Caspase Inhibitor VI

PKCδ–iPLA2–PGE2–PPARγ signaling cascade mediates TNF-α induced Claudin 1 expression in human lung carcinoma cells

Daisuke Iitaka a,b, Serisha Moodley a,c, Hiroki Shimizu a,b, Xiao-Hui Bai a, Mingyao Liu a,c,⁎

a b s t r a c t

Claudin 1 (CLDN1) is a critical component of tight junction adhesion complexes that maintains the structural in- tegrity of epithelial cell layers. Dysregulation of CLDN1 is associated with the growth and metastasis of human lung adenocarcinoma. TNF-α treatment was previously shown to increase expression of CLDN1 that mediated lung cancer cell morphology changes and migration. This study aimed to elucidate the molecular mechanisms involved in TNF-α induced CLDN1 expression in human lung carcinoma A549 cells. Chemical inhibition or siRNA downregulation of Src, PI3K, Akt, MAPKs, NFκB, caspase and PKC demonstrated that PKC, specifically PKCδ, is required for TNF-α induced CLDN1 expression. Further investigation of the PKC pathway revealed that CLDN1 expression is enhanced by the

Keywords: Claudin1 TNF pathway PKC Migration Lung cancer

1. Introduction

The epithelial cell layer is bridged by cell-to-cell adhesions, which consist of two major types of structures, adherens junctions and tight junctions (TJ) [1]. Also known as occluding junctions or zonae occludentes, TJs serve as the “glue” between cells and as barriers to maintain cell polarity [1]. TJs are involved in the transmission of signals from the cell membrane to the nucleus to regulate gene expression [2]. TJs are composed of a branching network of transmembrane proteins that include the claudins and occludins [1]. Claudins (CLDNs) are a family of 24 integral transmembrane proteins that are involved in the formation of tight junctions and in paracellular molecular transport [3–6]. CLDNs directly interact with the zona occludins proteins and actin to promote the structural integrity of tight junctions [6].
As cell adhesion molecules, CLDNs play a role in cell morphology, polarity and motility [7]. Dysregulation of claudin expression has been identified in various cancers, linking claudin dysregulation to epithelial to mesenchymal transition, formation of cancer stem cells and chemoresistance [7,8]. A decrease of CLDN1 was found in glioblastoma, melanoma, breast, esophageal, rectal, ovarian and uterine cancers [7,8]. The loss of CLDN function leads to loss of cell-to-cell adhesion resulting in invasion and metastasis of cancer cells [9]. In lung cancers, low CLDN1 expression is also indicative of lung adenocarcinoma, whereas, increase in CLDN1 expression is correlated with better prognosis and tumor suppressive activity [10,11]. On the other hand, increased expression of certain CLDNs, such as CLDNs 2, 4 and 5, have been implicated in early stage metastasis of cancers and the inhibition of these CLDNs suppresses tumor growth and metastasis [11,12]. CLDN up-regulation is observed in colon cancer, lung squamous cell carcinoma, hepatocellu- lar carcinoma, stomach, prostate and breast cancers [7]. In human lung carcinoma A549 cells, tumor necrosis factor-α TNF-α treatment selec- tively increased expression of CLDN1 that mediated cancer cell mor- phology changes and migration [13]. Thus, CLDN family members may play different roles at different stages of tumorigenesis and metastasis. CLDN expression and function are regulated at multiple levels by various molecules, like the inflammatory cytokines, TNF-α, interferon- γ and interleukin-13 [5,14]. However, the signal transduction pathways that regulate CLDN expression and function are largely unknown. The present study aimed to elucidate the molecular mechanisms involved in TNF-α induced CLDN1 expression in human lung carcinoma A549 cells. A signaling cascade of PKCδ–iPLA2–PGE2–PPARγ was identified in response to TNF-α stimulation for CLDN1 expression. Future investi- gation of this pathway may lead to new targets for anti-lung cancer therapies.

