1Department of Anatomy, Department of Biochemistry and Biophysics, and Cardiovascular Research Institute,University of California San Francisco School of Medicine, San Francisco, California 94143-2140; 2Department of Cell Biology, University of Arizona, Tucson, Arizona 85724-5044; and 3Renal-Electrolyte Division, University of Pittsburgh, Pittsburgh, Pennsylvania 15261
Submitted 3 September 2003 ; accepted in final form 19 October 2003
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ABSTRACT |
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c-met protooncogene; transepithelial resistance; Madin-Darby canine kidney cell
The morphogenetic effects of HGF suggested that HGF may affect tight junctions (TJs). Several previous studies analyzed the effects of HGF on the assembly of TJs during the formation of a polarized epithelial monolayer (23, 55). The results of these studies indicated that HGF could inhibit assembly of newly forming epithelial cell-cell junctions. Treatment of polarized endothelial or epithelial monolayers with HGF had varying effects, either increasing or decreasing transepithelial resistance (TER) (26, 27, 34, 53). However, the time frame or conditions of HGF treatment in these studies were not reported to induce morphogenetic cell rearrangements.
The effect of HGF on TJ function during morphogenetic cell rearrangements is not well understood. Initial models of HGF-induced morphogenesis proposed that stimulation of cell dissociation is important for cell rearrangements. However, we showed previously (4, 56) that during HGF-induced tubulogenesis or formation of pseudostratified layers from polarized monolayers cell-cell contacts are maintained and the TJ marker ZO-1 remains localized at sites of contact between cells that are rearranging. In light of these results we hypothesize that TJ function may be maintained during HGF-induced cell rearrangements. A two-dimensional model system in which polarized Madin-Darby canine kidney (MDCK) cell monolayers are plated on permeable filter supports and stimulated by HGF to form pseudostratified layers (4, 91) allows functional aspects of TJs to be studied during morphogenetic cell movements.
In this study we analyzed the effect of HGF on TJ function during stimulation of morphogenetic cell rearrangements. We report that HGF induced cells of a polarized MDCK cell monolayer to crawl over each other to form a pseudostratified layer. During the transition from a monolayer to a pseudostratified layer HGF caused a transient increase in MDCK cell TER and preserved a barrier to paracellular diffusion of solutes. In addition, we show that the effects of HGF on TER were not inducible by direct activation of c-met and may require interaction of HGF with low-affinity binding sites. Our analysis of TJ integrity during pseudostratified layer formation reveals that maintenance of functionally intact TJs is an important component of HGF-stimulated morphogenesis that is likely to be critical for tissue morphogenesis and repair.
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MATERIALS AND METHODS |
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Cells and culture conditions. MDCK type II cells, either nontransfected or expressing the wild-type rabbit pIgR (45), were maintained in MEM containing Earle's balanced salt solution (MEM-EBSS; Cellgro, Mediatech, Washington, DC) supplemented with 5% FBS (Hyclone, Logan, UT), 100 U/ml penicillin, and 100 mg/ml streptomycin in 5% CO2-95% air. MDCK type II cells transfected with pSV2-neo and either a -actin promoter expression vector (pBAT) alone (46) or a pBAT vector containing a trk/met hybrid receptor cDNA (WTH2) (86) were kindly provided by M. Weidner and W. Birchmeier (Max Delbrück Center for Molecular Medicine, Berlin, Germany). These cells were maintained as described above except that 700 µg/ml G418 was included in the medium.
Assay for TER. Cells were seeded at confluent density onto Transwells (catalog no. 3401, polycarbonate membrane, 12-mm diameter, 0.4-µm pore size; Costar, Cambridge, MA) and grown in MEM-EBSS-5% FBS for 4-6 days, with daily medium changes, before use. Confluent, polarized monolayers were treated basally with HGF, MAb DO24, or NGF that was diluted into MEM-EBSS-5% FBS. For antibody inhibition of HGF, rabbit polyclonal anti-HGF antibody was diluted to a final concentration of 27 µg/ml into MEM-EBSS-5% FBS containing 100 ng/ml HGF and rotated for 2 h at 4°C before use on cell cultures.
To determine TER, cultures were washed one time quickly and then equilibrated for 10 min with MEM containing Hanks' balanced salts, 0.6% BSA, and 20 mM HEPES, pH 7.3 (MEM-BSA) at 37°C. We used a Millicell-ERS instrument (Millipore Continental Water Systems, Bedford, MA) to measure TER, being careful to keep distances from the bottom of the well and between the electrodes standardized. Resistances were calculated after subtracting background values obtained from blank Transwells that had been cultured in parallel.
