression. This contradiction may be in part due to differences in the cell lines and tissue origin used.

In addition, we found that TNFα-induced PKA-dependent CREB1 activation was also involved in Cldn-3 upregulation. Several studies have found that TNFα stimulates adenylyl cyclase, thereby elevating cAMP levels (Delghandi et al., 2005; Wang et al., 2018). These findings imply that TNFα activates PKA, although this has not been experimentally verified. In addition to the canonical cAMP-PKA-CREB pathway, ERK may also activate CREB. This can occur either through the classical RAS-MEK-ERK pathway or through cAMP-EPAC-ERK pathway (Naqvi et al., 2014). Therefore, we tested for potential crosstalk between the two pathways. However, we found that ERK does not alter CREB1 phosphorylation. In addition, we found that PKA is not involved in NFκB activation,

67 suggesting that these are two distinct pathways involved in regulating the TNFα-induced Cldn-3 upregulation. Nevertheless, both of these pathways appear to be necessary for the upregulation of Cldn-3 by TNFα. In fact, many stimuli are known to simultaneously activate both of these pathways, suggesting that this may be a mechanism for inducing strong Cldn-3 upregulation.

Interestingly, we also found that CREB1 activation via treatment with cAMP analogues, that activate PKA and promote CREB1 phosphorylation, alone did not induce Cldn-3 upregulation. This suggests that although CREB1 is necessary for mediating TNFα-induced Cldn-3 increase, its activation alone is not sufficient to elevate Cldn-3.

It would be interesting to explore whether activation of the ERK/NFκB pathway by itself is sufficient for Cldn-3 expression in the absence of CREB1 activation. However, there were some challenges involved in doing so. I tried to activate the NFκB pathway without also activating PKA. Many stimuli affecting claudins (e.g. LPS and IL1-β) are known to activate both pathways. Therefore, I tried both genetic and pharmacological approaches, however, these were not successful. Firstly, NFκB activation through IKBα silencing deemed unsuccessful due to a fast turnover of the IKBα protein which resulted in overcoming of the downregulation. Second, a chemical compound, betulinic acid that has been shown to activate NFκB in other cells, failed to do so in tubular cells. Thus, we currently cannot conclude whether the NFκB pathway alone (in the absence of CREB1 activation) can upregulate Cldn-3.

The Cldn-3 promoter also contains binding sites for additional transcription sites, including Sp1, GATA-3/4, HGF, cdx homeodomain proteins, AP1 and Klf4/6. I found that Sp1 was also necessary for the TNFα-induced activation of Cldn-3. Interestingly, some studies have shown Sp1 to be regulated in an ERK-dependent manner, while other studies have suggested NFκB activity modulation by a synergistic interaction with Sp1 (Hirano et al., 1998; Savickiene et al., 2004). Further studies will have to clarify whether ERK is involved in activating Sp1 in TNFα treated tubular cells and whether this is upstream of NFκB activation or whether an Sp1-NFκB interaction modulates Cldn-3 expression. In addition, mutational analysis of a luciferase-coupled Cldn-3

68 promoter reporter construct can performed in order to determine whether these transcription factors bind directly to the promoter or whether they indirectly regulate the promoter through other factors and complexes. The interaction between the transcription factors and their direct binding to the endogenous Cldn-3 promoter should be verified. This can be done by performing a ChIP assay. Moreover, it would be interesting to explore the regulation of other transcription factors in response to TNFα as well.

The permeability functions of Cldn-3 are highly debated as many studies have reported contradictory findings in different cell lines. Moreover, the permeability properties of Cldn-3 in the tubules remains largely unknown. We hypothesized that if Cldn-3 is indeed functioning as a sealing protein in the tubules, its overexpression should result in an increase in TER, indicating a tightening of the cell layer. Thus, to assess the functional implications of Cldn-3 expression on permeability, we studied the effect of Cldn-3 overexpression on the TER in tubular cells which models the TNFα-induced upregulation of Cldn-3. We found that Cldn-3 overexpression increased TER in both Cldn-3 overexpressing cell lines, suggesting that in our tubular cell model, Cldn-3 seals off the TJs, thereby reducing paracellular permeability. As mentioned earlier, the exact role of the permeability functions of Cldn-3 are not definitive as many studies have provided conflicting results. Few studies conducted in Caco-2 and alveolar epithelial cells have shown Cldn-3 to be a pore-forming protein. However, the majority of studies have implicated the role of Cldn-3 as a barrier-forming TJ protein, as evidenced by increased TER and decreased solute permeability in MDCK II, parotid gland and brain endothelial cells. Nonetheless, the evidence that Cldn-3 is impermeable to both charged and uncharged solutes in MDCK II cells suggests that Cldn- 3 is a general barrier-forming protein in a kidney epithelium model.

We have previously shown that TNFα induced a biphasic change in transepithelial resistance (TER), with a decrease followed by an increase in TER (Amoozadeh et al., 2015). However, the role of Cldn-3 in mediating this change remained unknown. Based on my findings, it is possible that the late TNFα-induced increase in TER may be a consequence of the elevated Cldn-3 expression during this long-term TNFα treatment phase. This suggests that TNFα-induced

69 increase in Cldn-3 expression may contribute to the permeability changes induced by this cytokine. Our next step will be to determine if silencing Cldn-3 alters the effect of TNFα on permeability, as we have done earlier for Cldn-2 (Amoozadeh et al., 2015). Many observations demonstrate changes in Cldn-3 expression in highly proliferating cancer cells. Indeed, many claudins have been implicated in affecting proliferation. Thus, in addition to the effects on paracellular permeability, Cldn-3 may also play a role in modulating cell proliferation. We found that Cldn-3 silencing elevated p27kip1 levels, indicating suppressed cell proliferation. This can be further verified in the Cldn-3 overexpressed cells that we have generated. If Cldn-3 overexpression decreases p27kip1 levels, then Cldn-3 may indeed play a role in regulating cell proliferation. Cldn-3 overexpression has been shown to increase nephric tubulogenesis during early kidney development (Haddad et al., 2011). This is in line with my observation which suggested that Cldn-3 may promote proliferation. Thus, this implies that the overexpression of Cldn-3 in various cancers could also promote proliferation. However, the effect of Cldn-3 on proliferation is not straight-forward, since there are contradictory findings which suggest the opposite effect. Cldn-3 downregulation has been associated with increased proliferation of gastric cancer cells, indicating a vital role of Cldn-3 in the progression of early gastric cancer (Okugawa et al., 2012). Together, these studies reveal the importance of Cldn-3 in regulating cell proliferation in a highly context-dependent manner. However, whether Cldn-3 indeed reduces cell proliferation in tubular cells remains to be established. Similarly, it is not yet known whether TNFα-induced upregulation of Cldn-3 might have an opposite effect, contributing to augmented proliferation. We have previously described that Cldn-2 silencing reduced TNFα-induced proliferation in a GEF-H1/RhoA dependent manner (Dan et al., 2019). However, the effect of TNFα-induced upregulation of Cldn-3 on tubular cell proliferation and the underlying mechanisms need to be further explored.

In addition, some of our initial studies have revealed the role of Cldn-3 in cell migration as well. We observed that Cldn-3 silencing reduced cell migration in tubular cells (not shown). Thus, Cldn- 3 overexpression should promote migration, which would be in line with elevated Cldn-3 observed in some cancers. However, there are contradictions as the vast majority of studies have

70 attributed Cldn-3 expression to suppress migration in various cancer cells. This contradiction may be in part due to cell-specific effects. The Cldn-3-regulated reduction in migration, EMT and cancer metastasis indicates a protective effect that Cldn-3 imparts in these cells. On the other hand, in the tubules, Cldn-3 may play a vital role in migration and development of cells following wounding or tubular injury. This is also evident in our preliminary studies (not shown) using the mouse unilateral ureteral obstruction (UUO) model of kidney injury, where Cldn-3 was found to be upregulated at day 3 of injury. However, the molecular mechanisms involved in this regulation remain unknown.

Figure 5-1. Proposed mechanisms involved in regulating claudin-3 expression in tubular cells. Schematic representation of the mechanisms involved in TNFα-induced Cldn-3 upregulation in kidney tubular epithelial cells. TNFα increased Cldn-3 synthesis through two distinct mechanisms: ERK-dependent NFκB activation and PKA-dependent CREB1 activation. TNFα induced phosphorylation of p65 on two sites, serine 536 and serine 276, mediated by ERK. TNFα-induced increase in Cldn-3 synthesis resulted in increased TJ localization and increased transepithelial resistance (TER). Cldn-3 may be involved in regulating cell proliferation and migration. Preliminary findings also show lysosomes to be involved in degrading Cldn-3.

5.2 Future Directions My research findings have uncovered several interesting questions that can be further investigated and are discussed below.

I have shown that TNFα upregulates Cldn-3 mRNA and protein expression through NFκB and CREB1, however, it is not known whether these transcription factors directly interact with the Cldn-3 promoter. For the endogenous promoter, this can be investigated by performing a ChIP assay. A luciferase-based promoter assay can also be performed to assess the effect of TNFα stimulation using a Cldn-3 promoter with mutations in the transcription factor binding sites. This will further confirm the regulation of Cldn-3 gene transcription by TNFα.

Additionally, the general mechanism of Cldn-3 degradation remains largely unknown and this should be investigated in the context of the tubules. Although I have shown that TNFα did not alter Cldn-3 degradation, we obtained initial data suggesting that Cldn-3 is degraded in the lysosomes, however, whether TNFα alters its degradation needs to be further investigated.

In addition, post-translational modifications have important roles in the control of claudin stability. However, very little is known regarding the importance of phosphorylation on Cldn-3 stability in the tubules. Some studies have implicated Cldn-3 phosphorylation to be involved in Cldn-3 delocalization from the TJs. However, this needs to be verified in the context of the tubules.

Moreover, the functional consequences of TNFα-induced Cldn-3 upregulation should be further explored. We showed that Cldn-3 overexpression increased TER; however, in order to further confirm its role on permeability, Cldn-3 silencing experiments should also be conducted. This can be done via siRNA-mediated silencing of Cldn-3 and the effect should be compared with control cells. If Cldn-3 silencing decreases TER, this would further confirm Cldn-3 to be a barrier-forming protein in the tubular cell model. In addition, the permeability function of Cldn-3 can be further investigated by measuring the paracellular flux of ions in cells where Cldn-3 is either overexpressed or knocked-down.

Finally, additional functional assays need to be conducted in order to validate the functional role of Cldn-3 in the tubules, and to determine the importance of the TNFα-induced upregulation of Cldn-3 on permeability and wound healing. Furthermore, although my study has elucidated some of the molecular mechanisms involved in Cldn-3 regulation in a kidney tubular cell model, these effects need to be validated in vivo to further assess its significance in kidney injury, regeneration and disease. This can be done by TNFα injection into mice and analyzing Cldn-3 expression in kidney tissues and assessing solute permeability across the nephron. In addition, the effect of Cldn-3 KO on tubular function and development, under both basal conditions and in kidney injury models, should also be further studied. It would also be interesting to determine whether the TNFα-induced upregulation of Cldn-3 expression imparts a protective effect in a kidney injury model.

5.3 Conclusions In summary, my study provided novel insights into the signaling mechanisms involved in Cldn-3 expression regulation in response to TNFα in tubular epithelial cells. Particularly, I have shown that TNFα induces an upregulation of Cldn-3 mRNA and protein expression. I also confirmed this effect using TNFα from different sources and verified this effect in other epithelial cell lines. I have also shown that this effect occurs at the synthesis level and that TNFα does not alter Cldn- 3 degradation. I also identified two distinct signaling pathways involved in mediating TNFα- induced upregulation of Cldn-3. TNFα induced activation of the transcription factors NFκB and CREB1 through ERK- and PKA-dependent mechanisms, respectively, and both pathways were needed to mediate this effect. Cldn-3 overexpression increased TER, suggesting Cldn-3 might be a tight junction sealing protein in tubular cells. Finally, I showed that Cldn-3 silencing increased p27kip1, which can lead to reduced cell proliferation.

Taken together, my study reveals the mechanism of Cldn-3 regulation in response to a pro- inflammatory cytokine and its contribution to tubular barrier function. Cldn-3 may also play a vital role in modulating other key cellular events, such as proliferation and migration, which may have implications in various pathological conditions, including cancer and inflammation.

REFERENCES

Agarwal, R., D’Souza, T., & Morin, P. J. (2005). Claudin-3 and Claudin-4 Expression in Ovarian Epithelial Cells Enhances Invasion and Is Associated with Increased Matrix Metalloproteinase-2 Activity. Cancer Research, 65(16), 7378–7385. https://doi.org/10.1158/0008-5472.CAN-05-1036 Ahmad, R., Kumar, B., Chen, Z., Chen, X., Müller, D., Lele, S. M., … Singh, A. B. (2017). Loss of claudin-3 expression induces IL6/gp130/Stat3 signaling to promote colon cancer malignancy by hyperactivating Wnt/β-catenin signaling. Oncogene, 36(47), 6592–6604. https://doi.org/10.1038/onc.2017.259 Aljameeli, A., Thakkar, A., & Shah, G. (2017). Calcitonin receptor increases invasion of prostate cancer cells by recruiting zonula occludens-1 and promoting PKA-mediated TJ disassembly. Cellular Signalling, 36, 1–13. https://doi.org/10.1016/j.cellsig.2017.04.008 Amasheh, M., Fromm, A., Krug, S. M., Amasheh, S., Andres, S., Zeitz, M., … Schulzke, J.-D. (2010). TNFα-induced and berberine-antagonized tight junction barrier impairment via tyrosine kinase, Akt and NFκB signaling. Journal of Cell Science, 123(23), 4145–4155. https://doi.org/10.1242/jcs.070896 Amasheh, S. (2002). Claudin-2 expression induces cation-selective channels in tight junctions of epithelial cells. Journal of Cell Science, 115(24), 4969–4976. https://doi.org/10.1242/jcs.00165 Amoozadeh, Y., Anwer, S., Dan, Q., Venugopal, S., Shi, Y., Branchard, E., … Szászi, K. (2018). Cell confluence regulates claudin-2 expression: possible role for ZO-1 and Rac. American Journal of Physiology-Cell Physiology, 314(3), C366–C378. https://doi.org/10.1152/ajpcell.00234.2017 Amoozadeh, Y., Dan, Q., Anwer, S., Huang, H. H., Barbieri, V., Waheed, F., … Szászi, K. (2017). Tumor Necrosis Factor-α Increases Claudin-1, 4, and 7 Expression in Tubular Cells: Role in Permeability Changes. Journal of Cellular Physiology, 232(8), 2210–2220. https://doi.org/10.1002/jcp.25736 Amoozadeh, Y., Dan, Q., Xiao, J., Waheed, F., & Szászi, K. (2015). Tumor necrosis factor-α induces a biphasic change in claudin-2 expression in tubular epithelial cells: role in barrier functions. American Journal of Physiology-Cell Physiology, 309(1), C38–C50. https://doi.org/10.1152/ajpcell.00388.2014 Anderson, J. M. (2001). Molecular structure of tight junctions and their role in epithelial transport. News in Physiological Sciences : An International Journal of Physiology Produced Jointly by the International Union of Physiological Sciences and the American Physiological Society, 16, 126–130. Retrieved from http://www.ncbi.nlm.nih.gov/pubmed/11443232 Anderson, J. M., & Van Itallie, C. M. (2009). Physiology and Function of the Tight Junction. Cold Spring Harbor Perspectives in Biology, 1(2), a002584–a002584. https://doi.org/10.1101/cshperspect.a002584 Asaka, M., Hirase, T., Hashimoto-Komatsu, A., & Node, K. (2011). Rab5a-mediated localization of claudin-1 is regulated by proteasomes in endothelial cells. American Journal of Physiology. Cell Physiology, 300(1), C87-96. https://doi.org/10.1152/ajpcell.00565.2010 Asiedu, M. K., Ingle, J. N., Behrens, M. D., Radisky, D. C., & Knutson, K. L. (2011).

