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Vol. 20, Issue 7, 1916-1925, April 1, 2009
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Epithelial Pathobiology Research Unit, Department of Pathology and Laboratory Medicine, Emory University School of Medicine, Atlanta, GA 30322
Submitted October 9, 2008;
Revised January 2, 2009;
Accepted January 22, 2009
Monitoring Editor: Ben Margolis
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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; Laukoetter et al., 2007), endothelial cell migration (Naik et al., 2003a; Bazzoni et al., 2005), angiogenesis (Naik et al., 2003a), platelet aggregation (Babinska et al., 2002), and leukocyte adhesion (Martin-Padura et al., 1998; Del Maschio et al., 1999). Recent reports suggest that JAM-A not only directly decreases paracellular permeability (Mandell et al., 2005; Laukoetter et al., 2007) but also promotes epithelial cell migration (Severson et al., 2008), which is critical for the maintenance of intestinal barrier in health and disease (Lacy, 1988; Fenteany et al., 2000). The relationship between these functions and JAM-A structure remains poorly understood.
The mature JAM-A protein is composed of two extracellular immunoglobulin (Ig)-like loops, a membrane-spanning segment, and a cytoplasmic tail ending in a postsynaptic density 95/disc-large/zona occludens (PDZ) binding motif. The membrane distal most extracellular Ig loop mediates homodimerization between JAM-A proteins on the same cell (Prota et al., 2003; Guglielmi et al., 2006; Severson et al., 2008) (cis) and potentially mediates interactions between JAM-A molecules on adjacent cells (Kostrewa et al., 2001) (trans). Cis-dimerization has been shown to be necessary for JAM-A regulation of epithelial cell invasion (Severson et al., 2008) and epithelial barrier recovery (Liu et al., 2000). The PDZ binding motif of JAM-A has been reported to associate with the PDZ domain-containing proteins Afadin and ZO-1 in endothelial and epithelial cells (Yamamoto et al., 1997; Ebnet et al., 2000). However, the signaling events downstream of JAM-A dimerization that mediate these functional effects are not understood.
Based on our previous findings (Severson et al., 2008), we hypothesized that cis-dimerization of JAM-A results in close apposition of JAM-A PDZ binding motifs that facilitates interaction between scaffolding proteins, which in turn activates signaling molecules. However, biologically relevant scaffolding and signaling proteins downstream of the dimerized JAM-A complex have not yet been defined.
In this study, we validated the above-mentioned hypothesis that JAM-A dimerization and its association with specific scaffolding proteins results in activation of signaling molecules that regulate epithelial cell migration. We demonstrate 1) coassociation of the Rap1 guanine nucleotide exchange factor PDZ-GEF2 and Afadin with JAM-A; 2) Afadin and PDZ-GEF2 association requires JAM-A expression; 3) decreased expression of JAM-A, Afadin, PDZ-GEF2, or Rap1A results in analogous decreased β1 integrin protein levels and reduced cell migration; 4) and PDZ-GEF2 activates Rap1A to regulate β1 integrin and cell migration. Our results support the hypothesis that JAM-A dimerization is required for the assembly of a protein complex containing PDZ-GEF2 and Afadin, which activates Rap1A, thereby stabilizing β1 integrin protein levels and regulating cell migration. These findings demonstrate that JAM-A regulates cell migration and, by extension, implicates JAM-A as a key molecule that facilitates wound healing after physical or inflammatory injury.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Immunoblots
Monolayers of epithelial cells that were between 60 and 90% confluent were homogenized in radioimmunoprecipitation assay (RIPA) lysis buffer containing 20 mM Tris, 50 mM NaCl, 2 mM EDTA, 2 mM EGTA, 1% sodium deoxycholate, 1% Triton X-100, and 0.1% SDS, pH 7.4. Lysis buffer was supplemented with protease and phosphatase inhibitor cocktails from Sigma-Aldrich (1:100 dilution). Protein concentrations in lysates were quantified by bicinchoninic acid assay. Lysates were cleared by centrifugation and immediately boiled in reducing SDS sample buffer. Additionally, isolated colonic epithelial cells from JAM-A–deficient and control mice were analyzed for β1 integrin and Rap1 by immunoblot analysis for three animals. Distal colonic epithelial cell lysates were prepared after the serosa and external longitudinal layer of the muscularis propria were stripped away. Isolated epithelial sheets were subsequently lysed in RIPA buffer. Mice were genotyped and raised as reported previously (Laukoetter et al., 2007). SDS-polyacrylamide gel electrophoresis (PAGE) and immunoblots were performed by standard methods. Tubulin was used as a protein loading control. Each immunoblot shown is representative of at least three independent experiments.