Inulin diffusion measurements. Apical to basolateral 14C-labeled inulin leakage was measured across MDCK cell cultures grown on 12-mm Costar Transwells. MDCK cell cultures were plated at confluent density and grown 4-5 days in MEM-5% FBS followed by 24 h in MEM-5% FBS alone or MEM-5% FBS + 100 ng/ml rhHGF. To measure inulin diffusion, cultures were washed once with MEM-BSA at 37°C and then 0.5 ml of MEM-BSA containing 1.25 x 105 cpm of [14C]inulin was placed in the apical compartment and 1 ml of MEM-BSA in the basal well. These volumes were chosen because they result in matching fluid levels across the filter. Cultures were maintained at 37°C, and aliquots were collected, 20 µl from the apical side and 40 µl from the basal side at 1, 2, 4, and 8 h after addition of [14C]inulin. Controls included both blank filters and MDCK cultures that were grown as above in MEM-5% FBS and transferred 12 h before the diffusion assay to MEM suspension medium (S-MEM, GIBCO-BRL, Gaithersburg, MD) containing 5% dialyzed FBS and 2 µM CaCl2 ("low-Ca2+" cultures). The inulin diffusion assay for low-Ca2+ cultures was carried out in MEM-BSA containing 2 µM CaCl2. The aliquots were counted in a liquid scintillation counter (Beckman Instruments, Irvine, CA).
Assay for cell scattering. Cells were trypsinized from confluent 10-cm tissue culture plates, resuspended in MEM-EBSS-5% FBS, and counted in a hemocytometer. Two milliliters of medium containing cells at a density of 5 x 104 cells/ml was replated into each 35-mm well. Cells were cultured for 8-9 h in a 5% CO2-95% air incubator to allow cells to attach and form small islands containing 5-20 cells each. Cultures were then treated for 24-28 h with HGF, MAb DO24, or NGF and photographed with a Nikon camera attached to a Zeiss inverted microscope outfitted with phase and Hoffman Modulation Contrast optics.
Morphogenesis assay. Cells were cultured on Transwells as described for the TER analysis. For coculture experiments, MDCK cells expressing pIgR were mixed with nontransfected MDCK cells before plating onto Transwells, so that 10% of the cells plated contained pIgR. Polarized monolayers were treated for 20-24 h with 100 ng/ml HGF. Alternatively, polarized monolayers of nontransfected MDCK cells or MDCK cells transfected with pIgR, pSV2-neo/pBAT vector, or trk/met chimeric receptor were treated as above with HGF or NGF at concentrations between 2.5 and 2,500 ng/ml. All samples were processed for immunofluorescence or electron microscopy as described in Immunofluorescence and confocal microscopy and Electron microscopy.
Immunofluorescence and confocal microscopy. Transwell filter cultures were rinsed at room temperature with PBS, pH 7.4, containing 1 mM CaCl2 and 0.5 mM MgCl2 (PBS+), fixed for 30 min with 4% paraformaldehyde in PBS+, permeabilized for 30 min with 0.025% saponin in PBS+, rinsed with PBS+, and quenched for 10 min with 75 mM NH4Cl-20 mM glycine in PBS+, pH 8.0. Nonspecific binding sites were blocked by rocking for 10 min in PBS+-0.025% saponin-0.7% fish skin gelatin (block buffer) followed by 10 min in block buffer with 0.1 mg/ml boiled RNase A. Filters were incubated with primary antibody diluted in block buffer, either for 60 min in a humidified chamber at 37°C or overnight at 4°C. Primary antibody concentrations were as follows: rr1 MAb supernatant, 3:1; R40.76 anti ZO-1 ascites, 1:150; R40.76 anti ZO-1 MAb supernatant, 3:1; guinea pig anti-secretory component, 1:130. After extensive washing with PBS+-saponin and blocking buffer, cells were incubated for 30 min at 37°C in a humidified chamber in a block buffer solution containing fluorophore-conjugated secondary antibodies, diluted 1:100, and ppI, diluted 1:1,000 from a 3-4 mg/ml stock. Samples were washed extensively, postfixed with 4% paraformaldehyde in 0.1 M cacodylate buffer, pH 7.4, and mounted in Vectashield (Vector Labs, Burlingame, CA).
Confocal images were collected with a krypton-argon laser with K1 and K2 filter sets coupled to a Bio-Rad MRC600 confocal head and an Optiphot II Nikon microscope with a Plan Apo 60x 1.4 NA objective. Transwell filter samples were imaged either in the X-Y plane or in the X-Z plane with a motor step size of 0.5 or 1 µm, Kalman filtering with 5 frames/image, and diaphragm set at 1/3 open. The data were analyzed with Comos software. Images were converted to TIFF format, and composites of images were prepared with Adobe Photoshop (Adobe, Mountain View, CA) on a Macintosh computer (Apple Computer, Cupertino, CA).
Electron microscopy. For ultrastructural analysis, MDCK cells on Transwell filters were fixed 60 min on ice with 1.3% glutaraldehyde, 1 mM CaCl2, 1 mM MgCl2, 0.05% ruthenium red, and 67 mM sodium cacodylate, pH 7.4, on the apical side of the Transwell and 67 mM cacodylate, pH 7.4, on the opposing side. These samples were then postfixedfor 3 h at room temperature in 1.7% OsO4, 0.05% ruthenium red, and 67 mM sodium cacodylate, pH 7.4, counterstained overnight with 0.5% uranyl acetate, dehydrated in ethanol, and embedded in epon. Thin sections were cut in the X-Z plane of the Transwell with a diamond knife (Diatome, Fort Washington, PA) and were observed at 80 kV in a Zeiss EM-10 electron microscope.
Statistical analysis. Data are expressed as means ± SE. Statistical analysis was performed by Student's t-test, one-way ANOVA, and Tukey honestly significant difference multiple-comparison test. P < 0.05 was considered statistically significant.