TGFbeta/TNF(alpha)-mediated epithelial-mesenchymal transition generates breast cancer stem cells with a claudin-low phenotype. Cancer Research, 71(13), 4707–4719. https://doi.org/10.1158/0008-5472.CAN-10-4554 Bai, L., Zhang, Z., Zhang, H., Li, X., Yu, Q., Lin, H., & Yang, W. (2008). HIV-1 Tat protein alter the tight junction integrity and function of retinal pigment epithelium: an in vitro study. BMC Infectious Diseases, 8(1), 77. https://doi.org/10.1186/1471-2334-8-77 Balkovetz, D. F. (2006). Claudins at the gate: determinants of renal epithelial tight junction paracellular permeability. American Journal of Physiology-Renal Physiology, 290(3), F572– F579. https://doi.org/10.1152/ajprenal.00135.2005 BARTHOLOMAY, A. F., ROBIN, E. D., VALLEE, R. L., & WACKER, W. E. (1956). Zinc metabolism in hepatic dysfunction. I. Serum zinc concentrations in Laënnec’s cirrhosis and their validation by sequential analysis. The New England Journal of Medicine, 255(9), 403–408. https://doi.org/10.1056/NEJM195608302550901 Bartholow, T. L., Chandran, U. R., Becich, M. J., & Parwani, A. V. (2011). Immunohistochemical profiles of claudin-3 in primary and metastatic prostatic adenocarcinoma. Diagnostic Pathology, 6(1), 12. https://doi.org/10.1186/1746-1596-6-12 Baskić, D., Popović, S., Ristić, P., & Arsenijević, N. N. (2006). Analysis of cycloheximide-induced apoptosis in human leukocytes: Fluorescence microscopy using annexin V/propidium iodide versus acridin orange/ethidium bromide. Cell Biology International, 30(11), 924– 932. https://doi.org/10.1016/j.cellbi.2006.06.016 Baud, L., Perez, J., Friedlander, G., & Ardaillou, R. (1988). Tumor necrosis factor stimulates prostaglandin production and cyclic AMP levels in rat cultured mesangial cells. FEBS Letters, 239(1), 50–54. https://doi.org/10.1016/0014-5793(88)80543-x Baumgartner, H. K., Rudolph, M. C., Ramanathan, P., Burns, V., Webb, P., Bitler, B. G., … Neville, M. C. (2017). Developmental Expression of Claudins in the Mammary Gland. Journal of Mammary Gland Biology and Neoplasia, 22(2), 141–157. https://doi.org/10.1007/s10911- 017-9379-6 Baumholtz, A. I., Simard, A., Nikolopoulou, E., Oosenbrug, M., Collins, M. M., Piontek, A., … Ryan, A. K. (2017). Claudins are essential for cell shape changes and convergent extension movements during neural tube closure. Developmental Biology, 428(1), 25–38. https://doi.org/10.1016/j.ydbio.2017.05.013 Bhat, A. A., Ahmad, R., Uppada, S. B., Singh, A. B., & Dhawan, P. (2016). Claudin-1 promotes TNF-α-induced epithelial-mesenchymal transition and migration in colorectal adenocarcinoma cells. Experimental Cell Research, 349(1), 119–127. https://doi.org/10.1016/j.yexcr.2016.10.005 Bhat, A. A., Uppada, S., Achkar, I. W., Hashem, S., Yadav, S. K., Shanmugakonar, M., … Uddin, S. (2019). Tight Junction Proteins and Signaling Pathways in Cancer and Inflammation: A Functional Crosstalk. Frontiers in Physiology, 9, 1942. https://doi.org/10.3389/fphys.2018.01942 Blake, J. F., Burkard, M., Chan, J., Chen, H., Chou, K. J., Diaz, D., … Schwarz, J. B. (2016). Discovery of (S)-1-(1-(4-Chloro-3-fluorophenyl)-2-hydroxyethyl)-4-(2-((1-methyl-1H- pyrazol-5-yl)amino)pyrimidin-4-yl)pyridin-2(1H)-one (GDC-0994), an Extracellular Signal- Regulated Kinase 1/2 (ERK1/2) Inhibitor in Early Clinical Development. Journal of Medicinal Chemistry, 59(12), 5650–5660. https://doi.org/10.1021/acs.jmedchem.6b00389

Blanchard, A. A., Skliris, G. P., Watson, P. H., Murphy, L. C., Penner, C., Tomes, L., … Myal, Y. (2009). Claudins 1, 3, and 4 protein expression in ER negative breast cancer correlates with markers of the basal phenotype. Virchows Archiv : An International Journal of Pathology, 454(6), 647–656. https://doi.org/10.1007/s00428-009-0770-6 Blaskewicz, C. D., Pudney, J., & Anderson, D. J. (2011). Structure and Function of Intercellular Junctions in Human Cervical and Vaginal Mucosal Epithelia1. Biology of Reproduction, 85(1), 97–104. https://doi.org/10.1095/biolreprod.110.090423 Brenner, D., Blaser, H., & Mak, T. W. (2015). Regulation of tumour necrosis factor signalling: live or let die. Nature Reviews Immunology, 15(6), 362–374. https://doi.org/10.1038/nri3834 Brilli Skvarca, L., Han, H. I., Espiritu, E. B., Missinato, M. A., Rochon, E. R., McDaniels, M. D., … Hukriede, N. A. (2019). Enhancing regeneration after acute kidney injury by promoting cellular dedifferentiation in zebrafish. Disease Models & Mechanisms, 12(4), dmm037390. https://doi.org/10.1242/dmm.037390 Bruewer, M., Luegering, A., Kucharzik, T., Parkos, C. A., Madara, J. L., Hopkins, A. M., & Nusrat, A. (2003). Proinflammatory Cytokines Disrupt Epithelial Barrier Function by Apoptosis- Independent Mechanisms. The Journal of Immunology, 171(11), 6164–6172. https://doi.org/10.4049/jimmunol.171.11.6164 Butt, A. M., Khan, I. B., Hussain, M., Idress, M., Lu, J., & Tong, Y. (2012a). Role of post translational modifications and novel crosstalk between phosphorylation and O-beta- GlcNAc modifications in human claudin-1, -3 and -4. Molecular Biology Reports, 39(2), 1359–1369. https://doi.org/10.1007/s11033-011-0870-7 Butt, A. M., Khan, I. B., Hussain, M., Idress, M., Lu, J., & Tong, Y. (2012b). Role of post translational modifications and novel crosstalk between phosphorylation and O-beta- GlcNAc modifications in human claudin-1, -3 and -4. Molecular Biology Reports, 39(2), 1359–1369. https://doi.org/10.1007/s11033-011-0870-7 Campanale, J. P., Sun, T. Y., & Montell, D. J. (2017). Development and dynamics of cell polarity at a glance. Journal of Cell Science, 130(7), 1201–1207. https://doi.org/10.1242/jcs.188599 Capaldo, C. T., Farkas, A. E., Hilgarth, R. S., Krug, S. M., Wolf, M. F., Benedik, J. K., … Nusrat, A. (2014). Proinflammatory cytokine-induced tight junction remodeling through dynamic self- assembly of claudins. Molecular Biology of the Cell, 25(18), 2710. https://doi.org/10.1091/MBC.E14-02-0773 Carranza, M. L., Rousselot, M., Chibalin, A. V., Bertorello, A. M., Favre, H., & Féraille, E. (1998). Protein kinase A induces recruitment of active Na+, K+-ATPase units to the plasma membrane of rat proximal convoluted tubule cells. Journal of Physiology, 511(1), 235–243. https://doi.org/10.1111/j.1469-7793.1998.235bi.x Challier, J. C., Dubernard, G., Galtier, M., Bintein, T., Vervelle, C., Raison, D., … Uzan, S. (2005). Junctions and adhesion molecules in first trimester and term human placentas. Cellular and Molecular Biology (Noisy-Le-Grand, France), 51 Suppl, OL713-22. Retrieved from http://www.ncbi.nlm.nih.gov/pubmed/16171570 Che, J., Yue, D., Zhang, B., Zhang, H., Huo, Y., Gao, L., … Cao, B. (2018). Claudin-3 Inhibits Lung Squamous Cell Carcinoma Cell Epithelial-mesenchymal Transition and Invasion via Suppression of the Wnt/β-catenin Signaling Pathway. International Journal of Medical Sciences, 15(4), 339–351. https://doi.org/10.7150/ijms.22927 Chen, G., Zhao, X., Tan, Z., Wang, D., Luo, D., Zhang, P., … Li, L. (2018). Investigation of the role

of cullin 4A overexpression in human liver cancer. Molecular Medicine Reports, 18(3), 2531–2540. https://doi.org/10.3892/mmr.2018.9233 Chen, X., Song, P., Fan, P., He, T., Jacobs, D., Levesque, C. L., … Ma, X. (2018). Moderate Dietary Protein Restriction Optimized Gut Microbiota and Mucosal Barrier in Growing Pig Model. Frontiers in Cellular and Infection Microbiology, 8, 246. https://doi.org/10.3389/fcimb.2018.00246 Chow, Y.-W., & Wang, H.-L. (2002). Functional Modulation of P2×2 Receptors by Cyclic AMP- Dependent Protein Kinase. Journal of Neurochemistry, 70(6), 2606–2612. https://doi.org/10.1046/j.1471-4159.1998.70062606.x Claude, P. (1973). FRACTURE FACES OF ZONULAE OCCLUDENTES FROM "TIGHT" AND "LEAKY" EPITHELIA. The Journal of Cell Biology, 58(2), 390–400. https://doi.org/10.1083/jcb.58.2.390 Claverie-Martin, F. (2015). Familial hypomagnesaemia with hypercalciuria and nephrocalcinosis: Clinical and molecular characteristics. Clinical Kidney Journal, Vol. 8, pp. 656–664. https://doi.org/10.1093/ckj/sfv081 Colegio, O. R., Itallie, C. Van, Rahner, C., & Anderson, J. M. (2003). Claudin extracellular domains determine paracellular charge selectivity and resistance but not tight junction fibril architecture. American Journal of Physiology-Cell Physiology, 284(6), C1346–C1354. https://doi.org/10.1152/ajpcell.00547.2002 Coutinho-Camillo, C. M., Lourenço, S. V., da Fonseca, F. P., & Soares, F. A. (2011). Claudin expression is dysregulated in prostate adenocarcinomas but does not correlate with main clinicopathological parameters. Pathology, 43(2), 143–148. https://doi.org/10.1097/PAT.0b013e3283428099 Coyne, C. B., Gambling, T. M., Boucher, R. C., Carson, J. L., & Johnson, L. G. (2003). Role of claudin interactions in airway tight junctional permeability. American Journal of Physiology. Lung Cellular and Molecular Physiology, 285(5), L1166-78. https://doi.org/10.1152/ajplung.00182.2003 Croteau, J. D., Engle, E. L., Queen, S. E., Shirk, E. N., & Zink, M. C. (2017). Marked Enteropathy in an Accelerated Macaque Model of AIDS. The American Journal of Pathology, 187(3), 589– 604. https://doi.org/10.1016/j.ajpath.2016.10.019 Cummins, P. M. (2012). Occludin: One Protein, Many Forms. Molecular and Cellular Biology, 32(2), 242–250. https://doi.org/10.1128/MCB.06029-11 D’Souza, T., Agarwal, R., & Morin, P. J. (2005). Phosphorylation of Claudin-3 at Threonine 192 by cAMP-dependent Protein Kinase Regulates Tight Junction Barrier Function in Ovarian Cancer Cells. Journal of Biological Chemistry, 280(28), 26233–26240. https://doi.org/10.1074/jbc.M502003200 Dahlgren, D., & Lennernäs, H. (2019). Intestinal Permeability and Drug Absorption: Predictive Experimental, Computational and In Vivo Approaches. Pharmaceutics, 11(8), 411. https://doi.org/10.3390/pharmaceutics11080411 Dalton, J. E., Cruickshank, S. M., Egan, C. E., Mears, R., Newton, D. J., Andrew, E. M., … Carding, S. R. (2006). Intraepithelial gammadelta+ lymphocytes maintain the integrity of intestinal epithelial tight junctions in response to infection. Gastroenterology, 131(3), 818–829. https://doi.org/10.1053/j.gastro.2006.06.003 Dan, Q., Shi, Y., Rabani, R., Venugopal, S., Xiao, J., Anwer, S., … Szászi, K. (2019). Claudin-2