Immunoprecipitations (IPs)
Cells were grown to between 60 and 90% confluence in T-25 flasks (Corning) and then lysed with 1 ml of RIPA buffer and dounced. Samples were then centrifuged to remove cell debris, and 100 µl of supernatant was saved for input sample. The remaining supernatant was used for IPs. Sample was precleared for 45 min at 4C with Sepharose beads followed by incubation for 1 h at 4C with protein G-coupled to Sepharose (GE Healthcare, Chalfont St. Giles, Buckinghamshire, United Kingdom). Beads were washed three times with sample buffer. Beads were boiled for 15 min with SDS sample buffer, and the entire sample was loaded for analysis by immunoblot. Each IP was repeated at least three times.
Immunofluorescence (IF) Microscopy
Cells were grown on 0.3-µm pore transwell filters (Corning Life Sciences, Lowell, MA), fixed in 100% ethanol at –20°C for 20 min, and blocked in 1% BSA in HBSS+ for 1 h. Primary antibodies were diluted in blocking buffer and incubated with cells for 1 h at 25°C. The cells were washed in HBSS+ and then incubated in fluorescently labeled secondary antibodies for 45 min at room temperature. Labeled cells were then washed and mounted in Prolong Antifade Agent (Invitrogen, Carlsbad, CA). A laser scanning microscope (Carl Zeiss, Jena, Germany) was used to capture confocal fluorescence images.
DIC Microscopy
Pictures were obtained using an Axiovert 35 light microscope at 5x power by using the DIC 0.4 filter and saved using Axiomatic imaging software (Carl Zeiss). Images were then imported directly into Adobe Photoshop (Adobe Systems, Mountain View, CA) and saved as TIFF files for figures.
Antibodies
The murine monoclonal anti-JAM-A antibody J10.4 was described previously (Liu et al., 2000), and the rabbit polyclonal anti-PDZ-GEF was a kind gift from Dr. N. Mochizuki (Sakurai et al., 2006). Other antibodies were commercially purchased: polyclonal rabbit anti-JAM-A (Zymed Laboratories, South San Francisco, CA), monoclonal mouse anti-Afadin (BD Transduction Laboratories, Lexington, KY), monoclonal mouse anti-tubulin (Sigma-Aldrich), polyclonal rabbit anti-actin (Sigma-Aldrich), rabbit polyclonal Rap1 (Millipore, Billerica, MA), monoclonal rabbit anti-β1 integrin (Novus Biologicals, Littleton, CO), monoclonal rabbit anti-JAM-A (Novus Biologicals), polyclonal rabbit ZO-1 (Zymed Laboratories), polyclonal rabbit anti-FLAG (Sigma-Aldrich), goat anti-rabbit-horseradish peroxidase (HRP) (Jackson ImmunoResearch Laboratories, West Grove, PA), and goat anti-mouse-HRP (Jackson ImmunoResearch Laboratories).
Small Interfering RNA (siRNA) Experiments and DNA Transfection
For siRNA protein targets, four to 10 oligonucleotides were designed for each target, and protein down-regulation was verified by immunoblot. For each candidate protein tested, two promising siRNA targets that led to effective protein down-regulation were identified and were then separately tested for effects on β1 integrin levels and cell migration. Two separate oligonucleotides for each candidate molecule were tested to diminish the possibility of siRNA-mediated interferon and off-target effect and to increase the specificity of findings (Echeverri et al., 2006). The two siRNA oligomers that resulted in the greatest decrease in protein expression were combined and used for the experiments presented in this report. siRNA oligonucleotides were obtained from Dharmacon RNA Technologies (Lafayette, CO) or QIAGEN (Valencia, CA). Controls included a scramble control from QIAGEN (AllStars Negative Control) as well as mock-transfected controls. All targets used are listed in Table 1. Every transfection was done at a total concentration of 50 nM siRNA. Transfections were performed using HiPerFect (QIAGEN) according to the manufacturer's instructions. Assays were performed 72 h after transfection.