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RESULTS |
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TJs are important cell-cell junctions that encircle individual epithelial cells and function to maintain segregated apical and basolateral subdomains and provide paracellular permeability barriers between epithelial cells in vivo (39). We hypothesized that during morphogenetic cell movements these cell-cell junctions are altered. Therefore, we examined the effect of HGF on TJs during formation of pseudostratified layers. The localization of ZO-1, a cytoplasmic plaque protein of TJs, was determined with confocal microscopy. We examined sections through control and HGF-treated Transwell filter-grown MDCK cell cultures in both X-Z and X-Y planes. In X-Z cross sections through untreated MDCK monolayers ZO-1 was localized at the apical-most aspect of lateral cell-cell borders (Fig. 2A, arrow). After HGF-induced morphogenesis, ZO-1 appeared in X-Z confocal images as punctate spots at sites of cell-cell contact at several levels through the pseudostratified layer (Fig. 2B). In contrast to untreated monolayers, HGF-treated cultures often had multiple spots of ZO-1 along an individual cell border (Fig. 2B'', arrowhead). To determine whether HGF affected the formation of TJ rings that normally surround epithelial cells, we collected X-Y confocal sections through basal, central, and apical planes of untreated and HGF-treated MDCK Transwell cultures. ZO-1 staining appeared within a discrete apical plane in untreated MDCK cell monolayers, where it formed a bright ring that outlined the plasma membrane of each cell-cell border (Fig. 2, C, E, and G). After HGF-induced formation of pseudostratified layers, X-Y confocal images showed that ZO-1 was found in all planes of the culturebasal, central, and apical (Fig. 2, D, F, and H). ZO-1 staining in individual X-Y planes of HGF-treated pseudostratified layers appeared discontinuous. However, by examining projections of confocal images, obtained by summing all X-Y confocal sections through a sample, we found that there were complete rings of ZO-1 around each cell in both monolayers and pseudostratified layers (Fig. 2, I and J). This indicates that, although HGF induced changes in cell shape and the localization of ZO-1, staining of ZO-1 in pseudostratified layers depicts morphologically intact TJ belts. These results raised the possibility that TJs remain functionally intact during morphogenesis despite their irregular organization.
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To determine whether the morphologically complete rings of ZO-1 represent functionally intact TJs we tested the integrity of TJs during HGF-induced morphogenesis. Ruthenium red is a heavy metal conjugate that is too large to cross a functionally intact TJ and can be visualized with electron microscopy (35, 36, 83). Ruthenium red was added to the apical side of either untreated or HGF-treated Transwell filter cultures of MDCK cells during fixation and processing for electron microscopy. X-Z sections were prepared of MDCK cell monolayers and pseudostratified layers, and images were collected to determine whether ruthenium red crossed the cell layers. In electron micrographs, shown in Fig. 3, we observed that TJs between cells in HGF-treated cultures were no longer localized solely at the apical aspect of the plasma membrane but were also found at different levels along the lateral membrane (Fig. 3B, arrows), in agreement with the detection of ZO-1 in different planes of section mentioned above. However, ruthenium red remained at the apical side of both untreated and HGF-treated MDCK cell layers. In addition, ruthenium red added basally never reached the apical cell surface, although it appeared to encircle portions of cells that angled through the pseudostratified layer underneath the apical-most cells and were sectioned below the level of the TJs (not shown). Together, our results indicate that TJs are morphologically and functionally intact, maintaining paracellular permeability barriers during HGF-induced rearrangement of a monolayer into a pseudostratified layer.
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To quantitatively measure the functional integrity of TJs during HGF-induced cell rearrangements we measured TER in response to treatment with HGF. In low-resistance MDCK strain II cells, TER is comprised of transcellular and paracellular resistors in parallel for which the transcellular resistance is two orders of magnitude greater than the paracellular resistance. Therefore, for these cells TER is a direct indicator for tightness of the paracellular seal and an instantaneous measure of paracellular permeability (21, 61). MDCK cells were plated and maintained at confluent density on Transwell filters to form electrically tight, polarized monolayers and then cultured for an additional 20 h in the presence or absence of 100 ng/ml rhHGF. TER measurements are shown in Fig. 4A. In the absence of HGF, TJs between MDCK cells in monolayers maintained a TER of 100
-cm2. Surprisingly, the TER of MDCK cell cultures treated with HGF was increased more than twofold compared with untreated cultures. Similar results were obtained with a clone of MDCK cells transfected with pIgR, demonstrating that the effect of HGF on TER is not clone specific. In addition, HGF treatment from the basal side of the filter alone was sufficient to induce a maximal increase in TER (data not shown; Ref. 13). Therefore, all further experiments were carried out by treating monolayers with HGF on the basolateral side only. To directly test that the effect on TER was due specifically to HGF we treated the basolateral side of filter-grown MDCK cell monolayers with medium that had been preincubated with both 100 ng/ml HGF and anti-HGF polyclonal antibodies. Figure 4A shows that antibody pretreatment almost completely blocked the HGF-induced increase in TER, confirming that the effect on TER is due to HGF. These results indicate that HGF activation of a basolaterally localized receptor mechanism causes an increase in MDCK cell monolayer TER.