2. Materials and methods

2.1. Cell line, antibodies and other reagents

Human lung adenocarcinoma A549 cells were maintained in DMEM, supplemented with 10% FBS, 1% penicillin–streptomycin, and 1% gluta- mine. Cells were cultured in a standard humidified incubator at 37 °C with 5% CO2. Antibody for CLDN1 was from Invitrogen Corporation (Camarillo, CA). Antibodies for phospho-PKCδ, PKCδ, phospho-cPLA2 and cPLA2 were from Cell Signaling Technology (Danvers, MA). Anti- body for iPLA2 was from Merck Millipore (Billerica, MA). Antibody for GAPDH was from Abcam (Cambridge, UK). Horseradish peroxidase (HRP)-conjugated anti-rabbit secondary antibodies were from Jackson ImmunoResearch Laboratories (West Baltimore Pike, PA). Recombinant human TNF-α were purchased from Austral Biologicals (San Ramon, CA). Actinomycin D, cycloheximide, ammonium chloride (NH4Cl), ALLN, and phorbol 12,13-dibutyrate (PDBu) were from Sigma Aldrich (St. Louis, MO). Pepstatin A, caspase 3 si-RNA and PPARγ si-RNA were from Santa Cruz Biotechnology (Santa Cruz, CA). MG132, CAY10502, FKGK11, methyl arachidonyl fluorophosphonate (MAF), prostaglandin E2 (PGE2), 15-keto PGE2, rosiglitazone, troglitazone and GW6992 were from Cayman Chemical (Ann Arbor, MI). PD98059, SP600125, pyrolidine dithiocarbamate (PDTC) and caffeic acid phenylethyl ester (CAPE), Ly294002, Wortmannin, Akt inhibitor II, PP2, Ro-31-8220, bisindolylmaleimide I (BIM I), Go¨6976, Go¨6983, BAPTA/AM, EGTA/AM and Rottlerin were from EMD Biosciences (Darmstadt, Germany). BOC-D-FMK and z-VAD-FMK were from R&D Systems Inc. (Minneapolis, MN).

2.2. Protein studies and immunoblot analysis

Cells were lysed with modified radioimmunoprecipitation assay buffer (50 mmol/L Tris–HCl, pH 7.5; 150 mmol/L NaCl; 2 mmol/L EGTA; 2 mmol/L EDTA; and 1% Triton X-100) containing 10 mg/mL each of aprotinin, leupeptin, pepstatin, 1 mmol/L phenylmethylsulfonyl fluoride, 1 mmol/L Na3VO4 and 10 mmol/L NaF. Protein concentration was measured by Pierce BCA Protein Assay Kit (Thermo, Rockford, IL). Cell lysates containing equal amounts of total proteins were separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis and then transferred to a nitrocellulose membrane. Non-specific binding sites on the membrane were blocked by incubation with 5% non-fat milk in 150 mM NaCl and 20 mM Tris-HCl, pH 7.6 (TBS) for 1 h at room temperature. The membranes were probed with indicated anti- bodies overnight at 4 °C. After washing with TBS containing 0.1% Tween-20 (TBST), the membrane was incubated with horseradish peroxidase-conjugated goat anti-rabbit IgG for 1 h at room temperature. After washing with TBST, peroxidase activity was detected using SuperSignal West Dura Chemiluminescent Substrate (ThermoScientific, Rockford, IL). The intensities of protein bands were quantified using ImageJ software (NIH).

2.3. SiRNA transfection

Loss-of-function was done using siRNA targeted to caspase 3 or PPARγ, and non-targeted siRNA was used as a control. Each siRNA (50 nmol/L) was transfected into A549 cells using Oligofectamine (Invitrogen, St Louis, MO), according to the manufacturer’s instructions. The medium containing siRNA was replaced with fresh medium after 24 h. Cells were reseeded and treated with different reagents 24 h after cell reseeding.

2.4. Cell proliferation assay

Cell proliferation was evaluated using CellTiter 96® Aqueous One Solution Cell Proliferation Assay (Promega, Madison, WI)) at 0, 8, 18 and 24 h after stimulation. Cells were seeded into 96-well plates and incubated for 24 h prior to the treatments. The media was replaced with 100 μL fresh media containing 20 ng/mL TNF-α. CellTiter 96® AQueous One Solution Reagent (20 μL) was added into each well and the plates were incubated at 37 °C, 5% CO2 for 1 h. The plate was read on a 96 well plate reader (ThermoLab system, Opsys, MR) at 490 nm. The background absorbance of media alone was subtracted from the sample absorbance. The proliferation index was calculated as the mean absorbance of cells at the indicated time point divided by the mean absorbance of cells at 0 h after stimulation and expressed as mean and standard deviation.

2.5. Measurement of PGE2 levels

Cells were plated in 96 well plates and cultured overnight. The next day, cells were treated with TNF-α or PDBu for 2 h. Samples of media were taken and assayed for both intracellular PGE2 and total PGE2 levels according to the manufacturer’s instructions using Amersham Prostaglandin E2 Biotrak Enzymeimmunoassay (EIA) Kit from GE Healthcare (Ann Arbor, MI).

2.6. Wound healing assay

Wounds were created in confluent cells using a pipette tip. The confluent cells were serum starved before wounding. Cells were then treated with 10% FBS, and/or 50 ng/mL TNFα, PGE2, 15-keto PGE2, PDBu, rosiglitazone, troglitazone, BIM1, Ro31-8820, Rottlerin, CAPE, PDTC, ATK, FKGK, MAF and GW6992 Cells were incubated at 37 °C, 5% CO2 for 0, 6, 12 and 18 h. Assays were repeated four times for each condition.