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). Cells were used in assays 48 h after transfection.
Rap1 Activity Assay
A Rap1 activity assay was performed according to the manufacturer's instructions (Millipore). Briefly, cells were lysed in a Tris and Triton X-based lysis buffer at 4°C. Cell debris was removed by centrifugation, and 100 µl of sample was saved as input to determine total Rap1 levels. Then, 50 µg of protein for each sample was incubated at 4°C for 45 min with Ral-GDS agarose beads to bind active Rap1. Beads were washed three times with the lysis buffer followed by boiling in SDS sample buffer. The entire sample was analyzed by immunoblot with detection using a polyclonal Rap1 antibody from Millipore. Each activity assay was repeated at least three times.
Surface Biotinylation
Cells were treated with siRNA as described above. At 72 h after treatment, cells were placed on ice, rinsed with phosphate-buffered saline, and then cells were treated with EZ-link-Sulfo-NHS-LC-Biotin (Pierce Chemical, Rockford, IL) according to the manufacturer's protocol. Cells were lysed as described above, and ImmunoPure Immobilized Monomeric Avadin (Pierce Chemical) was then used to isolate the labeled surface proteins. Proteins were removed from the Avadin beads by boiling in SDS sample buffer followed by Western blot and probing for β1 integrin.
Scratch Wound Assay
Cells were seeded in 24-well plates at a density resulting in 50–70% confluence 24 h later. Cells were treated with siRNA 24 h after seeding. Wounds were made in confluent monolayers 48 h after siRNA treatment by using a pipet tip under vacuum suction. A straight line was drawn across the bottom of each well, and then pictures were taken with the line at the bottom of the viewing field. Wounds were measured from the exact vertical middle of each picture such that the initial measurement and the measurement 16 h later were taken from the same vertical spot in each well. The average starting wound width was 702 ± 51 µm, and there was no statistical difference in the sizes of initial wound widths in any of the experiments. Representative images are shown for each figure, and results are reported as percentage of wound closed. Percentage of wound closure is equal to (initial width – width at 16 h)/initial width. Measurements were taken in pixels, which directly correlate to distance in a linear manner. Data are from a representative experiment of three independent experiments with four sample replicates per experiment.
Constructs and Cloning
A PDZ-GEF2 construct was cloned from SK-CO15 cDNA. mRNA from SK-CO15 cells, which were 75% confluent, was isolated using TRIzol (Invitrogen). cDNA was then produced using the first-strand cDNA synthesis kit (Invitrogen). PDZ-GEF2 was amplified by polymerase chain reaction (PCR) with Phusion (New England Biolabs, Ipswich, MA) polymerase and an N-terminal FLAG tag, and BamHI and XhoI restriction sites were added in the primers. The forward primer containing the FLAG tag was as follows: 5'-aataaagcttgccaccATGGATTACAAGGACGACGATGACAAGATGAACTCACCCGTGGACCCT-3'. The reverse primer was 5'-TTCAAGAAATGTCCTGTAAGTT-3'. The PCR product was inserted into pCDNA3.0 by using the engineered restriction sites and sequence was verified.
Reverse Transcription (RT)-PCR
RT-PCR was performed using the SuperScript III one-step PCR kit from Invitrogen according to the manufacturer's instructions. The annealing Tm used for all primers was 57°C. The PDZ-GEF1 and two primers used have been described and verified previously (Schultess et al., 2005).