suppresses GEF-H1, RHOA, and MRTF thereby impacting proliferation and profibrotic phenotype of tubular cells. Journal of Biological Chemistry, jbc.RA118.006484. https://doi.org/10.1074/jbc.RA118.006484 Danzinger, S., Tan, Y. Y., Rudas, M., Kastner, M.-T., Weingartshofer, S., Muhr, D., & Singer, C. F. (2019). Association of Cytokeratin 5 and Claudin 3 expression with BRCA1 and BRCA2 germline mutations in women with early breast cancer. BMC Cancer, 19(1), 695. https://doi.org/10.1186/s12885-019-5908-6 Daugherty, B. L., Mateescu, M., Patel, A. S., Wade, K., Kimura, S., Gonzales, L. W., … Koval, M. (2004). Developmental regulation of claudin localization by fetal alveolar epithelial cells. American Journal of Physiology. Lung Cellular and Molecular Physiology, 287(6), L1266-73. https://doi.org/10.1152/ajplung.00423.2003 Davies, S. P., Reddy, H., Caivano, M., & Cohen, P. (2000). Specificity and mechanism of action of some commonly used protein kinase inhibitors. Biochemical Journal, 351(1), 95–105. https://doi.org/10.1042/0264-6021:3510095 De Rooij, J., Zwartkruis, F. J. T., Verheijen, M. H. G., Cool, R. H., Nijman, S. M. B., Wittinghofer, A., & Bos, J. L. (1998). Epac is a Rap1 guanine-nucleotide-exchange factor directly activated by cyclic AMP. Nature, 396(6710), 474–477. https://doi.org/10.1038/24884 de Souza, Waldemir F, Fortunato-Miranda, N., Robbs, B. K., de Araujo, W. M., de-Freitas-Junior, J. C., Bastos, L. G., … Morgado-Díaz, J. A. (2013). Claudin-3 overexpression increases the malignant potential of colorectal cancer cells: roles of ERK1/2 and PI3K-Akt as modulators of EGFR signaling. PloS One, 8(9), e74994. https://doi.org/10.1371/journal.pone.0074994 de Souza, Waldemir Fernandes, Barbosa, L. A., Liu, L., de Araujo, W. M., de-Freitas-Junior, J. C. M., Fortunato-Miranda, N., … Morgado-Díaz, J. A. (2014). Ouabain-induced alterations of the apical junctional complex involve α1 and β1 Na,K-ATPase downregulation and ERK1/2 activation independent of caveolae in colorectal cancer cells. The Journal of Membrane Biology, 247(1), 23–33. https://doi.org/10.1007/s00232-013-9607-y Delghandi, M. P., Johannessen, M., & Moens, U. (2005). The cAMP signalling pathway activates CREB through PKA, p38 and MSK1 in NIH 3T3 cells. Cellular Signalling, 17(11), 1343–1351. https://doi.org/10.1016/j.cellsig.2005.02.003 Diomede, F., Zingariello, M., Cavalcanti, M. F. X. B., Merciaro, I., Pizzicannella, J., De Isla, N., … Trubiani, O. (2017). MyD88/ERK/NFkB pathways and pro-inflammatory cytokines release in periodontal ligament stem cells stimulated by Porphyromonas gingivalis. European Journal of Histochemistry : EJH, 61(2), 2791. https://doi.org/10.4081/ejh.2017.2791 Dong, C. X., Zhao, W., Solomon, C., Rowland, K. J., Ackerley, C., Robine, S., … Brubaker, P. L. (2014). The intestinal epithelial insulin-like growth factor-1 receptor links glucagon-like peptide-2 action to gut barrier function. Endocrinology, 155(2), 370–379. https://doi.org/10.1210/en.2013-1871 DONNAHOO, K. K., SHAMES, B. D., HARKEN, A. H., & MELDRUM, D. R. (1999). REVIEW ARTICLE: THE ROLE OF TUMOR NECROSIS FACTOR IN RENAL ISCHEMIA-REPERFUSION INJURY. Journal of Urology, 162(1), 196–203. https://doi.org/10.1097/00005392-199907000-00068 Dörfel, M. J., & Huber, O. (2012). A phosphorylation hotspot within the occludin C-terminal domain. Annals of the New York Academy of Sciences, 1257(1), 38–44. https://doi.org/10.1111/j.1749-6632.2012.06536.x Durán, W. N. (2008). The double-edge sword of TNF-alpha in ischemia-reperfusion injury.

American Journal of Physiology. Heart and Circulatory Physiology, 295(6), H2221-2. https://doi.org/10.1152/ajpheart.01050.2008 Farquhar, M. G. (1963). JUNCTIONAL COMPLEXES IN VARIOUS EPITHELIA. The Journal of Cell Biology, 17(2), 375–412. https://doi.org/10.1083/jcb.17.2.375 Fernández-Blanco, J. A., Estévez, J., Shea-Donohue, T., Martínez, V., & Vergara, P. (2015). Changes in Epithelial Barrier Function in Response to Parasitic Infection: Implications for IBD Pathogenesis. Journal of Crohn’s & Colitis, 9(6), 463–476. https://doi.org/10.1093/ecco-jcc/jjv056 Fiorentino, M., Sapone, A., Senger, S., Camhi, S. S., Kadzielski, S. M., Buie, T. M., … Fasano, A. (2016). Blood-brain barrier and intestinal epithelial barrier alterations in autism spectrum disorders. Molecular Autism, 7(1), 49. https://doi.org/10.1186/s13229-016-0110-z Fleming, T. P., Javed, Q., & Hay, M. (1992). Epithelial differentiation and intercellular junction formation in the mouse early embryo. Development (Cambridge, England). Supplement, 105–112. Retrieved from http://www.ncbi.nlm.nih.gov/pubmed/1299356 Flores-Maldonado, C., Verdejo-Torres, O., Campos-Blázquez, J., Cabrera, A. R., García- Hernández, V., Rincón-Heredia, R., & Contreras, R. G. (2017). Lysosomal Degradation of Junctional Proteins. In Lysosomes - Associated Diseases and Methods to Study Their Function. https://doi.org/10.5772/intechopen.69370 Fromm, M., Piontek, J., Rosenthal, R., Günzel, D., & Krug, S. M. (2017). Tight junctions of the proximal tubule and their channel proteins. Pflugers Archiv : European Journal of Physiology, 469(7–8), 877–887. https://doi.org/10.1007/s00424-017-2001-3 Fujita-Yoshigaki, J. (2011). Analysis of changes in the expression pattern of claudins using salivary acinar cells in primary culture. Methods in Molecular Biology (Clifton, N.J.), 762, 245–258. https://doi.org/10.1007/978-1-61779-185-7_17 Fujita, K., Katahira, J., Horiguchi, Y., Sonoda, N., Furuse, M., & Tsukita, S. (2000). Clostridium perfringens enterotoxin binds to the second extracellular loop of claudin-3, a tight junction integral membrane protein. FEBS Letters, 476(3), 258–261. https://doi.org/10.1016/S0014- 5793(00)01744-0 Furuse, M. (2010a). Molecular basis of the core structure of tight junctions. Cold Spring Harbor Perspectives in Biology, 2(1), a002907. https://doi.org/10.1101/cshperspect.a002907 Furuse, M. (2010b). Molecular basis of the core structure of tight junctions. Cold Spring Harbor Perspectives in Biology, 2(1), a002907. https://doi.org/10.1101/cshperspect.a002907 Furuse, M., Fujita, K., Hiiragi, T., Fujimoto, K., & Tsukita, S. (1998). Claudin-1 and -2: Novel Integral Membrane Proteins Localizing at Tight Junctions with No Sequence Similarity to Occludin. The Journal of Cell Biology, 141(7), 1539–1550. https://doi.org/10.1083/jcb.141.7.1539 Furuse, M., Sasaki, H., & Tsukita, S. (1999). Manner of Interaction of Heterogeneous Claudin Species within and between Tight Junction Strands. The Journal of Cell Biology, 147(4), 891–903. https://doi.org/10.1083/jcb.147.4.891 Gao, Y., Li, S., Wang, J., Luo, C., Zhao, S., & Zheng, N. (2017). Modulation of Intestinal Epithelial Permeability in Differentiated Caco-2 Cells Exposed to Aflatoxin M1 and Ochratoxin A Individually or Collectively. Toxins, 10(1), 13. https://doi.org/10.3390/toxins10010013 Garcia-Hernandez, V., Quiros, M., & Nusrat, A. (2017). Intestinal epithelial claudins: expression and regulation in homeostasis and inflammation. Annals of the New York Academy of

Sciences, 1397(1), 66–79. https://doi.org/10.1111/nyas.13360 Gibson, D. L., Ma, C., Rosenberger, C. M., Bergstrom, K. S. B., Valdez, Y., Huang, J. T., … Vallance, B. A. (2008). Toll-like receptor 2 plays a critical role in maintaining mucosal integrity during Citrobacter rodentium-induced colitis. Cellular Microbiology, 10(2), 388–403. https://doi.org/10.1111/j.1462-5822.2007.01052.x Gogarten, W., Lindeman, K. S., Hirshman, C. A., & Emala, C. W. (2003). Tumor Necrosis Factor Alpha Stimulates Adenylyl Cyclase Activity in Human Myometrial Cells1. Biology of Reproduction, 68(3), 751–757. https://doi.org/10.1095/biolreprod.102.004168 Gonzalez-Mariscal, L., Del CarmenNamorado, M., Martin, D., Sierra, G., & Reyes, J. L. (2006). 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, 21(9), 2391–2398. https://doi.org/10.1093/ndt/gfl255 González-Mariscal, L, Betanzos, A., Nava, P., & Jaramillo, B. E. (2003). Tight junction proteins. Progress in Biophysics and Molecular Biology, 81(1), 1–44. Retrieved from http://www.ncbi.nlm.nih.gov/pubmed/12475568 González-Mariscal, Lorenza, Tapia, R., & Chamorro, D. (2008). Crosstalk of tight junction components with signaling pathways. Biochimica et Biophysica Acta (BBA) - Biomembranes, 1778(3), 729–756. https://doi.org/10.1016/j.bbamem.2007.08.018 Goswami, P., Das, P., Verma, A. K., Prakash, S., Das, T. K., Nag, T. C., … Makharia, G. K. (2014). Are alterations of tight junctions at molecular and ultrastructural level different in duodenal biopsies of patients with celiac disease and Crohn’s disease? Virchows Archiv : An International Journal of Pathology, 465(5), 521–530. https://doi.org/10.1007/s00428- 014-1651-1 Gradauer, K., Iida, M., Watari, A., Kataoka, M., Yamashita, S., Kondoh, M., & Buckley, S. T. (2017). Dodecylmaltoside Modulates Bicellular Tight Junction Contacts To Promote Enhanced Permeability. Molecular Pharmaceutics, 14(12), 4734–4740. https://doi.org/10.1021/acs.molpharmaceut.7b00297 Grell, M., Zimmermann, G., Gottfried, E., Chen, C. M., Grünwald, U., Huang, D. C., … Strasser, A. (1999). Induction of cell death by tumour necrosis factor (TNF) receptor 2,CD40 and CD30: a role for TNF-R1 activation by endogenous membrane-anchored TNF. The EMBO Journal, 18(11), 3034–3043. https://doi.org/10.1093/emboj/18.11.3034 Grosse, B., Degrouard, J., Jaillard, D., & Cassio, D. (2013). Build them up and break them down: Tight junctions of cell lines expressing typical hepatocyte polarity with a varied repertoire of claudins. Tissue Barriers, 1(4), e25210. https://doi.org/10.4161/tisb.25210 Grotegut, S., von Schweinitz, D., Christofori, G., & Lehembre, F. (2006). Hepatocyte growth factor induces cell scattering through MAPK/Egr-1-mediated upregulation of Snail. The EMBO Journal, 25(15), 3534–3545. https://doi.org/10.1038/sj.emboj.7601213 Guillemot, L., Spadaro, D., & Citi, S. (2013). The junctional proteins cingulin and paracingulin modulate the expression of tight junction protein genes through GATA-4. PloS One, 8(2), e55873. https://doi.org/10.1371/journal.pone.0055873 Guntaka, S. R., Samak, G., Seth, A., LaRusso, N. F., & Rao, R. (2011). Epidermal growth factor protects the apical junctional complexes from hydrogen peroxide in bile duct epithelium. Laboratory Investigation; a Journal of Technical Methods and Pathology, 91(9), 1396–1409. https://doi.org/10.1038/labinvest.2011.73