2.7. Statistical analysis

ANOVA and the Student’s t test were used to evaluate continuous variables. Differences were considered significant when the relevant p value was b 0.05. These analyses were performed using statistical soft- ware JMP, version 8 (SAS Institute Inc., Cary, NC).

3. Results

3.1. TNF-α induced CLDN1 expression requires transcriptional and translational activation

In a previous study, we showed that TNF-α induced CLDN1 expres- sion in human lung carcinoma A549 cells after 24 h, 48 h, and 72 h [13]. To determine whether TNF-α treatment can affect CLDN1 expression at earlier time points, we challenged A549 cells with 20 ng/mL TNF-α for 15, 30, 45, 60, 120 and 360 min. TNF-α increased CLDN1 protein expres- sion after 15 min (data not shown). To investigate the mechanisms by which TNF-α induced CLDN1 expression, cells were treated with tran- scription inhibitor, actinomycin D (5 μg/mL), or translation inhibitor, cycloheximide (10 μg/mL). CLDN1 protein expression was significantly inhibited after either 2 h or 24 h stimulation (Fig. 1). Thus, TNF-α in- duced up-regulation of CLDN1 in requires both transcriptional and translational activation.
Lysosomes are responsible for degrading extracellular and trans- membrane proteins, whereas proteasomes degrade intracellular pro- teins [15]. TNF-α induced CLDN1 protein expression at 2 h was upregulated by the lysosome inhibitors, pepstatin a (100 μg/mL) or NH4Cl (10 mM), and by the proteasome inhibitors, MG132 (10 μM) or ALLN (20 μM) (Fig. 1). However, after 24 h, TNF-α in conjunction with MG132, reduced CLDN1 protein expression (Fig. 1). These results indicate that both lysosomes and proteasomes are involved in the regulation of CLDN1 expression and transient inhibition of these mech- anisms prevents CLDN1 degradation. On the other hand, prolonged pro- teasome inhibition by MG132 may have unexpected side effects on A549 cells.

3.2. TNF-α induced CLDN1 expression may be mediated through PKCδ

TNF-α can activate multiple signaling cascades via Src tyrosine kinase, PI3K and MAPK that influence downstream targets, including Akt, JNK and NFκB [16,17]. To investigate which pathway(s) are involved in TNF-α induced CLDN1 expression, cells were pre- incubated with PD98059 (MEK inhibitor, 10 μM), SB203580 (MAPK inhibitor, 10 μM), SP600125 (JNK inhibitor, 25 μM), PDTC (NFκB inhibi- tor, 50 μM), CAPE (NFκB inhibitor, 25 μM), Ly294002 (PI3K inhibitor, 10 μM), Wortmannin (PI3K inhibitor, 200 μM), Akt inhibitor II (10 μM) or PP2 (Src inhibitor, 10 μM) before treatment with TNF-α. TNF-α induced CLDN1 expression was inhibited only by the NFκB inhibitor, CAPE, but not by other inhibitors we tested (Fig. 2). TNF-α stimulation also induces caspase 3 activation by activating the TNF receptor associated death domain (TRADD) signaling cascade [18]. To investigate the relationship between caspase 3 and TNF-α induced CLDN1 expression, a caspase 3 specific inhibitor, BOC-D-FMK (50 μM) or a pan-caspase inhibitor, z-VAD-FMK (50 μM) was used. The caspase inhibitors had no significant effect on TNF-α induced CLDN1 expression, although they effectively blocked TNF-α induced caspase 3 activation (Fig. 3). SiRNA downregulation of caspase 3 also had no effect on TNF-α induced CLDN1 expression.
It has been shown that PKCδ altered the expression of tight junction proteins, including CLDN1, in pancreas, liver, skin and melanoma cell lines [19–22]. Furthermore, previous studies have shown that TNF-α increases PKC activity in A549 cells [23–25]. To test the role of PKC in mediating TNF-α induced CLDN1 expression, cells were treated with TNF-α or with the PKC agonist, PDBu (500 nM). TNF-α treatment in- creased the phosphorylation of PKCδ and CLDN1 expression after 2 h or 24 h, whereas PDBu treatment increased the phosphorylation of PKCδ and CLDN1 expression only after 2 h (Fig. 5A). Phorbol esters, as PKC activators, inhibit PKC activity after prolonged treatment [26]. In human bronchial epithelial BEAS-2B cells, PDBu-induced phosphoryla- tion of PKCδ peaked at 30 min and declined gradually over time [27]. Therefore, short exposure of cells to PDBu increased CLDN1 expression, while prolonged treatment of cells with PDBu inhibited PKC activity and CLDN1 expression.
To identify which PKC isozymes are involved in TNF-α induced CLDN1 expression, cells were pretreated with Bim1 (pan-PKC inhibitor, 2 μM), Ro31-8220 (PKCα, β, γ inhibitor, 3 μM), Gö 6976 (PKCα, β, inhib- itor, 1 μM), Gö 6983 (PKCα, β, γ inhibitor, 1 μM), BAPTA/AM (calcium chelator, 5 μM), EGTA/AM (calcium chelator, 1 μM) or Rottlerin (PKCδ inhibitor, 10 μM) 1 h before co-incubation with TNF-α. TNF-α induced CLDN1 expression was inhibited by the pan-PKC inhibitor, Bim1 and by the PKCδ inhibitor, Rottlerin (Fig. 4). By contrast, neither inhibition of classical PKCs nor chelation of calcium, which is required for classical PKC activation, inhibited TNF-α induced CLDN1 expression (Fig. 4). These results indicate that PKCδ is required for TNF-α induced CLDN1 expression in A549 cells.