Real-Time PCR
Quantitative real-time PCR was performed to determine mRNA levels. PCR amplification was performed using the GeneAmp 5700 sequence detection system (Applied Biosystems, Foster City, CA). PCR was run using the following protocol: initial activation at 94°C for 3 min, 40 cycles of 94°C for 15 s, 52°C for 30 s, and 72°C for 30 s. Each sample was performed in triplicate. Direct detection of PCR product was monitored in real-time by measuring the increase in fluorescence caused by the binding of SYBR Green I Dye (Invitrogen) to double-stranded DNA, by using the ABI Gene Amp 5700 sequence detection system (Applied Biosystems). Results of the real-time PCR data were represented as -fold change, which was determined by applying the formula 2
Ct, where Ct = Ct of target gene – Ct of endogenous control gene (signal regulatory peptide), and Ct = Ct of samples for target gene (siRNA treated) – Ct of the control for the target gene (mock siRNA treated). The primers used were as follows: Rap1A.F: 5'-TGGATACTGCAGGGACAGAGCAAT-3'; Rap1A.R: 5'-ACATCTTCCGTGTCCTTAACCCGT-3'; Rap1B.F: 5'-AGGCGTTGGAAAGTCTGCTTTGAC-3'; Rap1B.R: 5'-ATTGCTCCGTTCCTGCAGTATCCA-3'; PDZ-GEF1.F: 5'-AAATTCGTCACGTTGGCCGAATGG-3'; PDZ-GEF1.R: 5'-ACTCCGCCATTTCTTCTTCCGAGT-3'; PDZ-GEF2.F: 5'-TGTTGACTCCATGTCTGCAGCTCT-3'; and PDZ-GEF2.R: 5'-ACCCAGGGCCATGTTGACTATGAT-3'.
Statistics
For comparison of samples in experiments with only two groups, a Student's t test was used. One-way analysis of variance was used for comparisons in experiments with greater than two groups with post hoc analysis performed by GraphPad (GraphPad Software, San Diego, CA) to determine p values for sample groups compared with controls. p < 0.05 was considered significant in either case.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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; Ebnet et al., 2001; Severson et al., 2008), and this presumably requires JAM-A interactions with PDZ domain-containing scaffolding proteins. To identify scaffolding proteins that mediate JAM-A function, we focused on the candidate proteins Afadin and ZO-1, which have previously been reported to associate with JAM-A (Ebnet et al., 2000; Fukuhara et al., 2002). Specifically, we investigated whether these proteins are required for JAM-A regulation of cell migration.
JAM-A, ZO-1, and Afadin were immunolocalized in a model intestinal epithelial cell line SKCO-15. ZO-1 is a tight junction (TJ) protein, but Afadin has been reported to associate with Nectin(s) in adherens junctions (AJs) (Miyahara et al., 2000) and has also been observed to localize at both the TJs and AJs by electron microscopy (Yamamoto et al., 1997) and subcellular fractionation experiments (Vogelmann and Nelson, 2005). As shown in Figure 1A and Supplemental Figure 1A, JAM-A, Afadin, and ZO-1 colocalized at TJs.
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). Immunoprecipitated protein complexes were isolated using protein G-Sepharose followed by boiling in SDS-containing sample buffer. Supernatants were then subjected to SDS-PAGE followed by immunoblotting. Immunoblots were probed for JAM-A, ZO-1, and Afadin. As shown in Figure 1B Afadin coimmunoprecipitates with JAM-A, whereas ZO-1 was not detected in the protein complex (Supplemental Figure 1B). Although the conditions used in this experiment may have disrupted a potential interaction between ZO-1 and JAM-A, these data suggest that JAM-A coimmunoprecipitates with Afadin in a manner independent of potential interactions between JAM-A and ZO-1 (Figure 1B).
It has been reported previously that JAM-A regulates epithelial and endothelial cell migration (Naik et al., 2003a; Bazzoni et al., 2005; Severson et al., 2008). To confirm that JAM-A regulates cell migration in SKCO-15 cells, we evaluated the effect of decreased JAM-A protein levels on wound closure in a scratch wound assay. siRNA was used to decrease JAM-A protein expression in SKCO-15 cells, which are amenable to transfection. JAM-A siRNA treatment consistently resulted in a >80% decrease in JAM-A protein expression compared with transfection with scrambled siRNA controls as shown in Figure 2A. siRNA-mediated down-regulation of JAM-A protein expression inhibited wound closure compared with mock- or scramble siRNA-transfected cells (p < 0.01) (Figure 2, B and C). These results are consistent with our previous observations demonstrating that overexpression of JAM-A dimerization-defective mutants or JAM-A mutants lacking the PDZ binding motif decreases the rate of epithelial cell migration across permeable filters (Severson et al., 2008).