Günzel, D., & Yu, A. S. L. (2013). Claudins and the modulation of tight junction permeability. Physiological Reviews, 93(2), 525–569. https://doi.org/10.1152/physrev.00019.2012 Guo, R.-J., Suh, E. R., & Lynch, J. P. (2004). The role of Cdx proteins in intestinal development and cancer. Cancer Biology & Therapy, 3(7), 593–601. https://doi.org/10.4161/cbt.3.7.913 Gurevich, L. E., Kazantseva, I. A., Sokolova, I. N., Korsakova, N. A., Kuzovleva, E. I., Chistiakova, O. A., … Egorov, V. I. (n.d.). [Solid pseudopapillary tumors of the pancreas: clinical and morphological characteristics, specific features of their immunophenotype, and diagnostic problems]. Arkhiv Patologii, 76(5), 44–54. Retrieved from http://www.ncbi.nlm.nih.gov/pubmed/25543408 Gusti, V., Bennett, K. M., & Lo, D. D. (2014). CD137 signaling enhances tight junction resistance in intestinal epithelial cells. Physiological Reports, 2(8), e12090. https://doi.org/10.14814/phy2.12090 Gustin, J. A., Pincheira, R., Mayo, L. D., Ozes, O. N., Kessler, K. M., Baerwald, M. R., … Donner, D. B. (2004). Tumor necrosis factor activates CRE-binding protein through a p38 MAPK/MSK1 signaling pathway in endothelial cells. American Journal of Physiology-Cell Physiology, 286(3), C547–C555. https://doi.org/10.1152/ajpcell.00332.2002 Guttman, J. A., Li, Y., Wickham, M. E., Deng, W., Vogl, A. W., & Finlay, B. B. (2006). Attaching and effacing pathogen-induced tight junction disruption in vivo. Cellular Microbiology, 8(4), 634–645. https://doi.org/10.1111/j.1462-5822.2005.00656.x Haddad, N., El Andalousi, J., Khairallah, H., Yu, M., Ryan, A. K., & Gupta, I. R. (2011). The tight junction protein claudin-3 shows conserved expression in the nephric duct and ureteric bud and promotes tubulogenesis in vitro. American Journal of Physiology. Renal Physiology, 301(5), F1057-65. https://doi.org/10.1152/ajprenal.00497.2010 Hagen, S. J. (2017). Non-canonical functions of claudin proteins: Beyond the regulation of cell- cell adhesions. Tissue Barriers, 5(2), e1327839. https://doi.org/10.1080/21688370.2017.1327839 Haines, R. J., Beard, R. S., Chen, L., Eitnier, R. A., & Wu, M. H. (2016). Interleukin-1β Mediates β- Catenin-Driven Downregulation of Claudin-3 and Barrier Dysfunction in Caco2 Cells. Digestive Diseases and Sciences, 61(8), 2252–2261. https://doi.org/10.1007/s10620-016- 4145-y Hartsock, A., & Nelson, W. J. (2008, March). Adherens and tight junctions: Structure, function and connections to the actin cytoskeleton. Biochimica et Biophysica Acta - Biomembranes, Vol. 1778, pp. 660–669. https://doi.org/10.1016/j.bbamem.2007.07.012 Hashimoto, K., Oshima, T., Tomita, T., Kim, Y., Matsumoto, T., Joh, T., & Miwa, H. (2008). Oxidative stress induces gastric epithelial permeability through claudin-3. Biochemical and Biophysical Research Communications, 376(1), 154–157. https://doi.org/10.1016/j.bbrc.2008.08.140 Hashizume, A., Ueno, T., Furuse, M., Tsukita, S., Nakanishi, Y., & Hieda, Y. (2004). Expression patterns of claudin family of tight junction membrane proteins in developing mouse submandibular gland. Developmental Dynamics : An Official Publication of the American Association of Anatomists, 231(2), 425–431. https://doi.org/10.1002/dvdy.20142 Hayden, M. S., & Ghosh, S. (2014). Regulation of NF-κB by TNF family cytokines. Seminars in Immunology, 26(3), 253–266. https://doi.org/10.1016/j.smim.2014.05.004 Hei, Y. J., MacDonell, K. L., McNeill, J. H., & Diamond, J. (1991). Lack of correlation between

activation of cyclic AMP-dependent protein kinase and inhibition of contraction of rat vas deferens by cyclic AMP analogs. Molecular Pharmacology, 39(2), 233–238. Retrieved from http://www.ncbi.nlm.nih.gov/pubmed/1847496 Hichino, A., Okamoto, M., Taga, S., Akizuki, R., Endo, S., Matsunaga, T., & Ikari, A. (2017). Down- regulation of Claudin-2 Expression and Proliferation by Epigenetic Inhibitors in Human Lung Adenocarcinoma A549 Cells. The Journal of Biological Chemistry, 292(6), 2411–2421. https://doi.org/10.1074/jbc.M116.762807 Hirano, F., Tanaka, H., Hirano, Y., Hiramoto, M., Handa, H., Makino, I., & Scheidereit, C. (1998). Functional Interference of Sp1 and NF-κB through the Same DNA Binding Site. Molecular and Cellular Biology, 18(3), 1266–1274. https://doi.org/10.1128/MCB.18.3.1266 Hirsch, M., & Noske, W. (1993). The tight junction: Structure and function. Micron, 24(3), 325– 352. https://doi.org/10.1016/0968-4328(93)90057-8 Honda, H., Pazin, M. J., D’Souza, T., Ji, H., & Morin, P. J. (2007). Regulation of the CLDN3 gene in ovarian cancer cells. Cancer Biology & Therapy, 6(11), 1733–1742. https://doi.org/10.4161/cbt.6.11.4832 Hou, J., & Goodenough, D. A. (2010a). Claudin-16 and claudin-19 function in the thick ascending limb. Current Opinion in Nephrology and Hypertension, 19(5), 483–488. https://doi.org/10.1097/MNH.0b013e32833b7125 Hou, J., & Goodenough, D. A. (2010b). Claudin-16 and claudin-19 function in the thick ascending limb. Current Opinion in Nephrology and Hypertension, 19(5), 483. https://doi.org/10.1097/MNH.0B013E32833B7125 Hu, J., Nakano, H., Sakurai, H., & Colburn, N. H. (2004). Insufficient p65 phosphorylation at S536 specifically contributes to the lack of NF- B activation and transformation in resistant JB6 cells. Carcinogenesis, 25(10), 1991–2003. https://doi.org/10.1093/carcin/bgh198 Huang, Y.-H., Bao, Y., Peng, W., Goldberg, M., Love, K., Bumcrot, D. A., … Sawicki, J. A. (2009). Claudin-3 gene silencing with siRNA suppresses ovarian tumor growth and metastasis. Proceedings of the National Academy of Sciences, 106(9), 3426–3430. https://doi.org/10.1073/pnas.0813348106 Hustler, A., Eardley, I., Hinley, J., Pearson, J., Wezel, F., Radvanyi, F., … Southgate, J. (2018). Differential transcription factor expression by human epithelial cells of buccal and urothelial derivation. Experimental Cell Research, 369(2), 284–294. https://doi.org/10.1016/j.yexcr.2018.05.031 Idriss, H. T., & Naismith, J. H. (2000). TNF? and the TNF receptor superfamily: Structure-function relationship(s). Microscopy Research and Technique, 50(3), 184–195. https://doi.org/10.1002/1097-0029(20000801)50:3<184::AID-JEMT2>3.0.CO;2-H Ikari, A., Matsumoto, S., Harada, H., Takagi, K., Hayashi, H., Suzuki, Y., … Miwa, M. (2006). Phosphorylation of paracellin-1 at Ser217 by protein kinase A is essential for localization in tight junctions. Journal of Cell Science, 119(9), 1781–1789. https://doi.org/10.1242/jcs.02901 Ikenouchi, J., Matsuda, M., Furuse, M., & Tsukita, S. (2003). Regulation of tight junctions during the epithelium-mesenchyme transition: direct repression of the gene expression of claudins/occludin by Snail. Journal of Cell Science, 116(10), 1959–1967. https://doi.org/10.1242/jcs.00389 Insel, P. A., & Ostrom, R. S. (2003, June). Forskolin as a tool for examining adenylyl cyclase

expression, regulation, and G protein signaling. Cellular and Molecular Neurobiology, Vol. 23, pp. 305–314. https://doi.org/10.1023/A:1023684503883 Irudayanathan, F. J., Wang, N., Wang, X., & Nangia, S. (2017). Architecture of the paracellular channels formed by claudins of the blood-brain barrier tight junctions. Annals of the New York Academy of Sciences, 1405(1), 131–146. https://doi.org/10.1111/nyas.13378 Itallie, Christina M. Van, Holmes, J., Bridges, A., Gookin, J. L., Coccaro, M. R., Proctor, W., … Anderson, J. M. (2008). The density of small tight junction pores varies among cell types and is increased by expression of claudin-2. Journal of Cell Science, 121(3), 298–305. https://doi.org/10.1242/JCS.021485 Itoh, M., Furuse, M., Morita, K., Kubota, K., Saitou, M., & Tsukita, S. (1999). Direct Binding of Three Tight Junction-Associated Maguks, Zo-1, Zo-2, and Zo-3, with the Cooh Termini of Claudins. The Journal of Cell Biology, 147(6), 1351–1363. https://doi.org/10.1083/jcb.147.6.1351 Jääskeläinen, A., Soini, Y., Jukkola-Vuorinen, A., Auvinen, P., Haapasaari, K.-M., & Karihtala, P. (2018). High-level cytoplasmic claudin 3 expression is an independent predictor of poor survival in triple-negative breast cancer. BMC Cancer, 18(1), 223. https://doi.org/10.1186/s12885-018-4141-z Jung, H., Jun, K. H., Jung, J. H., Chin, H. M., & Park, W. B. (2011). The Expression of Claudin-1, Claudin-2, Claudin-3, and Claudin-4 in Gastric Cancer Tissue. Journal of Surgical Research, 167(2), e185–e191. https://doi.org/10.1016/j.jss.2010.02.010 Kaarteenaho-Wiik, R., & Soini, Y. (2009). Claudin-1, -2, -3, -4, -5, and -7 in usual interstitial pneumonia and sarcoidosis. The Journal of Histochemistry and Cytochemistry : Official Journal of the Histochemistry Society, 57(3), 187–195. https://doi.org/10.1369/jhc.2008.951566 Kaarteenaho, R., Merikallio, H., Lehtonen, S., Harju, T., & Soini, Y. (2010). Divergent expression of claudin -1, -3, -4, -5 and -7 in developing human lung. Respiratory Research, 11(1), 59. https://doi.org/10.1186/1465-9921-11-59 Kakiashvili, E., Dan, Q., Vandermeer, M., Zhang, Y., Waheed, F., Pham, M., & Szászi, K. (2011). The Epidermal Growth Factor Receptor Mediates Tumor Necrosis Factor-α-induced Activation of the ERK/GEF-H1/RhoA Pathway in Tubular Epithelium. Journal of Biological Chemistry, 286(11), 9268–9279. https://doi.org/10.1074/jbc.M110.179903 Kalliolias, G. D., & Ivashkiv, L. B. (2016). TNF biology, pathogenic mechanisms and emerging therapeutic strategies. Nature Reviews Rheumatology, 12(1), 49–62. https://doi.org/10.1038/nrrheum.2015.169 Karasov, W. H. (2017). Integrative physiology of transcellular and paracellular intestinal absorption. The Journal of Experimental Biology, 220(Pt 14), 2495–2501. https://doi.org/10.1242/jeb.144048 Karin, M. (1999, September 24). The beginning of the end: IκB kinase (IKK) and NF-κB activation. Journal of Biological Chemistry, Vol. 274, pp. 27339–27342. https://doi.org/10.1074/jbc.274.39.27339 Kase, H., Iwahashi, K., Nakanishi, S., Matsuda, Y., Yamada, K., Takahashi, M., … Kaneko, M. (1987). K-252 compounds, novel and potent inhibitors of protein kinase C and cyclic nucleotide-dependent protein kinases. Biochemical and Biophysical Research Communications, 142(2), 436–440. https://doi.org/10.1016/0006-291x(87)90293-2

Kempe, S., Kestler, H., Lasar, A., & Wirth, T. (2005). NF- B controls the global pro-inflammatory response in endothelial cells: evidence for the regulation of a pro-atherogenic program. Nucleic Acids Research, 33(16), 5308–5319. https://doi.org/10.1093/nar/gki836 Khan, N., & Asif, A. R. (2015). Transcriptional regulators of claudins in epithelial tight junctions. Mediators of Inflammation, 2015, 219843. https://doi.org/10.1155/2015/219843 Khan, S. B., Cook, H. T., Bhangal, G., Smith, J., Tam, F. W. K., & Pusey, C. D. (2005). Antibody blockade of TNF-α reduces inflammation and scarring in experimental crescentic glomerulonephritis. Kidney International, 67(5), 1812–1820. https://doi.org/10.1111/J.1523-1755.2005.00279.X Kim, B.-G., Lee, P.-H., Lee, S.-H., Park, C.-S., & Jang, A.-S. (2018). Impact of ozone on claudins and tight junctions in the lungs. Environmental Toxicology, 33(7), 798–806. https://doi.org/10.1002/tox.22566 Kim, J. S., Kim, E. J., Lee, S., Tan, X., Liu, X., Park, S., … Ahn, Y.-H. (2019). MiR-34a and miR-34b/c have distinct effects on the suppression of lung adenocarcinomas. Experimental & Molecular Medicine, 51(1), 9. https://doi.org/10.1038/s12276-018-0203-1 Kirk, A., Campbell, S., Bass, P., Mason, J., & Collins, J. (2010). Differential expression of claudin tight junction proteins in the human cortical nephron. Nephrology Dialysis Transplantation, 25(7), 2107–2119. https://doi.org/10.1093/ndt/gfq006 Kiuchi-Saishin, Y., Gotoh, S., Furuse, M., Takasuga, A., Tano, Y., & Tsukita, S. (2002). Differential expression patterns of claudins, tight junction membrane proteins, in mouse nephron segments. Journal of the American Society of Nephrology : JASN, 13(4), 875–886. Retrieved from http://www.ncbi.nlm.nih.gov/pubmed/11912246 Kobayashi, K., Miwa, H., & Yasui, M. (2010). Inflammatory mediators weaken the amniotic membrane barrier through disruption of tight junctions. The Journal of Physiology, 588(Pt 24), 4859–4869. https://doi.org/10.1113/jphysiol.2010.197764 Kobayashi, K., Miwa, H., & Yasui, M. (2011). Progesterone maintains amniotic tight junctions during midpregnancy in mice. Molecular and Cellular Endocrinology, 337(1–2), 36–42. https://doi.org/10.1016/j.mce.2011.01.019 Kobayashi, K., Tsugami, Y., Matsunaga, K., Oyama, S., Kuki, C., & Kumura, H. (2016). Prolactin and glucocorticoid signaling induces lactation-specific tight junctions concurrent with β- casein expression in mammary epithelial cells. Biochimica et Biophysica Acta, 1863(8), 2006–2016. https://doi.org/10.1016/j.bbamcr.2016.04.023 Kooij, G., Kopplin, K., Blasig, R., Stuiver, M., Koning, N., Goverse, G., … de Vries, H. E. (2014a). Disturbed function of the blood-cerebrospinal fluid barrier aggravates neuroinflammation. Acta Neuropathologica, 128(2), 267–277. https://doi.org/10.1007/s00401- 013-1227-1 Kooij, G., Kopplin, K., Blasig, R., Stuiver, M., Koning, N., Goverse, G., … de Vries, H. E. (2014b). Disturbed function of the blood-cerebrospinal fluid barrier aggravates neuroinflammation. Acta Neuropathologica, 128(2), 267–277. https://doi.org/10.1007/s00401- 013-1227-1 Kowalczyk, A. P., & Green, K. J. (2013). Structure, function, and regulation of desmosomes. In Progress in Molecular Biology and Translational Science (Vol. 116, pp. 95–118). https://doi.org/10.1016/B978-0-12-394311-8.00005-4 Krause, G., Protze, J., & Piontek, J. (2015). Assembly and function of claudins: Structure–