3.3. TNF-α induced CLDN1 expression requires PLA2 and PGE2

Phospholipases A2 (PLA2) are a family of enzymes that cleave phospholipids resulting in the release of arachidonic acid, which is modified into inflammation and immunity signaling molecules, prosta- glandins and leukotrienes [28]. TNF-α stimulation increased cytosolic PLA2 (cPLA2) and Ca2+ independent PLA2 (iPLA2) mRNA levels in A549 cells [13]. Furthermore, treatment of A549 cells with a PKC inhibitor blocked TNF-α induced PLA2 protein expression [29]. Hence, we checked the expression of PKCδ, cPLA2 and iPLA2 in A549 cells after TNF-α or PDBu treatment. After 2 h, TNF-α or PDBu increased the expression of PKCδ and its phosphorylation (Fig. 5a). TNF-α or PDBu also increased expression of iPLA2 but not expression of cPLA2 or the phosphorylation of cPLA2 (Fig. 5a).
To determine the role of iPLA2 in TNF-α induced CLDN1 expression, cells were treated with CAY10502 (cPLA2 specific inhibitor, 10 μM), FKGK11 (iPLA2 specific inhibitor, 10 μM) and MAF (pan-PLA2 inhibitor, 10 μM). The iPLA2 and the pan-PLA2 inhibitor blocked TNF-α induced CLDN1 expression, but the cPLA2 inhibitor had no significant effect on CLDN1 expression (Fig. 5b). TNF-α has been shown to induce a signifi- cant increase in prostaglandin E2 (PGE2), a downstream product of arachidonic acid, in A549 cells [30]. Using PGE2 ELISA, we found that the intracellular PGE2 level was significantly increased after 2 h TNF-α or PDBu stimulation, and iPLA2 or pan-PLA2 inhibitors (but not cPLA2 inhibitor) blocked TNF-α or PDBu induced PGE2 production (Fig. 5c). Moreover, TNF-α and PDBu did not affect the total PGE2 level, which is calculated as secreted and cytosolic PGE2 (Fig. 5c). These results indicate that iPLA2 plays an important role in TNF-α or PDBu induced PGE2 production.

3.4. PGE2, 15 keto-PGE2 and PPARγ regulate CLDN1 expression

PGE2 can function as a soluble mediator for intracellular and extra- cellular signaling [30]. However, the half-life of PGE2 is short and PGE2 is rapidly oxidized into 15-keto PGE2, a peroxisome proliferator- activated receptor γ (PPARγ) ligand [31–33]. Stimulate cells with PGE2 (1 μM) increased CLDN1 expression after 2 h but not after 24 h treatment (Fig. 6a). Moreover, A549 cells stimulated with either 15- keto PGE2 or one of the two PPARγ agonists, rosiglitazone (10 μM) or troglitazone (10 μM) increased CLDN1 expression after either 2 h or 24 h (Fig. 6b). Conversely, the PPARγ chemical inhibitor, GW 9662 (50 μM) or PPARγ siRNA reduced both the basal, as well as, the TNFα, PDBu, PGE2, 15-keto PGE2, rosiglitazone or troglitazone-induced CLDN1 expression after either 2 h or 24 h stimulation (Fig. 6c). These results suggest that PGE2 and PPARγ are important mediators of the TNF-α induced CLDN1 expression.