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), we reasoned that proteins which interact with this motif mediate the functional effects. To determine whether either Afadin or ZO-1 mediates the effects of JAM-A on cell migration, analogous experiments were performed using siRNA to decrease the expression of Afadin and ZO-1. As shown in Figure 2, siRNA-mediated down-regulation of Afadin expression, but not ZO-1, inhibited wound closure at 16 h (p < 0.001). These effects are likely due to changes in the rate of cell migration and not secondary to alterations in cell proliferation as the time course for these experiments (16 h) was sufficiently short as to minimize potential effects from proliferation (doubling time of SKCO-15 cells is 48 h). Because decreased expression of JAM-A and Afadin, but not ZO-1, decreased cell migration, and JAM-A was observed to associate in a protein complex with Afadin, these data are consistent with JAM-A signaling through Afadin to regulate cell migration.
JAM-A and Afadin Influence Cell Migration through Regulation of β1 Integrin Protein Levels
We reported previously that JAM-A regulation of cell migration is mediated by effects on β1 integrin protein levels. The decreased epithelial cell migration, which was observed after overexpression of dominant-negative JAM-A, was rescued by overexpression of β1 integrin (Severson et al., 2008). To evaluate whether Afadin regulates cell migration downstream of JAM-A, we analyzed β1 integrin protein levels after decreased expression of either protein. Diminished expression of JAM-A or Afadin, but not ZO-1, resulted in decreased levels of β1 integrin protein (Figure 3 and Supplemental Figure 1C). These findings suggest that JAM-A signals through Afadin to effect cell migration by regulating cellular levels of β1 integrin protein.
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; Severson et al., 2008). In in vitro studies, Rap1 has been reported to regulate integrin levels in epithelial cells (Mandell et al., 2005) as well as integrin activation in leukocytes (Reedquist et al., 2000). To verify these results in vivo, we determined the influence of JAM-A on β1 integrin protein levels and Rap1 activation in intestinal epithelial cells derived from JAM-A knockout mice. In the colonic epithelial cells from JAM-A knockout mice, β1 integrin protein levels were decreased by 50% compared with wild-type controls (p < 0.05) (Figure 4). Colonic epithelial cell lysates were incubated with Ral-GDS agarose beads to bind active Rap1, followed by SDS-PAGE. Immunoblots were then probed for Rap1. Total levels of Rap1 were unchanged compared with wild-type controls; however, active Rap1 levels were decreased by 49% (p < 0.05) (Figure 4). These results suggest that JAM-A regulates Rap1 activity and β1 integrin levels both in vitro and in vivo.
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). Because Rap1 consists of two closely related genes (paralogues), Rap1A and Rap1B, we performed experiments examining the individual roles of Rap1A and Rap1B in JAM-A/Afadin mediated regulation of β1 integrin levels and cell migration. To isolate the individual effects of Rap1A and Rap1B, siRNA specific to the mRNA sequences in the 3' untranslated region of Rap1A and Rap1B were used, because reliable and specific antibodies are not available. As shown in Figure 5B, siRNA-mediated down-regulation of Rap1A and Rap1B resulted in decreased expression of Rap1 as assessed by a pan-Rap1 antibody. By real-time RT-PCR, siRNA-mediated decreases in Rap1A or Rap1B reduced mRNA expression by 93 and 78%, respectively (Figure 5C). Interestingly, siRNA-mediated down-regulation of Rap1A, but not Rap1B, resulted in decreased β1 integrin levels. Furthermore, decreased expression of Rap1A, but not Rap1B, reduced cell migration compared with controls (Figure 5, D and E). These findings suggest that JAM-A and Afadin activate Rap1A, which in turn stabilizes β1 integrin levels and promotes cell migration.