function relationships based on homology models and crystal structures. Seminars in Cell & Developmental Biology, 42, 3–12. https://doi.org/10.1016/j.semcdb.2015.04.010 Krause, Gerd, Winkler, L., Mueller, S. L., Haseloff, R. F., Piontek, J., & Blasig, I. E. (2008). Structure and function of claudins. Biochimica et Biophysica Acta (BBA) - Biomembranes, 1778(3), 631–645. https://doi.org/10.1016/J.BBAMEM.2007.10.018 Krug, S. M., Günzel, D., Conrad, M. P., Rosenthal, R., Fromm, A., Amasheh, S., … Fromm, M. (2012). Claudin-17 forms tight junction channels with distinct anion selectivity. Cellular and Molecular Life Sciences, 69(16), 2765–2778. https://doi.org/10.1007/s00018-012-0949-x Kumar, V., Torben, W., Mansfield, J., Alvarez, X., Vande Stouwe, C., Li, J., … Mohan, M. (2019). Cannabinoid Attenuation of Intestinal Inflammation in Chronic SIV-Infected Rhesus Macaques Involves T Cell Modulation and Differential Expression of Micro-RNAs and Pro- inflammatory Genes. Frontiers in Immunology, 10, 914. https://doi.org/10.3389/fimmu.2019.00914 Lappi-Blanco, E., Lehtonen, S. T., Sormunen, R., Merikallio, H. M., Soini, Y., & Kaarteenaho, R. L. (2013). Divergence of tight and adherens junction factors in alveolar epithelium in pulmonary fibrosis. Human Pathology, 44(5), 895–907. https://doi.org/10.1016/j.humpath.2012.08.016 Lech, M., Gröbmayr, R., Ryu, M., Lorenz, G., Hartter, I., Mulay, S. R., … Anders, H.-J. (2014). Macrophage Phenotype Controls Long-Term AKI Outcomes—Kidney Regeneration versus Atrophy. Journal of the American Society of Nephrology, 25(2), 292–304. https://doi.org/10.1681/ASN.2013020152 Lee, N. P. Y., Tong, M. K., Leung, P. P., Chan, V. W., Leung, S., Tam, P.-C., … Luk, J. M. (2006). Kidney claudin-19: Localization in distal tubules and collecting ducts and dysregulation in polycystic renal disease. FEBS Letters, 580(3), 923–931. https://doi.org/10.1016/j.febslet.2006.01.019 Leemhuis, J., Boutillier, S., Schmidt, G., & Meyer, D. K. (2002). The protein kinase A inhibitor H89 acts on cell morphology by inhibiting Rho kinase. Journal of Pharmacology and Experimental Therapeutics, 300(3), 1000–1007. https://doi.org/10.1124/jpet.300.3.1000 Leong, K. G., & Karsan, A. (2000). Signaling pathways mediated by tumor necrosis factor alpha. Histology and Histopathology, 15(4), 1303–1325. https://doi.org/10.14670/HH-15.1303 Li, Jing, Li, Y.-X., Chen, M.-H., Li, J., Du, J., Shen, B., & Xia, X.-M. (2015). Changes in the phosphorylation of claudins during the course of experimental colitis. International Journal of Clinical and Experimental Pathology, 8(10), 12225–12233. Retrieved from http://www.ncbi.nlm.nih.gov/pubmed/26722407 Li, Jixi, Yin, Q., & Wu, H. (2013). Structural basis of signal transduction in the TNF receptor superfamily. Advances in Immunology, 119, 135–153. https://doi.org/10.1016/B978-0-12- 407707-2.00005-9 Li, Juan, Tu, Y., Jiang, L., Xu, H., & Zhang, S. (2012). [Expression and significance of Snail and Claudin-3 in non-small cell lung cancer]. Zhongguo Fei Ai Za Zhi = Chinese Journal of Lung Cancer, 15(10), 583–590. https://doi.org/10.3779/j.issn.1009-3419.2012.10.04 Li, Q., Zhang, Q., Wang, C., Liu, X., Li, N., & Li, J. (2009). Disruption of tight junctions during polymicrobial sepsis in vivo. The Journal of Pathology, 218(2), 210–221. https://doi.org/10.1002/path.2525 Li, Weishan, Li, R., Zhao, S., Jiang, C., Liu, Z., & Tang, X. (2018). Lithium Posttreatment Alleviates

Blood-Brain Barrier Injury After Intracerebral Hemorrhage in Rats. Neuroscience, 383, 129– 137. https://doi.org/10.1016/j.neuroscience.2018.05.001 Li, Wenjing, Xu, C., Wang, K., Ding, Y., & Ding, L. (2019). Non-tight junction-related function of claudin-7 in interacting with integrinβ1 to suppress colorectal cancer cell proliferation and migration. Cancer Management and Research, Volume 11, 1443–1451. https://doi.org/10.2147/CMAR.S188020 Li, X.-Y., Zhang, Y., Wang, X.-X., Jin, C., Wang, Y.-Q., Sun, T.-C., … Liu, Y.-X. (2018). Regulation of blood-testis barrier assembly in vivo by germ cells. FASEB Journal : Official Publication of the Federation of American Societies for Experimental Biology, 32(3), 1653–1664. https://doi.org/10.1096/fj.201700681R Li, Xiao, Wang, Q., Xu, H., Tao, L., Lu, J., Cai, L., & Wang, C. (2014). Somatostatin regulates tight junction proteins expression in colitis mice. International Journal of Clinical and Experimental Pathology, 7(5), 2153–2162. Retrieved from http://www.ncbi.nlm.nih.gov/pubmed/24966923 Li, Xuan, Kimura, H., Hirota, K., Kasuno, K., Torii, K., Okada, T., … Yoshida, H. (2005). Synergistic effect of hypoxia and TNF-α on production of PAI-1 in human proximal renal tubular cells. Kidney International, 68(2), 569–583. https://doi.org/10.1111/j.1523-1755.2005.00435.x Li, Y., Liu, B., Dillon, S. T., Fukudome, E. Y., Kheirbek, T., Sailhamer, E. A., … Alam, H. B. (2010a). Identification of a novel potential biomarker in a model of hemorrhagic shock and valproic acid treatment. The Journal of Surgical Research, 159(1), 474–481. https://doi.org/10.1016/j.jss.2009.04.011 Li, Y., Liu, B., Dillon, S. T., Fukudome, E. Y., Kheirbek, T., Sailhamer, E. A., … Alam, H. B. (2010b). Identification of a novel potential biomarker in a model of hemorrhagic shock and valproic acid treatment. The Journal of Surgical Research, 159(1), 474–481. https://doi.org/10.1016/j.jss.2009.04.011 Liang, X., Zhang, X.-H., Han, B.-C., Lei, W., Qi, Q.-R., Wang, T.-S., … Yang, Z.-M. (2013). Progesterone and heparin-binding epidermal growth factor-like growth factor regulate the expression of tight junction protein Claudin-3 during early pregnancy. Fertility and Sterility, 100(5), 1410–1418. https://doi.org/10.1016/j.fertnstert.2013.07.001 Lim, S., & Kaldis, P. (2013). Cdks, cyclins and CKIs: roles beyond cell cycle regulation. Development, 140(15), 3079–3093. https://doi.org/10.1242/dev.091744 Lim, T. S., Vedula, S. R. K., Hunziker, W., & Lim, C. T. (2008). Kinetics of Adhesion Mediated by Extracellular Loops of Claudin-2 as Revealed by Single-Molecule Force Spectroscopy. Journal of Molecular Biology, 381(3), 681–691. https://doi.org/10.1016/j.jmb.2008.06.009 Lin, X., Shang, X., Manorek, G., & Howell, S. B. (2013a). Regulation of the Epithelial- Mesenchymal Transition by Claudin-3 and Claudin-4. PloS One, 8(6), e67496. https://doi.org/10.1371/journal.pone.0067496 Lin, X., Shang, X., Manorek, G., & Howell, S. B. (2013b). Regulation of the Epithelial- Mesenchymal Transition by Claudin-3 and Claudin-4. PloS One, 8(6), e67496. https://doi.org/10.1371/journal.pone.0067496 Lipschutz, J. H., Li, S., Arisco, A., & Balkovetz, D. F. (2005). Extracellular Signal-regulated Kinases 1/2 Control Claudin-2 Expression in Madin-Darby Canine Kidney Strain I and II Cells. Journal of Biological Chemistry, 280(5), 3780–3788. https://doi.org/10.1074/jbc.M408122200 Mahajan, S. D., Aalinkeel, R., Sykes, D. E., Reynolds, J. L., Bindukumar, B., Adal, A., … Schwartz,

S. A. (2008). Methamphetamine alters blood brain barrier permeability via the modulation of tight junction expression: Implication for HIV-1 neuropathogenesis in the context of drug abuse. Brain Research, 1203, 133–148. https://doi.org/10.1016/j.brainres.2008.01.093 Mandel, I., Paperna, T., Volkowich, A., Merhav, M., Glass-Marmor, L., & Miller, A. (2012). The ubiquitin-proteasome pathway regulates claudin 5 degradation. Journal of Cellular Biochemistry, 113(7), 2415–2423. https://doi.org/10.1002/jcb.24118 Markov, A. G., Falchuk, E. L., Kruglova, N. M., Rybalchenko, O. V, Fromm, M., & Amasheh, S. (2014). Comparative analysis of theophylline and cholera toxin in rat colon reveals an induction of sealing tight junction proteins. Pflugers Archiv : European Journal of Physiology, 466(11), 2059–2065. https://doi.org/10.1007/s00424-014-1460-z Masili-Oku, S. M., Almeida, B. G. L. de, Bacchi, C. E., Filassi, J. R., Baracat, E. C., & Carvalho, F. M. (2017). Lymphocyte-predominant triple-negative breast carcinomas in premenopausal patients: Lower expression of basal immunohistochemical markers. Breast (Edinburgh, Scotland), 31, 34–39. https://doi.org/10.1016/j.breast.2016.10.012 Matsuda, M., Kubo, A., Furuse, M., & Tsukita, S. (2004). A peculiar internalization of claudins, tight junction-specific adhesion molecules, during the intercellular movement of epithelial cells. Journal of Cell Science, 117(7), 1247–1257. https://doi.org/10.1242/jcs.00972 Matter, K., & Balda, M. S. (2003). Signalling to and from tight junctions. Nature Reviews Molecular Cell Biology, 4(3), 225–237. https://doi.org/10.1038/nrm1055 Mattioli, I., Sebald, A., Bucher, C., Charles, R.-P., Nakano, H., Doi, T., … Schmitz, M. L. (2004). Transient and Selective NF-κB p65 Serine 536 Phosphorylation Induced by T Cell Costimulation Is Mediated by IκB Kinase β and Controls the Kinetics of p65 Nuclear Import. The Journal of Immunology, 172(10), 6336–6344. https://doi.org/10.4049/jimmunol.172.10.6336 McCabe, M. J., Allan, C. M., Foo, C. F. H., Nicholls, P. K., McTavish, K. J., & Stanton, P. G. (2012). Androgen initiates Sertoli cell tight junction formation in the hypogonadal (hpg) mouse. Biology of Reproduction, 87(2), 38. https://doi.org/10.1095/biolreprod.111.094318 Mees, S. T., Mennigen, R., Spieker, T., Rijcken, E., Senninger, N., Haier, J., & Bruewer, M. (2009). Expression of tight and adherens junction proteins in ulcerative colitis associated colorectal carcinoma: upregulation of claudin-1, claudin-3, claudin-4, and beta-catenin. International Journal of Colorectal Disease, 24(4), 361–368. https://doi.org/10.1007/s00384-009-0653-y Mei, M., Xiang, R.-L., Cong, X., Zhang, Y., Li, J., Yi, X., … Yu, G.-Y. (2015). Claudin-3 is required for modulation of paracellular permeability by TNF-α through ERK1/2/slug signaling axis in submandibular gland. Cellular Signalling, 27(10), 1915–1927. https://doi.org/10.1016/j.cellsig.2015.07.002 Meng, J., Holdcraft, R. W., Shima, J. E., Griswold, M. D., & Braun, R. E. (2005). Androgens regulate the permeability of the blood-testis barrier. Proceedings of the National Academy of Sciences of the United States of America, 102(46), 16696–16700. https://doi.org/10.1073/pnas.0506084102 Merino-Gracia, J., Costas-Insua, C., Canales, M. Á., & Rodríguez-Crespo, I. (2016). Insights into the C-terminal Peptide Binding Specificity of the PDZ Domain of Neuronal Nitric-oxide Synthase: CHARACTERIZATION OF THE INTERACTION WITH THE TIGHT JUNCTION PROTEIN