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To determine the time course of the effect of HGF on TER we cultured polarized Transwell filter-grown monolayers of MDCK cells for various time periods in the presence and absence of continuous HGF treatment. We found (Fig. 4B) that HGF caused a transient increase in TER, peaking around 24 h and then declining to baseline by 48 h. A similar time course of effect was found with the pIgR-expressing clone of MDCK, which had a baseline TER of 96 ± 3 -cm2 and an HGF-induced peak TER at 24 h of 251 ± 17
-cm2 (not shown), demonstrating that this effect is not clone specific. Thus the HGF-stimulated rise in TER correlates in time with the morphological transition of a monolayer into a pseudostratified layer and then declines to baseline.
To analyze the effect of HGF on the dynamic function of TJs during HGF-stimulated cell rearrangement we measured apical to basolateral [14C]inulin leakage across Transwell filter-grown MDCK cell cultures during stimulation of pseudostratified layer formation. In contrast to TER, which is an instantaneous measure of TJ functional integrity, [14C]inulin diffusion is measured over a period of several hours and therefore may be a more sensitive measure of apical to basolateral leakage. Tight, polarized MDCK cell monolayers were treated for 24 h in the absence or presence of HGF, and treatment was continued as apical to basolateral diffusion was measured by including [14C]inulin in the apical medium and collecting aliquots of both apical and basolateral medium at various time points (Fig. 5). We found that there was no significant increase in the amount of apical to basolateral [14C]inulin diffusion within the first 2 h of measurement. A small but statistically insignificant increase in the amount of basolateral [14C]inulin was detectable 4-8 h after apical application of [14C]inulin. In contrast, in samples treated with low-Ca2+ medium complete equilibration of [14C]inulin is detected, demonstrating that TJ function is completely lost. This suggests that the overall integrity of TJs is maintained in HGF-treated cultures during the 8-h time period in which [14C]inulin diffusion was measured.
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Our initial studies suggested that the concentration of HGF required to cause an increase in TER was much higher than that required to induce scattering (not shown). To analyze the amount of HGF required to stimulate these different activities we tested different concentrations of rhHGF (0-250 ng/ml) for effects on TER vs. scattering. In Fig. 6, we show that MDCK cells plated and allowed to form small colonies (Fig. 6A) elicited a full scattering response when treated with either 2.5 (Fig. 6B) or 100 (Fig. 6C) ng/ml HGF. In contrast, the dose-response of TER to HGF (Fig. 6D) demonstrates that 2.5 ng/ml HGF had no effect on TER and that peak effects were induced by HGF concentrations of 100 ng/ml.
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The c-met protooncogene is identified as the HGF receptor that transduces all known functions of HGF (86). This receptor is basolaterally localized in polarized MDCK cells (13), suggesting it as a likely candidate to transduce the HGF signal, resulting in an increase in TER. To test whether direct activation of c-met causes an increase in TER similar to that seen with HGF we took the following two approaches: 1) stimulation of MDCK cells with an activating antibody of c-met, DO24 (57-59), and 2) NGF stimulation of transfected MDCK cells expressing a trk/met chimeric receptor containing the ligand binding domain of the NGF receptor and the transmembrane and tyrosine kinase domains of c-met (86).
The scattering activity of 0, 2, and 10 nM DO24 is shown in Fig. 7, A-C. DO24 induced a complete MDCK cell scattering response at a concentration of 2 nM (compare Fig. 7A with Fig. 7, B and C). The effect of DO24 on TER was tested at 2, 10, 100, and 200 nM, concentrations that included and far exceeded the doses necessary to induce scattering. Results in Fig. 7D show that DO24 induced a small increase in TER that was similar at all concentrations tested, demonstrating that direct antibody activation of c-met produced only a partial effect on TER.
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Weidner et al. (86) showed previously that all known biological effects of HGF, including stimulation of cell scattering, invasiveness, morphogenesis, and proliferation, are inducible by NGF treatment of MDCK cells that are expressing a trk/met chimeric receptor. This demonstrated that direct activation of c-met transduces these cellular responses. The effects of both HGF and NGF could be tested with trk/met-transfected MDCK cells because they retain expression of the endogenous c-met receptor. Therefore, we compared the effects of HGF and NGF on the TER of control (pSV2-neo/pBAT vector) or trk/met-transfected MDCK cells to determine whether the same mechanism, direct activation of c-met, also transduces this response. We first assayed scattering as a functional measure of HGF and NGF activity in our culture system. Similar to the results shown above for nontransfected or pIgR-transfected MDCK cells, treatment of control transfected (Fig. 8C) or trk/met (Fig. 8D) clones with 2.5 ng/ml HGF produced a complete scattering response compared with untreated cultures (Fig. 8, A and B). As expected, treatment with either 2.5 or 250 ng/ml NGF had no effect on scattering of control clones (compare Fig. 8, A, E, and G). However, 2.5 ng/ml NGF induced a complete scattering response with trk/met-expressing MDCK cells (compare Figs. 8, B, F, and H) similar to the effect of 2.5 ng/ml HGF (compare Fig. 8, D and F). This confirmed that stimulation of the trk/met chimera with NGF fully activated c-met transduction of the scattering response.