3.5. TNF-α, PGE2, 15 keto-PGE2 and PPARγ agonist promotes cell migration

Our previous study showed that CLDN1 was important for TNF-α induced A549 cell migration [13]. Therefore, we investigated the role of the TNF-α induced PKCδ–iPLA2–PGE2–PPARγ signaling cascade in the promotion of A549 cell migration. In a wound-healing assay, TNF-α, PDBu, PGE2, 15-keto PGE2 and PPARγ agonists significantly promoted wound closure by reducing the cell free area at 6 h, 12 h, 18 h and 24 h after wounding (Fig. 7a). To clarify whether the wound closure was due to increased cell proliferation or cell migra- tion, we measured cell proliferation using an MTS assay. Compared to control cells, TNF-α had no effect on cell proliferation but the other agonists reduced cell proliferation within 24 h of treatment (Fig. 7b). Therefore, the PKCδ–iPLA2–PGE2–PPARγ pathway not only mediates TNF-α induced CLDN1 expression, but also CLDN1 related cell migration.

4. Discussion

In an effort to understand the molecular mechanism that regulates CLDN expression and function, we have identified the PKCδ–iPLA2– PGE2–PPARγ signaling cascade as a new signal transduction pathway for TNF-α induced CLDN1 expression in human lung cancer A549 cells (Fig. 8). Moreover, this signal transduction pathway is directly related to CLDN1 mediated cell migration.
TNF-α signaling is one of the most extensively studied pathways. Upon binding to TNF-α, TNF receptors form trimers resulting in cluster- ing of the intracellular death domains [16,34]. The death domains inter- act with the adaptor protein, TRADD, which serves as a platform to recruit other proteins, such as TNF-associated factor 2 (TRAF-2) or Fas-associated protein with death domain (FADD), and leads to activa- tion of NFκB, MAPK or caspases [16–18,35,36]. In the present study, the NFκB inhibitor CAPE, but not another NFκB inhibitor PDTC, signifi- cantly prevented TNF-α induced CLDN1 expression. CAPE inhibits protein tyrosine kinase mediated NFκB activation, preventing transloca- tion of NFκB to the nucleus and its binding to DNA, resulting in suppres- sion of lipoxygenase/cyclooxygenase pathway; whereas PDTC inhibits the release of IκB from NFκB and NOS synthase mRNA translation resulting in suppression of immunogenic pathways [37–39]. It is possi- ble that NFκB influences transcriptional activity in TNF-α induced CLDN1 regulation in relation to the lipoxygenase/cyclooxygenase path- way. In addition, inhibition of ERK, JNK, PI3K, Akt and Src failed to inhibit TNF-α induced CLDN1 expression. It is interesting to notice that these commonly studied pathways do not play a major role in TNF-α induced CLDN1 expression, suggesting the complexity of TNF-induced signaling. Another possible mechanism of TNF-α induced CLDN1 regulation is through proteolysis. Our results indicate that CLDN1 degradation is mediated through both lysosome and proteasome-dependent mecha- nisms. However, TNF-α treatment did not block either lysosomal or proteasomal degradation of CLDN1. Since our results strongly indicate that TNF-α induces rapid expression of CLDN1, which is regulated at both transcriptional and translational levels, we continued to explore other pathways activated by TNF-α stimulation.
Several studies have shown that TNF-α increases PKC activity in A549 cells [23–25]. The PKC family is divided into classical, novel and atypical PKCs [40]. PKCδ, a member of the novel PKC subfamily, has been shown to alter the expression of tight junction proteins, including CLDN1, in different cell types [20,40]. We demonstrated that TNF-α induced CLDN1 expression is reduced by the PKCδ inhibitor, Rottlerin. Although the specificity and mechanisms of Rottlerin have been questioned [41], it has been commonly used as an effective inhibitor for PKCδ [42]. Moreover, short exposure of