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). We proposed a model of JAM-A function involving dimerization of JAM-A that brings scaffolding proteins into proximity to facilitate downstream signaling. Because our results suggest that JAM-A coassociates with Afadin, and decreased expression of either protein has similar functional and biochemical effects, we hypothesize that JAM-A and Afadin are part of a complex that activates Rap1A. Interestingly, Afadin has been shown to directly associate with Rap1A (Linnemann et al., 1999; Hoshino et al., 2005). Furthermore, activation of Rap1A is dependent on guanine nucleotide exchange factors (GEFs). Given the dependence of JAM-A function on dimerization, we reasoned that a likely additional component of the JAM-A–associated signaling complex would be a Rap1-specific GEF. Experiments were thus performed to identify GEF molecules that contain PDZ binding domains and potentially coassociate with JAM-A. By RT-PCR (data not shown) and immunoblot, we observed expression of two Rap1-specific GEFs that contain PDZ binding domains, PDZ-GEF1 and PDZ-GEF2 (Rebhun et al., 2000), in SKCO-15 cells (Figure 6A). We next used siRNA to selectively decrease expression of PDZ-GEF1 and PDZ-GEF2 to determine antibody specificity. As detected by immunoblot, decreased expression of PDZ-GEF1 or PDZ-GEF2 revealed a partial reduction in total PDZ-GEF expression (Figure 6A). Specificity of siRNA mediated down-regulation was confirmed by real time RT-PCR, which revealed that PDZ-GEF1 and PDZ-GEF2 mRNA were each specifically decreased by 77 and 90% respectively (Figure 6B). To determine whether either protein was involved in JAM-A signaling, we performed additional siRNA experiments. As shown in Figure 6, A, D and E, decreased expression of PDZ-GEF2, but not PDZ-GEF1, resulted in decreased β1 integrin levels and decreased cell migration. Interestingly, down-regulation of either PDZ-GEF1 or PDZ-GEF2 was associated with significantly decreased total Rap1 activity (*p < 0.05). Notably, a larger decrease in total Rap1 activity was observed when expression of PDZ-GEF1 and PDZ-GEF2 were both simultaneously decreased (Figure 6C). These data suggest that PDZ-GEF2 and not PDZ-GEF1 activates Rap1A to stabilize β1 integrin levels and promote cell migration, whereas PDZ-GEF1 may activate Rap1B.
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JAM-A Coassociates with PDZ-GEF2
Given that PDZ-GEF2 contains a PDZ domain that has the potential to interact with the PDZ binding motif on JAM-A in concert with identical functional effects after decreased expression of either protein, experiments were performed to determine whether JAM-A and PDZ-GEF2 associate in a signaling complex. In particular, because similar phenotypes were seen with loss of JAM-A, Afadin, Rap1A, and PDZ-GEF2, we hypothesized that PDZ-GEF2 may form a complex with dimerized JAM-A and Afadin that regulates activation of Rap1A. Decreased expression of any of the components of such a complex would be predicted to reduce the levels of active Rap1A and lead to the functional and biochemical changes observed in this report. To test this hypothesis, colocalization and coimmunoprecipitation studies of JAM-A with PDZ-GEF2 were performed.
As shown in Figure 7A, JAM-A and PDZ-GEF colocalize at cell–cell junctions as observed by immunofluorescence labeling in SKCO-15 cells at the XZ and XY planes. In experiments using JAM-A mAb J10.4, coimmunoprecipitation of JAM-A with endogenous PDZ-GEF was observed (Figure 7B). Furthermore, to determine whether Afadin was also associated in a complex with PDZ-GEF2, coimmunoprecipitation experiments with an anti-Afadin antibody were performed in SKCO-15 cells transfected with FLAG-tagged PDZ-GEF2. The model based on our findings predicts that the association of PDZ-GEF2 with Afadin would be dependent on JAM-A. To directly test this hypothesis, we treated cells with control siRNA or with siRNA targeting JAM-A expression. As shown in Figure 7C, Afadin coimmunoprecipitated with JAM-A and FLAG-PDZ-GEF2 in lysates from control cells that express JAM-A. However, in cells with decreased JAM-A expression, FLAG-PDZ-GEF2 did not coimmunoprecipitate with Afadin. We hypothesized that JAM-A associated with the PDZ domain of PDZ-GEF2 through either direct or indirect interactions with linker scaffolding proteins. To further test this hypothesis, a truncated FLAG-PDZ-GEF construct was made in pCDNA3.0 that contained the putative PDZ binding domain of PDZ-GEF2 (amino acids 530-609). As can be seen in Figure 7D, from lysates of cells transfected with the tagged PDZ binding domain-containing construct, anti-FLAG antibody successfully immunoprecipitated JAM-A. These findings lend further support to a role of the PDZ domain of PDZ-GEF2 in mediating association of PDZ-GEF2 with JAM-A. Whether this association occurs by direct or indirect means awaits further binding studies with purified recombinant proteins. Our combined findings provide evidence that JAM-A dimerization forms the nucleus of a complex containing Afadin and PDZ-GEF2, which in turn activates Rap1A to stabilize β1 integrin levels and promote cell migration.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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; Vetrano et al., 2008). In particular, there was significantly increased colonic mucosal injury and inflammation in JAM-A–deficient animals, compared with wild-type controls. These in vivo data are consistent with our in vitro observations implicating JAM-A as an important regulator of epithelial wound healing.