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We then compared the effects of NGF and HGF on TER. Figure 8I shows that HGF caused an increase in the TER of monolayer cultures of both control-transfected and trk/met MDCK cells, with peak effects at 100 ng/ml. The peak effects of HGF on TER of control-transfected MDCK (pSV2-neo/pBAT vector) were similar to those in nontransfected and pIgR-transfected MDCK clones, resulting in a two- to three- fold increase in TER. trk/met-Transfected MDCK cells responded to HGF with a three- to fourfold increase in TER. In contrast, NGF, at concentrations between 2 and 25,000 ng/ml, did not have any effect on TER of either control or trk/met-transfected MDCK cell cultures (Fig. 8J). In addition, the ability of HGF (2.5, 25, 50, 100, 250, and 500 ng/ml) or NGF (2.5, 25, 250, 500, and 2,500 ng/ml) to induce trk/met MDCK cell pseudostratified layer morphogenesis was tested. Representative X-Z confocal sections are shown in Fig. 9. In untreated trk/met MDCK cell monolayers the nuclei were basally located and E-cadherin was basolateral. Cells remained in polarized monolayers after treatment with HGF at 2.5 ng/ml. At 25 and 50 ng/ml HGF, nuclei appeared more uneven but cells were still in monolayers (not shown). Pseudostratified layers were formed at 100, 250, and 500 ng/ml HGF. In contrast, NGF did not induce pseudostratified layer morphogenesis at any concentration tested. Therefore, our data provide the surprising evidence that direct activation of the trk/met chimeric receptor with NGF induces scattering but is not sufficient to cause pseudostratified layer morphogenesis or an increase in TER.
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DISCUSSION |
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We had initially hypothesized that HGF would cause a decrease in MDCK cell TER during morphogenesis. This hypothesis was based on the previously reported ability of HGF to induce scattering of MDCK cells (75, 76), disrupt TJ function (53), or inhibit TJ assembly after calcium switch (23, 26). Instead, after testing four different subclones of MDCK type II cells with recombinant human HGF concentrations ranging from 2.5 to 500 ng/ml, we found that HGF increased TER (Figs. 4, 6, and 8). Our results were not due to alterations in transcellular resistance, because HGF does not affect the function of ion channels in the transcellular pathway (53). The increase in TER was transient, peaking at 24 h and then declining to baseline (Fig. 4). Treatment of Transwell filter-grown cultures of MDCK cells with HGF for up to 72 h did not decrease TER below baseline. The timing of the HGF-induced increase in TER paralleled that of HGF stimulation of morphogenesis, demonstrating that HGF induced cell rearrangements without loss of TJ functional integrity during the transition of a polarized MDCK cell monolayer into a pseudostratified layer.
The increase in TER that we observed in response to HGF contrasts with studies by Nusrat et al. (53) in which recombinant human HGF caused a decrease in TER of monolayers of the human intestinal cell line T84 and mouse HGF caused a similar decrease with MDCK cells. In our hands, culturing MDCK cells similarly to Nusrat et al. on cross-linked or non-cross-linked collagen-coated filters lowered the baseline TER but fold increases in TER in response to either 100 or 500 ng/ml rhHGF were similar to those observed with uncoated Transwell filters. Nusrat et al. (53) reported that cell polarity was not altered by HGF treatment, suggesting that cell rearrangements were not induced. Therefore, the difference between our TER results and those of Nusrat et al. may depend on whether cell rearrangements were induced by HGF.
HGF may have differential effects depending on the initial state of cell polarity and cell-cell and cell-substrate adhesion. Variable effects of HGF on TER of endothelial and epithelial monolayers have been reported by others, showing a decrease (26, 27), an increase (34), or no effect (34) on TER. Culture environment, cell type, initial cell polarity, cell-substrate and cell-cell adhesion, dose of HGF, and treatment times varied considerably among these studies and may have contributed to the differences in effect. For example, it was reported that monolayers of lung endothelial cells but not alveolar cells increased TER in response to HGF (34), and several studies showed that altering cell-substrate interactions changed responsiveness to HGF (62, 68). These results suggest that the effects of HGF are highly context dependent.
The context-dependent effects of HGF discussed above may be important for processes such as wound healing. In wound healing the context of cells at a wound edge is different from that of cells that are more distant from a wound, and it is critical to induce rearrangements of cells that are adjacent to a wound edge without compromising the overall paracellular tightness of the epithelium. During wound repair in several tissues such as liver, kidney, lung, and gastric mucosa, HGF production and activation are increased and HGF contributes to acceleration of wound repair (40, 53, 66, 80, 88). One mechanism of wound healing, called restitution (33, 44), is used by epithelial cell sheets in adult organisms and involves the stimulation of filopodia and lamella extension that is similar to cell crawling induced by HGF (17). Fenteany et al. (16) demonstrated, using an MDCK epithelial cell monolayer model system for this type of wound repair, that cells both around the wound margin and several rows distant migrate in to close a wound. Our results suggest that HGF's ability to induce motility of cells within a tight monolayer while modulating but not compromising the overall TJ function of the epithelial cell layer may be an important component of wound healing.