TNF-α increases expression of PKCδ and its phosphorylation. Furthermore, the PKC activator, PDBu, increased expression of CLDN1. Previous literature notes that PKC acti- vation rapidly increases PLA2 activity in macrophages and PKC inhibi- tion reduces TNF-α induced PLA2 expression in human bronchial epithelial BEAS-2B cells [29,43]. Similarly, our investigation revealed that both TNF-α and PDBu increased expression of PKCδ, phosphoryla- tion of PKCδ, and the downstream PKC target, iPLA2. Therefore, PKC, specifically the PKCδ, is involved in mediating TNF-α induced CLDN1 expression.
Interestingly, this result echoes the relationship between TNF-α in- duced CLDN1 expression and the lipoxygenase/cyclooxygenase pathway that was hinted at in the NFκB inhibitor data. As we know, PLA2 catalyzes the hydrolysis of phospholipids to release arachidonic acid, which enters the cyclooxgenase pathway to form prostaglandins and the lipoxygensase pathway to form leukotrienes [44]. PLA2 has four main isotypes, including secreted PLA2 (sPLA2), cytosolic calcium dependent PLA2 (cPLA2), calcium independent PLA2 (iPLA2), and lipoprotein-associated PLA2 (lp-PLA2) [44]. Using microarray and bioin- formatics, we previously found that TNFα increases cPLA2 and iPLA2 mRNA levels in A549 cells [13]. In our current study, TNF-α was shown to enhance protein expression of iPLA2 but not cPLA2. To study the downstream effects of TNF-α stimulation on the PKC–cyclooxygen- ase pathway, we investigated the expression of PGE2, which is formed by PGE synthase catalysis of prostaglandin H2 (PGH2) [45]. A previous study showed that either TNF-α or PKC activation induces a rapid increase of PGE2 in A549 cells [30,46]. Moreover, PGE2 was shown to affect the architecture and function of tight junctions and their proteins, including CLDN1 [47]. Our results show that pan-PLA2 and iPLA2 inhib- itors, but not cPLA2 inhibitors, reduced TNF-α or PDBu induced CLDN1 expression, as well as, intracellular PGE2 expression. Furthermore, PGE2 and its oxidized form, 15-keto PGE2, effectively increased CLDN1 expression and promoted cell migration. Thus, CLDN1 expression is reg- ulated by prostaglandin activity in A549 cells.
The oxidized prostaglandin PGE2 metabolite, 15-keto PGE2 is a novel PPARγ ligand [33,48]. The PPARs (peroxisome proliferator- activated receptor) are members of the nuclear receptor superfamily of ligand-inducible transcription factors [49]. The PPARs control the expression of genes involved in adipogenesis, lipid metabolism, inflam- mation and maintenance of metabolic homeostasis [50]. There are three types PPARs, PPARα, PPARβ/δ and PPARγ, and each PPAR isoform has unique functions [48,50]. Since we demonstrated that 15-keto PGE2 is involved in TNF-α induced CLDN1 expression and a previous study showed that rosiglitazone, a PPARγ agonist, induced CLDN1 expression in human nasal epithelial cells, we investigated the role of PPARγ [51]. In the present study, two PPARγ agonists, rosiglitazone and troglitazone, increased CLDN1 expression. Moreover, PPARγ siRNA blocked the CLDN1 expression induced by either TNF-α, PDBu, PGE2, 15-keto PGE2 or the PPARγ agonists. Furthermore, PPARγ agonists increased cell migration, suggesting Caspase Inhibitor VI that PPARγ is a down-stream medi- ator of this signaling cascade for TNF-α induced CLDN1 expression and cell migration (Fig. 8).
The novel signaling cascade that regulates TNF-α induced CLDN1 expression and function should be studied in other cancer cells. This signal pathway may also used by cells to mediate other gene/protein expression. Future studies may elaborate on the effect of TNF-α induced CLDN1 on the epithelial to mesenchymal transition, tumor growth and invasion in vivo, and possible therapeutic intervention related to this signaling pathway.