CLAUDIN-3. The Journal of Biological Chemistry, 291(22), 11581–11595. https://doi.org/10.1074/jbc.M116.724427 Meyer, T. N., Schwesinger, C., Bush, K. T., Stuart, R. O., Rose, D. W., Shah, M. M., … Nigam, S. K. (2004). Spatiotemporal regulation of morphogenetic molecules during in vitro branching of the isolated ureteric bud: toward a model of branching through budding in the developing kidney. Developmental Biology, 275(1), 44–67. https://doi.org/10.1016/j.ydbio.2004.07.022 Michikawa, H., Fujita-Yoshigaki, J., & Sugiya, H. (2008). Enhancement of barrier function by overexpression of claudin-4 in tight junctions of submandibular gland cells. Cell and Tissue Research, 334(2), 255–264. https://doi.org/10.1007/s00441-008-0689-2 Milatz, S., Krug, S. M., Rosenthal, R., Günzel, D., Müller, D., Schulzke, J.-D., … Fromm, M. (2010). Claudin-3 acts as a sealing component of the tight junction for ions of either charge and uncharged solutes. Biochimica et Biophysica Acta, 1798(11), 2048–2057. https://doi.org/10.1016/j.bbamem.2010.07.014 Milatz, S., Piontek, J., Schulzke, J.-D., Blasig, I. E., Fromm, M., & Günzel, D. (2015). Probing the cis-arrangement of prototype tight junction proteins claudin-1 and claudin-3. The Biochemical Journal, 468(3), 449–458. https://doi.org/10.1042/BJ20150148 MISSERI, R., MELDRUM, D. R., DAGHER, P., HILE, K., RINK, R. C., & MELDRUM, K. K. (2004). UNILATERAL URETERAL OBSTRUCTION INDUCES RENAL TUBULAR CELL PRODUCTION OF TUMOR NECROSIS FACTOR-α INDEPENDENT OF INFLAMMATORY CELL INFILTRATION. Journal of Urology, 172(4 Part 2), 1595–1599. https://doi.org/10.1097/01.ju.0000138902.57626.70 Mitchell, L. A., Overgaard, C. E., Ward, C., Margulies, S. S., & Koval, M. (2011). Differential effects of claudin-3 and claudin-4 on alveolar epithelial barrier function. American Journal of Physiology-Lung Cellular and Molecular Physiology, 301(1), L40–L49. https://doi.org/10.1152/ajplung.00299.2010 Miyoshi, Y., Tanabe, S., & Suzuki, T. (2016). Cellular zinc is required for intestinal epithelial barrier maintenance via the regulation of claudin-3 and occludin expression. American Journal of Physiology. Gastrointestinal and Liver Physiology, 311(1), G105-16. https://doi.org/10.1152/ajpgi.00405.2015 Mohammadi, M. T., & Dehghani, G. A. (2014). Acute hypertension induces brain injury and blood-brain barrier disruption through reduction of claudins mRNA expression in rat. Pathology, Research and Practice, 210(12), 985–990. https://doi.org/10.1016/j.prp.2014.05.007 Moldvay, J., Jäckel, M., Páska, C., Soltész, I., Schaff, Z., & Kiss, A. (2007). Distinct claudin expression profile in histologic subtypes of lung cancer. Lung Cancer (Amsterdam, Netherlands), 57(2), 159–167. https://doi.org/10.1016/j.lungcan.2007.02.018 Molina-Jijón, E., Rodríguez-Muñoz, R., Namorado, M. del C., Pedraza-Chaverri, J., & Reyes, J. L. (2014). Oxidative stress induces claudin-2 nitration in experimental type 1 diabetic nephropathy. Free Radical Biology and Medicine, 72, 162–175. https://doi.org/10.1016/j.freeradbiomed.2014.03.040 Morita, K., Furuse, M., Fujimoto, K., & Tsukita, S. (1999). Claudin multigene family encoding four-transmembrane domain protein components of tight junction strands. Proceedings of the National Academy of Sciences of the United States of America, 96(2), 511–516.

https://doi.org/10.1073/pnas.96.2.511 Murakami, T., Felinski, E. A., & Antonetti, D. A. (2009). Occludin Phosphorylation and Ubiquitination Regulate Tight Junction Trafficking and Vascular Endothelial Growth Factor- induced Permeability. Journal of Biological Chemistry, 284(31), 21036–21046. https://doi.org/10.1074/jbc.M109.016766 Murray, A. J. (2008, June 3). Pharmacological PKA inhibition: All may not be what it seems. Science Signaling, Vol. 1. https://doi.org/10.1126/scisignal.122re4 Nagaoka, K., Zhang, H., Watanabe, G., & Taya, K. (2013). Epithelial cell differentiation regulated by MicroRNA-200a in mammary glands. PloS One, 8(6), e65127. https://doi.org/10.1371/journal.pone.0065127 Naqvi, S., Martin, K. J., & Arthur, J. S. C. (2014). CREB phosphorylation at Ser 133 regulates transcription via distinct mechanisms downstream of cAMP and MAPK signalling. Biochemical Journal, 458(3), 469–479. https://doi.org/10.1042/BJ20131115 NAVARRO, J., & MORAFERNANDEZ, C. (2006). The role of TNF-α in diabetic nephropathy: Pathogenic and therapeutic implications. Cytokine & Growth Factor Reviews, 17(6), 441– 450. https://doi.org/10.1016/j.cytogfr.2006.09.011 Nicholson, M. D. O., Lindsay, L. A., & Murphy, C. R. (2010). Ovarian hormones control the changing expression of claudins and occludin in rat uterine epithelial cells during early pregnancy. Acta Histochemica, 112(1), 42–52. https://doi.org/10.1016/j.acthis.2008.07.003 Nomme, J., Antanasijevic, A., Caffrey, M., Van Itallie, C. M., Anderson, J. M., Fanning, A. S., & Lavie, A. (2015). Structural Basis of a Key Factor Regulating the Affinity between the Zonula Occludens First PDZ Domain and Claudins. The Journal of Biological Chemistry, 290(27), 16595–16606. https://doi.org/10.1074/jbc.M115.646695 Nozaki, Y., Hino, S., Ri, J., Sakai, K., Nagare, Y., Kawanishi, M., … Matsumura, I. (2017). Lipopolysaccharide-Induced Acute Kidney Injury Is Dependent on an IL-18 Receptor Signaling Pathway. International Journal of Molecular Sciences, 18(12). https://doi.org/10.3390/ijms18122777 Obrig, T. G., Culp, W. J., McKeehan, W. L., & Hardesty, B. (1971). The mechanism by which cycloheximide and related glutarimide antibiotics inhibit peptide synthesis on reticulocyte ribosomes. Journal of Biological Chemistry, 246(1), 174–181. Okazaki, T., Sakon, S., Sasazuki, T., Sakurai, H., Doi, T., Yagita, H., … Nakano, H. (2003). Phosphorylation of serine 276 is essential for p65 NF-kappaB subunit-dependent cellular responses. Biochemical and Biophysical Research Communications, 300(4), 807–812. https://doi.org/10.1016/s0006-291x(02)02932-7 Okugawa, T., Oshima, T., Chen, X., Hori, K., Tomita, T., Fukui, H., … Miwa, H. (2012). Down- regulation of claudin-3 is associated with proliferative potential in early gastric cancers. Digestive Diseases and Sciences, 57(6), 1562–1567. https://doi.org/10.1007/s10620-012- 2043-5 Oleinikova, N. A., Kharlova, O. A., Mal’kov, P. G., & Danilova, N. V. (2017). [The expression of claudins in colonic neoplasms]. Arkhiv Patologii, 79(1), 28–35. https://doi.org/10.17116/patol201779128-35 Pahl, H. L. (1999, November 22). Activators and target genes of Rel/NF-κB transcription factors. Oncogene, Vol. 18, pp. 6853–6866. https://doi.org/10.1038/sj.onc.1203239

Pahle, J., Menzel, L., Niesler, N., Kobelt, D., Aumann, J., Rivera, M., & Walther, W. (2017). Rapid eradication of colon carcinoma by Clostridium perfringens Enterotoxin suicidal gene therapy. BMC Cancer, 17(1), 129. https://doi.org/10.1186/s12885-017-3123-x Papadopoulos, D., Shihan, M., & Scheiner-Bobis, G. (2018). Physiological implications of DHEAS- induced non-classical steroid hormone signaling. The Journal of Steroid Biochemistry and Molecular Biology, 179, 73–78. https://doi.org/10.1016/j.jsbmb.2017.10.002 Parikh, N. U., Aalinkeel, R., Reynolds, J. L., Nair, B. B., Sykes, D. E., Mammen, M. J., … Mahajan, S. D. (2015). Galectin-1 suppresses methamphetamine induced neuroinflammation in human brain microvascular endothelial cells: Neuroprotective role in maintaining blood brain barrier integrity. Brain Research, 1624, 175–187. https://doi.org/10.1016/j.brainres.2015.07.033 Pearce, S. C., Mani, V., Boddicker, R. L., Johnson, J. S., Weber, T. E., Ross, J. W., … Gabler, N. K. (2013). Heat Stress Reduces Intestinal Barrier Integrity and Favors Intestinal Glucose Transport in Growing Pigs. PLoS ONE, 8(8), e70215. https://doi.org/10.1371/journal.pone.0070215 Peng, S., Wang, S.-B., Singh, D., Zhao, P. Y. C., Davis, K., Chen, B., … Rizzolo, L. J. (2016). Claudin- 3 and claudin-19 partially restore native phenotype to ARPE-19 cells via effects on tight junctions and gene expression. Experimental Eye Research, 151, 179–189. https://doi.org/10.1016/j.exer.2016.08.021 Peppi, M., & Ghabriel, M. N. (2004). Tissue-specific expression of the tight junction proteins claudins and occludin in the rat salivary glands. Journal of Anatomy, 205(4), 257–266. https://doi.org/10.1111/j.0021-8782.2004.00332.x Pierce, J. W., Schoenleber, R., Jesmok, G., Best, J., Moore, S. A., Collins, T., & Gerritsen, M. E. (1997). Novel inhibitors of cytokine-induced IκBα phosphorylation and endothelial cell adhesion molecule expression show anti-inflammatory effects in vivo. Journal of Biological Chemistry, 272(34), 21096–21103. https://doi.org/10.1074/jbc.272.34.21096 Piontek, A., Rossa, J., Protze, J., Wolburg, H., Hempel, C., Günzel, D., … Piontek, J. (2017). Polar and charged extracellular residues conserved among barrier-forming claudins contribute to tight junction strand formation. Annals of the New York Academy of Sciences, 1397(1), 143–156. https://doi.org/10.1111/nyas.13341 Plain, A., & Alexander, R. T. (2018). Claudins and nephrolithiasis. Current Opinion in Nephrology and Hypertension, 27(4), 268–276. https://doi.org/10.1097/MNH.0000000000000426 Pradhan-Sundd, T., Zhou, L., Vats, R., Jiang, A., Molina, L., Singh, S., … Monga, S. P. (2018). Dual catenin loss in murine liver causes tight junctional deregulation and progressive intrahepatic cholestasis. Hepatology (Baltimore, Md.), 67(6), 2320–2337. https://doi.org/10.1002/hep.29585 Prasad, S., Mingrino, R., Kaukinen, K., Hayes, K. L., Powell, R. M., MacDonald, T. T., & Collins, J. E. (2005). Inflammatory processes have differential effects on claudins 2, 3 and 4 in colonic epithelial cells. Laboratory Investigation; a Journal of Technical Methods and Pathology, 85(9), 1139–1162. https://doi.org/10.1038/labinvest.3700316 Raviola, G., & Raviola, E. (1978). Intercellular junctions in the ciliary epithelium. Investigative Ophthalmology & Visual Science, 17(10), 958–981. Retrieved from http://www.ncbi.nlm.nih.gov/pubmed/100466 Reiling, J., Bridle, K. R., Gijbels, M., Schaap, F. G., Jaskowski, L., Santrampurwala, N., … Fawcett,

J. (2017). Low-Dose Lipopolysaccharide Causes Biliary Injury by Blood Biliary Barrier Impairment in a Rat Hepatic Ischemia/Reperfusion Model. Liver Transplantation : Official Publication of the American Association for the Study of Liver Diseases and the International Liver Transplantation Society, 23(2), 194–206. https://doi.org/10.1002/lt.24681 Reyes, J. L., Lamas, M., Martin, D., Del Carmen Namorado, M., Islas, S., Luna, J., … González- Mariscal, L. (2002). The renal segmental distribution of claudins changes with development. Kidney International, 62(2), 476–487. https://doi.org/10.1046/j.1523- 1755.2002.00479.x Rimon, E., Sasson, R., Dantes, A., Land-Bracha, A., & Amsterdam, A. (2004). Gonadotropin- induced gene regulation in human granulosa cells obtained from IVF patients: modulation of genes coding for growth factors and their receptors and genes involved in cancer and other diseases. International Journal of Oncology, 24(5), 1325–1338. https://doi.org/10.3892/ijo.24.5.1325 Robbins, J. D., Boring, D. L., Tang, W. J., Shank, R., & Seamon, K. B. (1996). Forskolin carbamates: binding and activation studies with type I adenylyl cyclase. Journal of Medicinal Chemistry, 39(14), 2745–2752. https://doi.org/10.1021/jm960191+ Rodenburg, R. N. P., Snijder, J., van de Waterbeemd, M., Schouten, A., Granneman, J., Heck, A. J. R., & Gros, P. (2017). Stochastic palmitoylation of accessible cysteines in membrane proteins revealed by native mass spectrometry. Nature Communications, 8(1), 1280. https://doi.org/10.1038/s41467-017-01461-z Rodgers, L. S., & Fanning, A. S. (2011). Regulation of epithelial permeability by the actin cytoskeleton. Cytoskeleton, 68(12), 653–660. https://doi.org/10.1002/cm.20547 Romero, A., Ares, I., Ramos, E., Castellano, V., Martínez, M., Martínez-Larrañaga, M.-R., … Martínez, M.-A. (2016). Mycotoxins modify the barrier function of Caco-2 cells through differential gene expression of specific claudin isoforms: Protective effect of illite mineral clay. Toxicology, 353–354, 21–33. https://doi.org/10.1016/j.tox.2016.05.003 Rossa, J., Lorenz, D., Ringling, M., Veshnyakova, A., & Piontek, J. (2012). Overexpression of claudin-5 but not claudin-3 induces formation of trans-interaction-dependent multilamellar bodies. Annals of the New York Academy of Sciences, 1257(1), 59–66. https://doi.org/10.1111/j.1749-6632.2012.06546.x Roy, P., Mukherjee, T., Chatterjee, B., Vijayaragavan, B., Banoth, B., & Basak, S. (2017). Non- canonical NFκB mutations reinforce pro-survival TNF response in multiple myeloma through an autoregulatory RelB:p50 NFκB pathway. Oncogene, 36(10), 1417–1429. https://doi.org/10.1038/onc.2016.309 Sabio, G., & Davis, R. J. (2014). TNF and MAP kinase signalling pathways. Seminars in Immunology, 26(3), 237–245. https://doi.org/10.1016/j.smim.2014.02.009 Sakaguchi, T., Gu, X., Golden, H. M., Suh, E., Rhoads, D. B., & Reinecker, H.-C. (2002). Cloning of the Human Claudin-2 5′-Flanking Region Revealed a TATA-less Promoter with Conserved Binding Sites in Mouse and Human for Caudal-related Homeodomain Proteins and Hepatocyte Nuclear Factor-1α. Journal of Biological Chemistry, 277(24), 21361–21370. https://doi.org/10.1074/jbc.M110261200 Sakai, N., Chiba, H., Fujita, H., Akashi, Y., Osanai, M., Kojima, T., & Sawada, N. (2007). Expression patterns of claudin family of tight-junction proteins in the mouse prostate. Histochemistry