In addition to analyzing the effects of HGF on TER, we also tested the effect of HGF on paracellular transport of the nonionic molecule inulin. Previous studies have analyzed the effects of HGF on intestinal or renal epithelial permeability to inulin and have obtained results similar to ours showing a slight increase in inulin flux compared with untreated monolayers. However, the authors of these previous studies have drawn contrasting conclusions, suggesting either that marginal increases in inulin permeability induced by HGF indicate that monolayer integrity is preserved or that small increases in inulin flux indicate a loss of cell-cell junctional integrity (5, 53). In our study, in addition to testing the inulin permeability of control and HGF-treated MDCK cell monolayers, we also analyzed inulin permeability across monolayers that lack TJs by testing confluent MDCK cell monolayers in which cell-cell adhesion was removed by treatment with low-Ca2+ medium. Our results confirm that the effects of HGF on inulin permeability are minimal compared with monolayers in which cell-cell junctional integrity is lost. This effect is not due to increased transcellular transport, because previous studies showed that HGF decreases transcytosis (4). The small increase in inulin diffusion in HGF-treated cultures may reflect dynamic changes in TJs during cell rearrangements. For instance, transient openings and closings of individual TJs may initially trap and subsequently release inulin from between the multiple TJs that develop along a single lateral cell-cell border during morphogenesis.
In the present study, we used multiple approaches to analyze coordinately both morphological and functional effects of HGF on TJs from polarized epithelial monolayers as cells were stimulated to rearrange. Three-dimensional (3D) analyses of morphological alterations in both X-Y and X-Z planes provided a clearer understanding of how HGF alters TJs during cell rearrangements. For example, if we had analyzed only individual confocal sections in single planes we might have concluded that ZO-1 staining is disorganized and discontinuous after HGF treatment. In contrast, immunofluorescence microscopy without sectioning can give the impression that HGF did not affect ZO-1, similar to a projection of summed X-Y confocal sections. The 3D morphological analyses demonstrated that, in response to HGF, ZO-1 is rearranged but tight junctional rings encircling each cell are morphologically complete (Fig. 2). Our morphological and quantitative measures of TJ functional integrity together provide strong evidence that TJ functional integrity is maintained during HGF-induced morphogenesis.
Our results demonstrate that HGF induces an increase in TER and crawling of cells from a polarized MDCK monolayer to form a pseudostratified layer. However, the downstream signaling pathway that mediates these HGF-induced effects is less clear. Our results show that MAb DO24 activation of endogenous c-met or NGF activation of a trk/met chimera stimulated all of the previously identified effects of HGF on MDCK cells but did not affect TER or pseudostratified layer formation. Prat et al. (58) showed previously that the MAbs DN30 and DO24 both bind c-met and activate receptor dimerization and phosphorylation but the biological responses to these two ligands are very different. These results suggested that specific interactions of ligands with the c-met receptor can alter downstream responses. Our results indicate that direct activation of the c-met signaling pathway by non-HGF ligands that are known to induce motility, invasion, tubulogenesis, proliferation, and cell survival is not sufficient to mediate an increase in TER or pseudostratified layer formation.
We observed that the concentration of HGF required to stimulate an increase in TER is much higher than that required to stimulate scattering. Concentrations of HGF that are sufficient to stimulate an increase in MDCK cell TER are at saturating levels for the high-affinity HGF receptor c-met, which has a Kd equal to 20 pM (79). However, MDCK cells also contain lower-affinity, higher-capacity HGF binding sites (79, 89). These sites have an affinity that is 10-fold lower than the high-affinity receptor. The concentrations of HGF that induce an increase in TER are close to the KD of the low-affinity MDCK cell surface HGF binding sites. During tissue repair and disease the concentration of HGF increases significantly over baseline in both injured tissue and plasma (40, 66, 88). On the basis of our results we surmise that local concentrations of HGF surrounding cells in injured tissue are increased and interactions with low-affinity sites may affect HGF activity to stimulate cell movement and alterations in TER that contribute to tissue repair.
Previous studies determined that in some assay systems low-affinity binding interactions of HGF with the cell surface could be inhibited or disrupted by the presence of an excess of heparin (>1 µg/ml), suggesting that these sites are composed of glycosaminoglycans (48, 52, 89). Subsequently, specific HGF-binding heparan sulfate structures and sulfoglycolipids were identified. These sites are endogenously expressed at cell surfaces of various tissues, and their interactions with HGF are heparin sensitive (3, 30, 37). c-met was also found to contain potential glycosaminoglycan binding domains (10), suggesting that trimeric complexes may form between cell surface heparan sulfate proteoglycans or sulfoglycolipids, HGF, and c-met.
Several studies have shown that glycosaminoglycans affect HGF function. For example, the presence of soluble or substrate-bound heparin or other heparin-like sulfated oligosaccharides has been found to increase the mitogenic potential of HGF, induction of cell motility by HGF, oligomerization of HGF, and autophosphorylation of c-met (1, 14, 29, 48, 60, 65, 90, 92). In addition, recent evidence shows that binding of syndecan-1, a heparan sulfate proteoglycan, to HGF enhances c-met-induced downstream signaling responses (15). The results of these studies suggest that low-affinity HGF binding sites are important regulators of HGF-induced signaling. We found that low concentrations (0.1 µg/ml) of heparin potentiated the effect of submaximal doses (25 or 50 ng/ml) of HGF on MDCK cell TER (unpublished data). NGF and MAb DO24 do not bind heparin and most likely do not interact with low-affinity HGF binding sites. Naka et al. (48) showed previously that 0.1 µg/ml heparin does not disrupt binding of HGF to cell surface receptors. It is possible that 0.1 µg/ml heparin in our system increased oligomerization of HGF and potentiated HGF activity without interfering with HGF interaction with endogenous glycosaminoglycans or c-met, similar to effects shown by Zioncheck et al. (92) on hepatocyte mitogenic potency. We suggest two possible HGF signaling mechanisms to explain our results. Low-affinity sites may alter responses to HGF by 1) providing an independent HGF receptor pathway or 2) acting as cofactors in signaling pathways that involve c-met. Our results, together with those of others, suggest that interactions of HGF with low-affinity binding sites may be important for mediating the TER and pseudostratified layer morphogenesis responses to HGF.