References

[1] B.N. Giepmans, S.C. van Ijzendoorn, Biochim. Biophys. Acta 1788 (4) (2009) 820–831. http://dx.doi.org/10.1016/j.bbamem.2008.07.015.
[2] M.S. Balda, K. Matter, Trends Cell Biol. 13 (6) (2003) 310–318.
[3] S. Tsukita, M. Furuse, M. Itoh, Nat. Rev. Mol. Cell Biol. 2 (4) (2001) 285–293. http:// dx.doi.org/10.1038/35067088.
[4] K. Morita, M. Furuse, K. Fujimoto, S. Tsukita, Proc. Natl. Acad. Sci. U. S. A. 96 (2) (1999) 511–516.
[5] M.K. Findley, M. Koval, IUBMB life 61 (4) (2009) 431–437. http://dx.doi.org/ 10.1002/iub.175.
[6] M. Heiskala, P.A. Peterson, Y. Yang, Traffic 2 (2) (2001) 93–98.
[7] M.J. Kwon, Int. J. Mol. Sci. 14 (9) (2013) 18148–18180. http://dx.doi.org/10.3390/ ijms140918148.
[8] A.B. Singh, A. Sharma, J.J. Smith, M. Krishnan, X. Chen, S. Eschrich, M.K. Washington,
T.J. Yeatman, R.D. Beauchamp, P. Dhawan, Gastroenterology 141 (6) (2011) 2140–2153. http://dx.doi.org/10.1053/j.gastro.2011.08.038.
[9] T.A. Martin, W.G. Jiang, Biochim. Biophys. Acta 1788 (4) (2009) 872–891. http:// dx.doi.org/10.1016/j.bbamem.2008.11.005.
[10] K. Zhang, H.P. Yao, M.H. Wang, Carcinogenesis 29 (3) (2008) 552–559. http:// dx.doi.org/10.1093/carcin/bgn003.
[11] Y.C. Chao, S.H. Pan, S.C. Yang, S.L. Yu, T.F. Che, C.W. Lin, M.S. Tsai, G.C. Chang, C.H. Wu,
Y.Y. Wu, Y.C. Lee, T.M. Hong, P.C. Yang, Am. J. Respir. Crit. Care Med. 179 (2) (2009) 123–133. http://dx.doi.org/10.1164/rccm.200803-456OC.
[12] P. Michl, C. Barth, M. Buchholz, M.M. Lerch, M. Rolke, K.H. Holzmann, A. Menke, H. Fensterer, K. Giehl, M. Lohr, G. Leder, T. Iwamura, G. Adler, T.M. Gress, Cancer Res. 63 (19) (2003) 6265–6271.
[13] A. Shiozaki, X.H. Bai, G. Shen-Tu, S. Moodley, H. Takeshita, S.Y. Fung, Y. Wang, S. Keshavjee, M. Liu, PLoS One 7 (5) (2012) e38049. http://dx.doi.org/10.1371/ journal.pone.0038049.
[14] T. Kinugasa, T. Sakaguchi, X. Gu, H.C. Reinecker, Gastroenterology 118 (6) (2000) 1001–1011.
[15] J. Adams, Nat. Rev. Cancer 4 (5) (2004) 349–360. http://dx.doi.org/10.1038/ nrc1361.
[16] W.M. Chu, Cancer Lett. 328 (2) (2013) 222–225. http://dx.doi.org/10.1016/ j.canlet.2012.10.014.
[17] Y. Wu, B.P. Zhou, Br. J. Cancer 102 (4) (2010) 639–644. http://dx.doi.org/10.1038/ sj.bjc.6605530.
[18] H. Hsu, H.B. Shu, M.G. Pan, D.V. Goeddel, Cell 84 (2) (1996) 299–308.
[19] D. Kyuno, T. Kojima, T. Ito, H. Yamaguchi, M. Tsujiwaki, A. Takasawa, M. Murata, S. Tanaka, K. Hirata, N. Sawada, Cell Tissue Res. 346 (3) (2011) 369–381. http:// dx.doi.org/10.1007/s00441-011-1287-2.
[20] C.H. Yoon, M.J. Kim, M.J. Park, I.C. Park, S.G. Hwang, S. An, Y.H. Choi, G. Yoon, S.J. Lee, J. Biol. Chem. 285 (1) (2010) 226–233. http://dx.doi.org/10.1074/ jbc.M109.054189.
[21] Y.K. Lin, H.W. Chen, Y.L. Leu, Y.L. Yang, Y. Fang, J.H. Su Pang, T.L. Hwang, J. Ethnopharmacol. 145 (2) (2013) 614–620. http://dx.doi.org/10.1016/ j.jep.2012.11.044.
[22] P.D. Leotlela, M.S. Wade, P.H. Duray, M.J. Rhode, H.F. Brown, D.T. Rosenthal, S.K. Dissanayake, R. Earley, F.E. Indig, B.J. Nickoloff, D.D. Taub, O.P. Kallioniemi, P. Meltzer, P.J. Morin, A.T. Weeraratna, Oncogene 26 (26) (2007) 3846–3856. http:// dx.doi.org/10.1038/sj.onc.1210155.
[23] C.C. Chen, C.Y. Chou, Y.T. Sun, W.C. Huang, Cell. Signal. 13 (8) (2001) 543–553. http://dx.doi.org/10.1016/S0898-6568(01)00171-1.
[24] C.J. Clarke, J.M. Guthrie, Y.A. Hannun, Mol. Pharmacol. 74 (4) (2008) 1022–1032. http://dx.doi.org/10.1124/mol.108.046250.