This study highlights a signaling pathway downstream of JAM-A dimerization that regulates cell migration. We recently reported that in epithelial cells, JAM-A dimerization and the JAM-A PDZ binding motif are necessary for stabilization of β1 integrin and regulation of cell migration (Severson et al., 2008). These findings are consistent with those observed in endothelial cells where decreased cell migration in JAM-A–deficient endothelial cells was restored after transfection of cells with full-length JAM-A but not expression of JAM-A lacking the PDZ binding motif (Bazzoni et al., 2005). Furthermore, Naik et al. (2003b) reported that overexpression of JAM-A increased cell migration in endothelial cells through 
vβ3 integrin and activation of mitogen-activated protein kinase. Collectively, these observations indicate that JAM-A regulates cell migration through maintenance of cellular β1 integrin protein levels.
The mechanism(s) by which JAM-A mediates cellular functions are poorly understood. We hypothesized previously that JAM-A regulation of epithelial cell migration was dependent on its cis-dimerization, which brings at least two JAM-A PDZ binding motifs into close apposition. It was proposed that such closely apposed PDZ binding motifs would allow for interactions between different scaffolding proteins. The interactions of such scaffolding proteins might then result in signaling that regulates β1 integrin protein levels (Severson et al., 2008) and cell migration.
To further explore the above-mentioned hypothesis, we tested whether JAM-A associates with candidate PDZ domain-containing scaffolding/signaling molecules in polarized intestinal epithelial cells by using standard biochemical and immunolocalization techniques. We verified a functional linkage of these scaffolding/signaling molecules to JAM-A by comparing the effects of decreased expression of these associated proteins with those observed after decreased JAM-A expression. Using this approach, we observed that decreased expression of JAM-A, Afadin, PDZ-GEF2, or Rap1A, but not ZO-1, PDZ-GEF1, or Rap1B, resulted in reduced levels of β1 integrin protein as well as decreased rates of cell migration. Additionally, decreased expression JAM-A, Afadin, and PDZ-GEF2 decreased levels of active Rap1 in a similar manner. The identical biochemical and functional effects observed after decreased expression of these proteins suggest that they are either in the same signaling pathway or separately activate the same effectors. However, because JAM-A, Afadin, and PDZ-GEF2 form part of a complex that is dependent on JAM-A expression, it is likely that these proteins participate in a single signaling pathway.
Of the proteins discussed, only Afadin has been reported to have an effect on cell migration. Lorger and Moelling (2006) reported that down-regulation of Afadin isoform 3 in MCF10A cells led to an increase in cell migration; however, this effect was due to down-regulation of Afadin isoforms 3 and dependant upon interactions with actin (Lorger and Moelling, 2006). We report decreased migration after down-regulation of Afadin in SK-CO15 cells. This difference may be due to variations in the Afadin isoforms expressed, their ability to bind to actin, or the potential absence of an effect of Afadin on β1 integrin in MCF10A cells. Additionally, the PDZ binding motif of Afadin has been reported to bind to Rap1GAP, which inactivates Rap1 (Su et al., 2003). It is possible that JAM-A and Rap1GAP compete for binding of Afadin, because they interact with the same protein domain, and such competitive binding could produce cell-type–specific responses. However, these possibilities await elucidation in further studies.