In summary, we demonstrate that TJ functional integrity is maintained during HGF-induced epithelial morphogenesis. We show that in response to HGF individual cells of a polarized epithelial monolayer were stimulated to move relative to their neighbors to form a pseudostratified layer. Individual TJs may be modified during this process, but the overall function of the epithelial layer as a barrier to paracellular transport is maintained. In addition, the mechanism of HGF-induced increases in TER and stimulation of pseudostratified layer formation is not activated by non-HGF ligands that directly activate c-met. We suggest that low-affinity HGF binding sites are involved in mediating the HGF-specific effects. The ability of polarized epithelial cells to migrate in response to HGF without loss of TJ functional integrity is consistent with a critical role of this mechanism in processes such as wound healing. During wound healing, the context of cells adjacent to a wound margin is different from that of cells that are farther from the wound edge. Our results suggest that cells in these different contexts may modulate their response to HGF and contribute to wound healing without disrupting the overall TJ functional integrity of the epithelium.
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ACKNOWLEDGMENTS |
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Present address of G. Apodaca: Renal-Electrolyte Div., University of Pittsburgh, Pittsburgh, PA 15261.
GRANTS
A. L. Pollack was supported by American Heart Association Postdoctoral Fellowship AHA 0020628Z. G. Apodaca was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grant P01-AI-53194. This work was also supported by National Institute of Allergy and Infectious Diseases Grant P01-AI-05319 awarded to K. E. Mostov.
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
2. Anderson JM, Stevenson BR, Jesaitis LA, Goodenough DA, and Mooseker MS. Characterization of ZO-1, a protein component of the tight junction from mouse liver and Madin-Darby canine kidney cells. J Cell Biol 106: 1141-1149, 1988.[Abstract]
3. Ashikari S, Habuchi H, and Kimata K. Characterization of heparan sulfate oligosaccharides that bind to hepatocyte growth factor. J Biol Chem 270: 29586-29593, 1995.
4. Balkovetz DF, Pollack AL, and Mostov KE. Hepatocyte growth factor alters the polarity of Madin-Darby canine kidney cell monolayers. J Biol Chem 272: 3471-3477, 1997.
5. Balkovetz DF and Sambandam V. Dynamics of E-cadherin and gamma-catenin complexes during dedifferentiation of polarized MDCK cells. Kidney Int 56: 910-921, 1999.[CrossRef][ISI][Medline]
6. Bladt F, Riethmacher D, Isenmann S, Aguzzi A, and Birchmeier C. Essential role for the c-met receptor in the migration of myogenic precursor cells into the limb bud. Nature 376: 768-771, 1995.[CrossRef][ISI][Medline]
7. Breitfeld PP, Casanova JE, Harris JM, Simister NE, and Mostov KE. Expression and analysis of the polymeric immunoglobulin receptor in Madin-Darby canine kidney cells using retroviral vectors. Methods Cell Biol 32: 329-337, 1989.[ISI][Medline]
8. Breitfeld PP, Harris JM, and Mostov KE. Postendocytotic sorting of the ligand for the polymeric immunoglobulin receptor in Madin-Darby canine kidney cells. J Cell Biol 109: 475-486, 1989.[Abstract]
9. Bussolino F, Di Renzo MF, Ziche M, Bocchietto E, Olivero M, Naldini L, Gaudino G, Tamagnone L, Coffer A, and Comoglio PM. Hepatocyte growth factor is a potent angiogenic factor which stimulates endothelial cell motility and growth. J Cell Biol 119: 629-641, 1992.[Abstract]
10. Cardin AD and Weintraub HJ. Molecular modeling of protein-glycosaminoglycan interactions. Arteriosclerosis 9: 21-32, 1989.[Abstract]
11. Cooper CS, Park M, Blair DG, Tainsky MA, Huebner K, Croce CM, and Vande Woude GF. Molecular cloning of a new transforming gene from a chemically transformed human cell line. Nature 311: 29-34, 1984.[ISI][Medline]
12. Cornelison DD and Wold BJ. Single-cell analysis of regulatory gene expression in quiescent and activated mouse skeletal muscle satellite cells. Dev Biol 191: 270-283, 1997.[CrossRef][ISI][Medline]
13. Crepaldi T, Pollack AL, Prat M, Zborek A, Mostov K, and Comoglio PM. Targeting of the SF/HGF receptor to the basolateral domain of polarized epithelial cells. J Cell Biol 125: 313-320, 1994.[Abstract]
14. Deakin JA and Lyon M. Differential regulation of hepatocyte growth factor/scatter factor by cell surface proteoglycans and free glycosaminoglycan chains. J Cell Sci 112: 1999-2009, 1999.