and Cell Biology, 127(4), 457–462. https://doi.org/10.1007/s00418-007-0269-7 Santos, F. C. A., Falleiros-Júnior, L. R., Corradi, L. S., Vilamaior, P. S. L., & Taboga, S. R. (2007). Experimental endocrine therapies promote epithelial cytodifferentiation and ciliogenesis in the gerbil female prostate. Cell and Tissue Research, 328(3), 617–624. https://doi.org/10.1007/s00441-007-0381-y Satake, S., Semba, S., Matsuda, Y., Usami, Y., Chiba, H., Sawada, N., … Yokozaki, H. (2008). Cdx2 transcription factor regulates claudin-3 and claudin-4 expression during intestinal differentiation of gastric carcinoma. Pathology International, 58(3), 156–163. https://doi.org/10.1111/j.1440-1827.2007.02204.x Savickiene, J., Treigyte, G., Pivoriunas, A., Navakauskiene, R., & Magnusson, K.-E. (2004). Sp1 and NF-kappaB transcription factor activity in the regulation of the p21 and FasL promoters during promyelocytic leukemia cell monocytic differentiation and its associated apoptosis. Annals of the New York Academy of Sciences, 1030(1), 569–577. https://doi.org/10.1196/annals.1329.066 Scheurer, S. B., Rybak, J. N., Rösli, C., Neri, D., & Elia, G. (2004). Modulation of gene expression by hypoxia in human umbilical cord vein endothelial cells: A transcriptomic and proteomic study. Proteomics, 4(6), 1737–1760. https://doi.org/10.1002/pmic.200300689 Schlüter, M. A., & Margolis, B. (2012). Apicobasal polarity in the kidney. Experimental Cell Research, 318(9), 1033–1039. https://doi.org/10.1016/J.YEXCR.2012.02.028 Schmelzer, E., Wauthier, E., & Reid, L. M. (2006). The phenotypes of pluripotent human hepatic progenitors. Stem Cells (Dayton, Ohio), 24(8), 1852–1858. https://doi.org/10.1634/stemcells.2006-0036 Schmidt, H., Braubach, P., Schilpp, C., Lochbaum, R., Neuland, K., Thompson, K., … Wittekindt, O. H. (2019). IL-13 Impairs Tight Junctions in Airway Epithelia. International Journal of Molecular Sciences, 20(13), 3222. https://doi.org/10.3390/ijms20133222 Schulzke, J.-D., Günzel, D., John, L. J., & Fromm, M. (2012). Perspectives on tight junction research. Annals of the New York Academy of Sciences, 1257(1), 1–19. https://doi.org/10.1111/j.1749-6632.2012.06485.x Schumann, M., Günzel, D., Buergel, N., Richter, J. F., Troeger, H., May, C., … Schulzke, J.-D. (2012). Cell polarity-determining proteins Par-3 and PP-1 are involved in epithelial tight junction defects in coeliac disease. Gut, 61(2), 220–228. https://doi.org/10.1136/gutjnl- 2011-300123 Schütze, S., Wiegmann, K., Machleidt, T., & Krönke, M. (1995). TNF-induced activation of NF- kappa B. Immunobiology, 193(2–4), 193–203. Retrieved from http://www.ncbi.nlm.nih.gov/pubmed/8530143 Shang, X., Lin, X., Alvarez, E., Manorek, G., & Howell, S. B. (2012). Tight junction proteins claudin-3 and claudin-4 control tumor growth and metastases. Neoplasia (New York, N.Y.), 14(10), 974–985. https://doi.org/10.1593/neo.12942 Shen, L., Weber, C. R., Raleigh, D. R., Yu, D., & Turner, J. R. (2011). Tight Junction Pore and Leak Pathways: A Dynamic Duo. Annual Review of Physiology, 73(1), 283–309. https://doi.org/10.1146/annurev-physiol-012110-142150 Shi, J., Barakat, M., Chen, D., & Chen, L. (2018). Bicellular Tight Junctions and Wound Healing. International Journal of Molecular Sciences, 19(12), 3862. https://doi.org/10.3390/ijms19123862

Shiozaki, A., Bai, X., Shen-Tu, G., Moodley, S., Takeshita, H., Fung, S.-Y., … Liu, M. (2012). Claudin 1 Mediates TNFα-Induced Gene Expression and Cell Migration in Human Lung Carcinoma Cells. PLoS ONE, 7(5), e38049. https://doi.org/10.1371/journal.pone.0038049 Shiozaki, A., Shimizu, H., Ichikawa, D., Konishi, H., Komatsu, S., Kubota, T., … Otsuji, E. (2014). Claudin 1 mediates tumor necrosis factor alpha-induced cell migration in human gastric cancer cells. World Journal of Gastroenterology, 20(47), 17863–17876. https://doi.org/10.3748/wjg.v20.i47.17863 Sikora, M., Chrabąszcz, M., Waśkiel-Burnat, A., Rakowska, A., Olszewska, M., & Rudnicka, L. (2019). Claudin-3 - a new intestinal integrity marker in patients with psoriasis: association with disease severity. Journal of the European Academy of Dermatology and Venereology : JEADV, jdv.15700. https://doi.org/10.1111/jdv.15700 Singh, P., Bahrami, L., Castillo, A., & Majid, D. S. A. (2013). TNF-α type 2 receptor mediates renal inflammatory response to chronic angiotensin II administration with high salt intake in mice. American Journal of Physiology. Renal Physiology, 304(7), F991-9. https://doi.org/10.1152/ajprenal.00525.2012 Søfteland, J. M., Casselbrant, A., Biglarnia, A.-R., Linders, J., Hellström, M., Pesce, A., … Oltean, M. (2019). Intestinal Preservation Injury: A Comparison Between Rat, Porcine and Human Intestines. International Journal of Molecular Sciences, 20(13), 3135. https://doi.org/10.3390/ijms20133135 Someya, M., Kojima, T., Ogawa, M., Ninomiya, T., Nomura, K., Takasawa, A., … Sawada, N. (2013). Regulation of tight junctions by sex hormones in normal human endometrial epithelial cells and uterus cancer cell line Sawano. Cell and Tissue Research, 354(2), 481– 494. https://doi.org/10.1007/s00441-013-1676-9 Sonoda, N., Furuse, M., Sasaki, H., Yonemura, S., Katahira, J., Horiguchi, Y., & Tsukita, S. (1999). Clostridium perfringens enterotoxin fragment removes specific claudins from tight junction strands: Evidence for direct involvement of claudins in tight junction barrier. The Journal of Cell Biology, 147(1), 195–204. https://doi.org/10.1083/jcb.147.1.195 Stammler, A., Lüftner, B. U., Kliesch, S., Weidner, W., Bergmann, M., Middendorff, R., & Konrad, L. (2016). Highly Conserved Testicular Localization of Claudin-11 in Normal and Impaired Spermatogenesis. PloS One, 11(8), e0160349. https://doi.org/10.1371/journal.pone.0160349 Stokes, M. B., Foster, K., Markowitz, G. S., Ebrahimi, F., Hines, W., Kaufman, D., … D’Agati, V. D. (2005). Development of glomerulonephritis during anti-TNF-α therapy for rheumatoid arthritis. Nephrology Dialysis Transplantation, 20(7), 1400–1406. https://doi.org/10.1093/ndt/gfh832 Suominen, J., Wang, Y., Kaipia, A., & Toppari, J. (2004). Tumor necrosis factor-alpha (TNF-alpha) promotes cell survival during spermatogenesis, and this effect can be blocked by infliximab, a TNF-alpha antagonist. European Journal of Endocrinology, 151(5), 629–640. https://doi.org/10.1530/eje.0.1510629 Suzuki, T., Elias, B. C., Seth, A., Shen, L., Turner, J. R., Giorgianni, F., … Rao, R. (2009). PKC regulates occludin phosphorylation and epithelial tight junction integrity. Proceedings of the National Academy of Sciences, 106(1), 61–66. https://doi.org/10.1073/pnas.0802741106 Suzuki, Takuya, Yoshinaga, N., & Tanabe, S. (2011). Interleukin-6 (IL-6) Regulates Claudin-2

Expression and Tight Junction Permeability in Intestinal Epithelium. Journal of Biological Chemistry, 286(36), 31263–31271. https://doi.org/10.1074/jbc.M111.238147 Szakál, D. N., Gyorffy, H., Arató, A., Cseh, A., Molnár, K., Papp, M., … Veres, G. (2010). Mucosal expression of claudins 2, 3 and 4 in proximal and distal part of duodenum in children with coeliac disease. Virchows Archiv : An International Journal of Pathology, 456(3), 245–250. https://doi.org/10.1007/s00428-009-0879-7 Szaszi, K., & Amoozadeh, Y. (2014). New insights into functions, regulation, and pathological roles of tight junctions in kidney tubular epithelium. International Review of Cell and Molecular Biology, 308, 205–271. https://doi.org/10.1016/B978-0-12-800097-7.00006-3 Taddei, A., Giampietro, C., Conti, A., Orsenigo, F., Breviario, F., Pirazzoli, V., … Dejana, E. (2008). Endothelial adherens junctions control tight junctions by VE-cadherin-mediated upregulation of claudin-5. Nature Cell Biology, 10(8), 923–934. https://doi.org/10.1038/ncb1752 Takala, H., Saarnio, J., Wiik, H., & Soini, Y. (2007). Claudins 1, 3, 4, 5 and 7 in esophageal cancer: loss of claudin 3 and 4 expression is associated with metastatic behavior. APMIS : Acta Pathologica, Microbiologica, et Immunologica Scandinavica, 115(7), 838–847. https://doi.org/10.1111/j.1600-0463.2007.apm_656.x Takano, K., Kojima, T., Sawada, N., & Himi, T. (2014). Role of tight junctions in signal transduction: an update. EXCLI Journal, 13, 1145–1162. Retrieved from http://www.ncbi.nlm.nih.gov/pubmed/26417329 Takehara, M., Nishimura, T., Mima, S., Hoshino, T., & Mizushima, T. (2009). Effect of Claudin Expression on Paracellular Permeability, Migration and Invasion of Colonic Cancer Cells. Biological & Pharmaceutical Bulletin, 32(5), 825–831. https://doi.org/10.1248/bpb.32.825 Tamagawa, H., Takahashi, I., Furuse, M., Yoshitake-Kitano, Y., Tsukita, S., Ito, T., … Kiyono, H. (2003). Characteristics of claudin expression in follicle-associated epithelium of Peyer’s patches: preferential localization of claudin-4 at the apex of the dome region. Laboratory Investigation; a Journal of Technical Methods and Pathology, 83(7), 1045–1053. Retrieved from http://www.ncbi.nlm.nih.gov/pubmed/12861044 Tan, S., Duan, H., Xun, T., Ci, W., Qiu, J., Yu, F., … Liu, S. (2014). HIV-1 impairs human retinal pigment epithelial barrier function: possible association with the pathogenesis of HIV- associated retinopathy. Laboratory Investigation; a Journal of Technical Methods and Pathology, 94(7), 777–787. https://doi.org/10.1038/labinvest.2014.72 Tanaka, H., Imasato, M., Yamazaki, Y., Matsumoto, K., Kunimoto, K., Delpierre, J., … Tsukita, S. (2018). Claudin-3 regulates bile canalicular paracellular barrier and cholesterol gallstone core formation in mice. Journal of Hepatology, 69(6), 1308–1316. https://doi.org/10.1016/j.jhep.2018.08.025 Tarulli, G. A., Meachem, S. J., Schlatt, S., & Stanton, P. G. (2008). Regulation of testicular tight junctions by gonadotrophins in the adult Djungarian hamster in vivo. Reproduction (Cambridge, England), 135(6), 867–877. https://doi.org/10.1530/REP-07-0572 Tatum, R., Zhang, Y., Lu, Q., Kim, K., Jeansonne, B. G., & Chen, Y.-H. (2007). WNK4 phosphorylates ser 206 of claudin-7 and promotes paracellular Cl − permeability. FEBS Letters, 581(20), 3887–3891. https://doi.org/10.1016/j.febslet.2007.07.014 Tavakoli, S., & Xiao, L. (2019). Depletion of Intestinal Microbiome Partially Rescues Bone Loss in Sickle Cell Disease Male Mice. Scientific Reports, 9(1), 8659.

https://doi.org/10.1038/s41598-019-45270-4 Tebbe, B., Mankertz, J., Schwarz, C., Amasheh, S., Fromm, M., Assaf, C., … Orfanos, C. E. (2002). Tight junction proteins: a novel class of integral membrane proteins. Expression in human epidermis and in HaCaT keratinocytes. Archives of Dermatological Research, 294(1–2), 14– 18. https://doi.org/10.1007/s00403-001-0290-y Tian, L., Li, Y., Zhang, J., Chang, R., Li, J., & Huo, L. (2018). IL-9 promotes the pathogenesis of ulcerative colitis through STAT3/SOCS3 signaling. Bioscience Reports, 38(6), BSR20181521. https://doi.org/10.1042/BSR20181521 Tikiyani, V., & Babu, K. (2019). Claudins in the brain: Unconventional functions in neurons. Traffic, tra.12685. https://doi.org/10.1111/tra.12685 Tipsmark, C. K., Kiilerich, P., Nilsen, T. O., Ebbesson, L. O. E., Stefansson, S. O., & Madsen, S. S. (2008). Branchial expression patterns of claudin isoforms in Atlantic salmon during seawater acclimation and smoltification. American Journal of Physiology. Regulatory, Integrative and Comparative Physiology, 294(5), R1563-74. https://doi.org/10.1152/ajpregu.00915.2007 Tipsmark, C. K., & Madsen, S. S. (2012). Tricellulin, occludin and claudin-3 expression in salmon intestine and kidney during salinity adaptation. Comparative Biochemistry and Physiology. Part A, Molecular & Integrative Physiology, 162(4), 378–385. https://doi.org/10.1016/j.cbpa.2012.04.020 Tiwari, A., Loughner, C. L., Swamynathan, S., & Swamynathan, S. K. (2017). KLF4 Plays an Essential Role in Corneal Epithelial Homeostasis by Promoting Epithelial Cell Fate and Suppressing Epithelial-Mesenchymal Transition. Investigative Ophthalmology & Visual Science, 58(5), 2785–2795. https://doi.org/10.1167/iovs.17-21826 Tokuhara, Y., Morinishi, T., Matsunaga, T., Sakai, M., Sakai, T., Ohsaki, H., … Hirakawa, E. (2018). Nuclear expression of claudin-3 in human colorectal adenocarcinoma cell lines and tissues. Oncology Letters, 15(1), 99–108. https://doi.org/10.3892/ol.2017.7281 Tokumasu, R., Tamura, A., & Tsukita, S. (2017). Time- and dose-dependent claudin contribution to biological functions: Lessons from claudin-1 in skin. Tissue Barriers, 5(3), e1336194. https://doi.org/10.1080/21688370.2017.1336194 Truong, L. D., Gaber, L., & Eknoyan, G. (2011). Obstructive Uropathy. In Experimental Models for Renal Diseases (pp. 311–326). https://doi.org/10.1159/000314578 TSUKITA, S., & FURUSE, M. (2006). The Structure and Function of Claudins, Cell Adhesion Molecules at Tight Junctions. Annals of the New York Academy of Sciences, 915(1), 129– 135. https://doi.org/10.1111/j.1749-6632.2000.tb05235.x Twiss, F., Oldenkamp, M., Hiemstra, A., Zhou, H., Matheron, L., Mohammed, S., & de Rooij, J. (2013). HGF signaling regulates Claudin-3 dynamics through its C-terminal tyrosine residues. Tissue Barriers, 1(4), e27425. https://doi.org/10.4161/tisb.27425 Uluçkan, Ö., & Wagner, E. F. (2017). Chronic systemic inflammation originating from epithelial tissues. The FEBS Journal, 284(4), 505–516. https://doi.org/10.1111/febs.13904 Vagianos, K., Bector, S., McConnell, J., & Bernstein, C. N. (2007). Nutrition assessment of patients with inflammatory bowel disease. JPEN. Journal of Parenteral and Enteral Nutrition, 31(4), 311–319. https://doi.org/10.1177/0148607107031004311 Van Itallie, C. M., Gambling, T. M., Carson, J. L., & Anderson, J. M. (2005). Palmitoylation of claudins is required for efficient tight-junction localization. Journal of Cell Science, 118(7),