Although decreased expression of PDZ-GEF1 had no effect on β1 integrin levels or cell migration, there was a decrease in total Rap1 activity. Furthermore, simultaneous knockdown of PDZ-GEF1 and PDZ-GEF2 resulted in larger decreases in Rap1 activity than that observed with knockdown of either GEF alone. These observations would be explained if PDZ-GEF1 specifically regulates Rap1B; however, further studies are necessary to elucidate the functional consequences of decreased Rap1B activity.
Previously, our laboratory reported a correlation between Rap1B expression and β1 integrin levels (Mandell et al., 2005). In that study, we observed that siRNA-mediated down-regulation of Rap1B decreased β1 integrin levels. However, those experiments were performed using "smart-pools" of siRNA containing four separate siRNA oligonucleotides. Given that Rap1A and Rap1B are 95% conserved, we determined that the mixture of Rap1B siRNA targets previously used also resulted in partial down-regulation of Rap1A expression and thus decreased β1 integrin protein levels. In the current study, we performed real-time RT-PCR to confirm that the siRNA targets used do not cross-react between Rap1A and Rap1B. Taken together, the above-mentioned results suggest that active Rap1A, not activeRap1B, regulates β1 integrin protein levels and the rate of cell migration.
Together, the data from this and previous studies support the hypothesis that a complex of dimerized JAM-A, Afadin, and PDZ-GEF2 regulate β1 integrin at the protein level by controlling Rap1A activation. Such fine-tuning of β1 integrin levels could serve as a regulator of cell migration under a variety of circumstances. As highlighted in Figure 8, our observations support a model of JAM-A–mediated regulation of cell migration through the formation of a PDZ-dependent signaling complex. In this model, JAM-A dimerization leads to the close apposition PDZ-GEF2 and Afadin. Rap1A brought into close proximity to PDZ-GEF2 by Afadin is maintained in an active state to promote cell migration by stabilizing β1 integrin levels through, as of yet, unclear mechanisms. The protein interactions depicted in Figure 8 may not be direct but may occur via additional as of yet unidentified scaffolding proteins that may associate with the JAM-A protein complex.
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; Glading et al., 2007). In a model of junction reassembly after transient calcium depletion, JAM-A mAbs, such as J10.4, that block dimerization have been shown to slow the formation of tight and adherens junctions (Liu et al., 2000). Similarly, treatment with cAMP analogues that activate Rap1 increase the rate of junction resealing (Kooistra et al., 2005). Thus, the effect of inhibiting JAM-A dimerization of the rate of junctional formation is likely due to inhibition of Rap1 activation through by mechanism(s) similar to that outlined in this current report.
It is important to note that the murine crystal structure of JAM-A predicts the formation of dimers on the same cell (cis) that interact between cells (trans) (Kostrewa et al., 2001). Indeed, the signaling events outlined in Figure 8 would be enhanced through trans interactions of JAM-A cis-dimers between cells. Trans interactions as well as cis interactions would result in each JAM-A dimer having two trans binding sites, allowing for localized high density oligomerization of JAM-A between cells. Such localized high concentrations of JAM-A between cell–cell contacts would result in large protein complexes serving to amplify JAM-A–mediated signals. The nature of such predicted trans interactions between cis-dimers of JAM-A remain to be defined through further studies.
In summary, the findings in this report support a model of JAM-A dimerization-mediated signaling through interactions with Afadin and PDZ-GEF2 that result in activation of Rap1A, to stabilize β1 integrin levels and regulate cell migration. Additional studies are necessary to determine the mechanism by which activated Rap1A stabilizes β1 integrin levels in epithelial cells. We hypothesize that other reported JAM-A functions may use similar mechanisms for intracellular signaling; however, there are likely to be cell-type–specific signaling intermediates and effector proteins. Further studies are needed to determine whether the general mechanism described in this report is applicable to JAM-A–mediated regulation of other cellular processes.
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ACKNOWLEDGMENTS |
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Footnotes |
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Address correspondence to: Charles A. Parkos (cparkos{at}emory.edu)
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REFERENCES |
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