[25] C.H. Woo, J.H. Lim, J.H. Kim, Am. J. Physiol. Lung Cell. Mol. Physiol. 288 (2) (2005) L307–L316. http://dx.doi.org/10.1152/ajplung.00105.2004.
[26] Z. Lu, D. Liu, A. Hornia, W. Devonish, M. Pagano, D.A. Foster, Mol. Cell. Biol. 18 (2) (1998) 839–845.
[27] H. Xiao, X.H. Bai, A. Kapus, W.Y. Lu, A.S. Mak, M. Liu, Mol. Cell. Biol. 30 (23) (2010) 5545–5561. http://dx.doi.org/10.1128/MCB. 00382-10.
[28] M. Murakami, I. Kudo, J. Biochem. 131 (3) (2002) 285–292.
[29] T. Wu, T. Ikezono, C.W. Angus, J.H. Shelhamer, Biochim. Biophys. Acta 1310 (2) (1996) 175–184.
[30] B.J. Pettus, J. Bielawski, A.M. Porcelli, D.L. Reames, K.R. Johnson, J. Morrow, C.E. Chalfant, L.M. Obeid, Y.A. Hannun, FASEB J. 17 (11) (2003) 1411–1421. http:// dx.doi.org/10.1096/fj.02-1038com.
[31] N. Eguchi, H. Hayashi, Y. Urade, S. Ito, O. Hayaishi, J. Pharmacol. Exp. Ther. 247 (2) (1988) 671–679.
[32] P.J. Henry, A. D’Aprile, G. Self, T. Hong, T.S. Mann, J. Pharmacol. Exp. Ther. 314 (3) (2005) 995–1001. http://dx.doi.org/10.1124/jpet.105.086124.
[33] W.L. Chou, L.M. Chuang, C.C. Chou, A.H. Wang, J.A. Lawson, G.A. FitzGerald, Z.F. Chang, J. Biol. Chem. 282 (25) (2007) 18162–18172. http://dx.doi.org/10.1074/ jbc.M702289200.
[34] H. Wajant, K. Pfizenmaier, P. Scheurich, Cell Death Differ. 10 (1) (2003) 45–65. http://dx.doi.org/10.1038/sj.cdd.4401189.
[35] H. Wajant, Essays Biochem. 39 (2003) 53–71.
[36] G. Chen, D.V. Goeddel, Science 296 (5573) (2002) 1634–1635. http://dx.doi.org/ 10.1126/science.1071924.
[37] K. Natarajan, S. Singh, T.R. Burke, D. Grunberger, B.B. Aggarwal, Proc. Natl. Acad. Sci. U. S. A. 93 (17) (1996) 9090–9095. http://dx.doi.org/10.1073/pnas.93.17.9090.
[38] R. Schreck, B. Meier, D.N. Mannel, W. Droge, P.A. Baeuerle, J. Exp. Med. 175 (5) (1992) 1181–1194. http://dx.doi.org/10.1084/jem.175.5.1181.
[39] J.M. Griscavage, S. Wilk, L.J. Ignarro, Proc. Natl. Acad. Sci. U. S. A. 93 (8) (1996) 3308–3312.
[40] H. Mellor, P.J. Parker, Biochem. J. 332 (Pt 2) (1998) 281–292.
[41] S.P. Soltoff, Trends Pharmacol. Sci. 28 (9) (2007) 453–458. http://dx.doi.org/ 10.1016/j.tips.2007.07.003.
[42] M. Gschwendt, H.J. Muller, K. Kielbassa, R. Zang, W. Kittstein, G. Rincke, F. Marks, Biochem. Biophys. Res. Commun. 199 (1) (1994) 93–98. http://dx.doi.org/ 10.1006/bbrc.1994.1199.
[43] Z.H. Qiu, C.C. Leslie, J. Biol. Chem. 269 (30) (1994) 19480–19487.
[44] J.E. Burke, E.A. Dennis, J. Lipid Res. 50 (2009) S237–S242. http://dx.doi.org/10.1194/ jlr. R800033-JLR200 (Suppl.).
[45] J.Y. Park, M.H. Pillinger, S.B. Abramson, Clin. Immunol. 119 (3) (2006) 229–240. http://dx.doi.org/10.1016/j.clim.2006.01.016.
[46] M.S. Chang, B.C. Chen, M.T. Yu, J.R. Sheu, T.F. Chen, C.H. Lin, Cell. Signal. 17 (3) (2005) 299–310. http://dx.doi.org/10.1016/j.cellsig.2004.07.008.
[47] M.N. Tanaka, B.L. Diaz, W. de Souza, J.A. Morgado-Diaz, BMC Cell Biol. 9 (2008) 63. http://dx.doi.org/10.1186/1471-2121-9-63.
[48] T. Shiraki, N. Kamiya, S. Shiki, T.S. Kodama, A. Kakizuka, H. Jingami, J. Biol. Chem. 280 (14) (2005) 14145–14153. http://dx.doi.org/10.1074/jbc.M500901200.
[49] R.M. Evans, G.D. Barish, Y.X. Wang, Nat. Med. 10 (4) (2004) 355–361. http:// dx.doi.org/10.1038/nm1025.
[50] G.D. Barish, V.A. Narkar, R.M. Evans, J. Clin. Invest. 116 (3) (2006) 590–597. http:// dx.doi.org/10.1172/JCI27955.
[51] N. Ogasawara, T. Kojima, M. Go, T. Ohkuni, J. Koizumi, R. Kamekura, T. Masaki, M. Murata, S. Tanaka, J. Fuchimoto, T. Himi, N. Sawada, Pharmacol. Res. 61 (6) (2010) 489–498. http://dx.doi.org/10.1016/j.phrs.2010.03.002.