1427–1436. https://doi.org/10.1242/jcs.01735 Van Itallie, Christina M., & Anderson, J. M. (2004). The Molecular Physiology of Tight Junction Pores. Physiology, 19(6), 331–338. https://doi.org/10.1152/physiol.00027.2004 Van Itallie, Christina M, & Anderson, J. M. (2013). Claudin interactions in and out of the tight junction. Tissue Barriers, 1(3), e25247. https://doi.org/10.4161/tisb.25247 Van Zee, K. J., Kohno, T., Fischer, E., Rock, C. S., Moldawer, L. L., & Lowry, S. F. (1992). Tumor necrosis factor soluble receptors circulate during experimental and clinical inflammation and can protect against excessive tumor necrosis factor alpha in vitro and in vivo. Proceedings of the National Academy of Sciences, 89(11), 4845–4849. https://doi.org/10.1073/pnas.89.11.4845 Vincent, T., Neve, E. P. A., Johnson, J. R., Kukalev, A., Rojo, F., Albanell, J., … Fuxe, J. (2009). A SNAIL1-SMAD3/4 transcriptional repressor complex promotes TGF-beta mediated epithelial-mesenchymal transition. Nature Cell Biology, 11(8), 943–950. https://doi.org/10.1038/ncb1905 Wang, F., Daugherty, B., Keise, L. L., Wei, Z., Foley, J. P., Savani, R. C., & Koval, M. (2003). Heterogeneity of claudin expression by alveolar epithelial cells. American Journal of Respiratory Cell and Molecular Biology, 29(1), 62–70. https://doi.org/10.1165/rcmb.2002- 0180OC Wang, Haiming, & Yang, X. (2015). The expression patterns of tight junction protein claudin-1, - 3, and -4 in human gastric neoplasms and adjacent non-neoplastic tissues. International Journal of Clinical and Experimental Pathology, 8(1), 881–887. Retrieved from http://www.ncbi.nlm.nih.gov/pubmed/25755790 Wang, Haitao, Xu, J., Lazarovici, P., Quirion, R., & Zheng, W. (2018). cAMP Response Element- Binding Protein (CREB): A Possible Signaling Molecule Link in the Pathophysiology of Schizophrenia. Frontiers in Molecular Neuroscience, 11, 255. https://doi.org/10.3389/fnmol.2018.00255 Wang, Hongying, Moreau, F., Hirota, C. L., & MacNaughton, W. K. (2010). Proteinase-activated receptors induce interleukin-8 expression by intestinal epithelial cells through ERK/RSK90 activation and histone acetylation. The FASEB Journal, 24(6), 1971–1980. https://doi.org/10.1096/fj.09-137646 Wang, X.-X., Zhang, Y., Li, X.-Y., Li, J., Tang, J.-X., Li, Y.-Y., … Liu, Y.-X. (2019). Kruppel-like factor 6 regulates Sertoli cell blood-testis barrier. Frontiers in Bioscience (Landmark Edition), 24, 1316–1329. Retrieved from http://www.ncbi.nlm.nih.gov/pubmed/31136981 Wang, Yaxi, Sun, T., Sun, H., Yang, S., Li, D., & Zhou, D. (2017). SCF/C-Kit/JNK/AP-1 Signaling Pathway Promotes Claudin-3 Expression in Colonic Epithelium and Colorectal Carcinoma. International Journal of Molecular Sciences, 18(4), 765. https://doi.org/10.3390/ijms18040765 Wang, Ying, & Adjaye, J. (2011). A Cyclic AMP Analog, 8-Br-cAMP, Enhances the Induction of Pluripotency in Human Fibroblast Cells. Stem Cell Reviews and Reports, 7(2), 331–341. https://doi.org/10.1007/s12015-010-9209-3 Wang, Z., Wang, A., Gong, Z., Biviano, I., Liu, H., & Hu, J. (2019). Plasma claudin-3 is associated with tumor necrosis factor-alpha-induced intestinal endotoxemia in liver disease. Clinics and Research in Hepatology and Gastroenterology, 43(4), 410–416. https://doi.org/10.1016/j.clinre.2018.11.014

Watson, C. J., Hoare, C. J., Garrod, D. R., Carlson, G. L., & Warhurst, G. (2005). Interferon- gamma selectively increases epithelial permeability to large molecules by activating different populations of paracellular pores. Journal of Cell Science, 118(Pt 22), 5221–5230. https://doi.org/10.1242/jcs.02630 Weber, L.-M., Fischer, A., Scholz, S., Metzke, D., & Baumgart, D. C. (2017). Impact of Proinflammatory Cytokines and Mesalamine on the Expression of the Tight Junction Protein Claudin-1 in Intestinal Epithelial Cells. Gastroenterology, 152(5), S740. https://doi.org/10.1016/S0016-5085(17)32573-8 Wen, H., Watry, D. D., Marcondes, M. C. G., & Fox, H. S. (2004). Selective Decrease in Paracellular Conductance of Tight Junctions: Role of the First Extracellular Domain of Claudin-5. Molecular and Cellular Biology, 24(19), 8408–8417. https://doi.org/10.1128/MCB.24.19.8408-8417.2004 Winston, B. W., Lange-Carter, C. A., Gardner, A. M., Johnson, G. L., & Riches, D. W. (1995). Tumor necrosis factor alpha rapidly activates the mitogen-activated protein kinase (MAPK) cascade in a MAPK kinase kinase-dependent, c-Raf-1-independent fashion in mouse macrophages. Proceedings of the National Academy of Sciences of the United States of America, 92(5), 1614–1618. https://doi.org/10.1073/pnas.92.5.1614 Wu, J., Peng, D., Zhang, Y., Lu, Z., Voehler, M., Sanders, C. R., & Li, J. (2015). Biophysical characterization of interactions between the C-termini of peripheral nerve claudins and the PDZ₁ domain of zonula occludens. Biochemical and Biophysical Research Communications, 459(1), 87–93. https://doi.org/10.1016/j.bbrc.2015.02.075 Xia, X.-M., Wang, F.-Y., Zhou, J., Hu, K.-F., Li, S.-W., & Zou, B.-B. (2011). CXCR4 antagonist AMD3100 modulates claudin expression and intestinal barrier function in experimental colitis. PloS One, 6(11), e27282. https://doi.org/10.1371/journal.pone.0027282 Xiao, L., Rao, J. N., Cao, S., Liu, L., Chung, H. K., Zhang, Y., … Wang, J.-Y. (2016). Long noncoding RNA SPRY4-IT1 regulates intestinal epithelial barrier function by modulating the expression levels of tight junction proteins. Molecular Biology of the Cell, 27(4), 617–626. https://doi.org/10.1091/mbc.E15-10-0703 Xu, J., Lan, K. T., Su, T. H., Li, Z. F., Wan, Y., Gu, Q. Q., … Zou, X. (2017). [Clinicopathologic and prognosis features of Claudin-low breast cancers]. Zhonghua Bing Li Xue Za Zhi = Chinese Journal of Pathology, 46(9), 634–639. https://doi.org/10.3760/cma.j.issn.0529- 5807.2017.09.009 Xu, X. B., Hu, Y., Wang, Y., Lai, C. J., & Zhang, T. Y. (2014). Expression of claudin-3 in the esophagus and larynx of rat reflux model. Auris, Nasus, Larynx, 41(6), 539–542. https://doi.org/10.1016/j.anl.2014.05.011 Xue, Y., Zhang, H., Sun, X., & Zhu, M.-J. (2016). Metformin Improves Ileal Epithelial Barrier Function in Interleukin-10 Deficient Mice. PloS One, 11(12), e0168670. https://doi.org/10.1371/journal.pone.0168670 Yamaga, K., Murota, H., Tamura, A., Miyata, H., Ohmi, M., Kikuta, J., … Katayama, I. (2018). Claudin-3 Loss Causes Leakage of Sweat from the Sweat Gland to Contribute to the Pathogenesis of Atopic Dermatitis. The Journal of Investigative Dermatology, 138(6), 1279– 1287. https://doi.org/10.1016/j.jid.2017.11.040 Yamashita, M., & Passegué, E. (2019). TNF-α Coordinates Hematopoietic Stem Cell Survival and Myeloid Regeneration. Cell Stem Cell, 25(3), 357-372.e7.

https://doi.org/10.1016/j.stem.2019.05.019 Yang, Y., Cheon, S., Jung, M. K., Song, S. B., Kim, D., Kim, H. J., … Cho, D. (2015). Interleukin-18 enhances breast cancer cell migration via down-regulation of claudin-12 and induction of the p38 MAPK pathway. Biochemical and Biophysical Research Communications, 459(3), 379–386. https://doi.org/10.1016/j.bbrc.2015.02.108 Ying, C., Hong, W., Nianhui, Z., Chunlei, W., Kehe, H., & Cuiling, P. (2019). Nontoxic concentrations of OTA aggravate DON-induced intestinal barrier dysfunction in IPEC-J2 cells via activation of NF-κB signaling pathway. Toxicology Letters, 311, 114–124. https://doi.org/10.1016/j.toxlet.2019.04.021 Yokoyama, M., Narita, T., Sakurai, H., Katsumata-Kato, O., Sugiya, H., & Fujita-Yoshigaki, J. (2017). Maintenance of claudin-3 expression and the barrier functions of intercellular junctions in parotid acinar cells via the inhibition of Src signaling. Archives of Oral Biology, 81, 141–150. https://doi.org/10.1016/j.archoralbio.2017.05.007 Yuan, B., Zhou, S., Lu, Y., Liu, J., Jin, X., Wan, H., & Wang, F. (2015). Changes in the Expression and Distribution of Claudins, Increased Epithelial Apoptosis, and a Mannan-Binding Lectin- Associated Immune Response Lead to Barrier Dysfunction in Dextran Sodium Sulfate- Induced Rat Colitis. Gut and Liver, 9(6), 734–740. https://doi.org/10.5009/gnl14155 Yuan, X., Lin, X., Manorek, G., Kanatani, I., Cheung, L. H., Rosenblum, M. G., & Howell, S. B. (2009). Recombinant CPE fused to tumor necrosis factor targets human ovarian cancer cells expressing the claudin-3 and claudin-4 receptors. Molecular Cancer Therapeutics, 8(7), 1906–1915. https://doi.org/10.1158/1535-7163.MCT-09-0106 Zhang, L., Wang, Y., Zhang, B., Zhang, H., Zhou, M., Wei, M., … Wang, C. (2017). Claudin-3 expression increases the malignant potential of lung adenocarcinoma cells: role of epidermal growth factor receptor activation. Oncotarget, 8(14). https://doi.org/10.18632/oncotarget.14974 Zhang, W.-J., Wang, Y., Yu, K., Duan, J.-Z., Yao, W.-R., Wang, Y., … Yang, G.-B. (2014). Associated changes in the transcription levels of IL-17A and tight junction-associated genes in the duodenal mucosa of rhesus macaques repeatedly exposed to simian/human immunodeficiency virus. Experimental and Molecular Pathology, 97(2), 225–233. https://doi.org/10.1016/j.yexmp.2014.07.007 Zhang, Y., Wu, S., Lu, R., Zhou, D., Zhou, J., Carmeliet, G., … Sun, J. (2015). Tight junction CLDN2 gene is a direct target of the vitamin D receptor. Scientific Reports, 5(1), 10642. https://doi.org/10.1038/srep10642 Zhang, Z., Yu, W., Chen, S., Chen, Y., Chen, L., & Zhang, S. (2018). Methylation of the claudin‑3 promoter predicts the prognosis of advanced gastric adenocarcinoma. Oncology Reports, 40(1), 49–60. https://doi.org/10.3892/or.2018.6411 Zihni, C., Balda, M. S., & Matter, K. (2014). Signalling at tight junctions during epithelial differentiation and microbial pathogenesis. Journal of Cell Science, 127(16), 3401–3413. https://doi.org/10.1242/jcs.145029 Zihni, Ceniz, Mills, C., Matter, K., & Balda, M. S. (2016). Tight junctions: from simple barriers to multifunctional molecular gates. Nature Reviews Molecular Cell Biology, 17(9), 564–580. https://doi.org/10.1038/nrm.2016.80 Zuo, S., Ge, H., Li, Q., Zhang, X., Hu, R., Hu, S., … Feng, H. (2017). Artesunate Protected Blood- Brain Barrier via Sphingosine 1 Phosphate Receptor 1/Phosphatidylinositol 3 Kinase

Pathway After Subarachnoid Hemorrhage in Rats. Molecular Neurobiology, 54(2), 1213– 1228. https://doi.org/10.1007/s12035-016-9732-6

100