15. Derksen PW, Keehnen RM, Evers LM, van Oers MH, Spaargaren M, and Pals ST. Cell surface proteoglycan syndecan-1 mediates hepatocyte growth factor binding and promotes Met signaling in multiple myeloma. Blood 99: 1405-1410, 2002.
16. Fenteany G, Janmey PA, and Stossel TP. Signaling pathways and cell mechanics involved in wound closure by epithelial cell sheets. Curr Biol 10: 831-838, 2000.[CrossRef][ISI][Medline]
17. Geiser TK, Kazmierczak BI, Garrity-Ryan LK, Matthay MA, and Engel JN. Pseudomonas aeruginosa ExoT inhibits in vitro lung epithelial wound repair. Cell Microbiol 3: 223-236, 2001.[CrossRef][ISI][Medline]
18. Gherardi E, Gray J, Stoker M, Perryman M, and Furlong R. Purification of scatter factor, a fibroblast-derived basic protein that modulates epithelial interactions and movement. Proc Natl Acad Sci USA 86: 5844-5848, 1989.[Abstract]
19. Giordano S, Ponzetto C, Di Renzo MF, Cooper CS, and Comoglio PM. Tyrosine kinase receptor indistinguishable from the c-met protein. Nature 339: 155-156, 1989.[CrossRef][ISI][Medline]
20. Gohda E, Tsubouchi H, Nakayama H, Hirono S, Sakiyama O, Takahashi K, Miyazaki H, Hashimoto S, and Daikuhara Y. Purification and partial characterization of hepatocyte growth factor from plasma of a patient with fulminant hepatic failure. J Clin Invest 81: 414-419, 1988.[ISI][Medline]
21. Gonzalez-Mariscal L. The relationship between structure and function of tight junctions. In: Tight Junctions, edited by Cereijido M. Boca Raton, FL: CRC, 1991, p. 67-76.
22. Grant DS, Kleinman HK, Goldberg ID, Bhargava MM, Nickoloff BJ, Kinsella JL, Polverini P, and Rosen EM. Scatter factor induces blood vessel formation in vivo. Proc Natl Acad Sci USA 90: 1937-1941, 1993.[Abstract]
23. Grisendi S, Arpin M, and Crepaldi T. Effect of hepatocyte growth factor on assembly of zonula occludens-1 protein at the plasma membrane. J Cell Physiol 176: 465-471, 1998.[CrossRef][ISI][Medline]
24. Gumbiner B and Simons K. A functional assay for proteins involved in establishing an epithelial occluding barrier: identification of a uvomorulin-like polypeptide. J Cell Biol 102: 457-468, 1986.[Abstract]
25. Higuchi O and Nakamura T. Identification and change in the receptor for hepatocyte growth factor in rat liver after partial hepatectomy or induced hepatitis. Biochem Biophys Res Commun 176: 599-607, 1991.[ISI][Medline]
26. Hollande F, Blanc EM, Bali JP, Whitehead RH, Pelegrin A, Baldwin GS, and Choquet A. HGF regulates tight junctions in new nontumorigenic gastric epithelial cell line. Am J Physiol Gastrointest Liver Physiol 280: G910-G921, 2001.
27. Jiang WG, Martin TA, Matsumoto K, Nakamura T, and Mansel RE. Hepatocyte growth factor/scatter factor decreases the expression of occludin and transendothelial resistance (TER) and increases paracellular permeability in human vascular endothelial cells. J Cell Physiol 181: 319-329, 1999.[CrossRef][ISI][Medline]
28. Karp SL, Ortiz-Arduan A, Li S, and Neilson EG. Epithelial differentiation of metanephric mesenchymal cells after stimulation with hepatocyte growth factor or embryonic spinal cord. Proc Natl Acad Sci USA 91: 5286-5290, 1994.[Abstract]
29. Kato S, Ishii T, Hara H, Sugiura N, Kimata K, and Akamatsu N. Hepatocyte growth factor immobilized onto culture substrates through heparin and matrigel enhances DNA synthesis in primary rat hepatocytes. Exp Cell Res 211: 53-58, 1994.[CrossRef][ISI][Medline]
30. Kobayashi T, Honke K, Miyazaki T, Matsumoto K, Nakamura T, Ishizuka I, and Makita A. Hepatocyte growth factor specifically binds to sulfoglycolipids. J Biol Chem 269: 9817-9821, 1994.
31. Komada M and Kitamura N. The cell dissociation and motility triggered by scatter factor/hepatocyte growth factor are mediated through the cytoplasmic domain of the c-Met receptor. Oncogene 8: 2381-2390, 1993.[ISI][Medline]
32. Komada M, Miyazawa K, Ishii T, and Kitamura N. Characterization of hepatocyte-growth-factor receptors on Meth A cells. Eur J Biochem 204: 857-864, 1992.[Abstract]
33. Lacy ER and Ito S. Rapid epithelial restitution of the rat gastric mucosa after ethanol injury. Lab Invest 51: 573-583, 1984.[ISI][Medline]
34. Liu F, Schaphorst KL, Verin AD, Jacobs K, Birukova A, Day RM, Bogatcheva N, Bottaro DP, and Garcia JG. Hepatocyte growth factor enhances endothelial cell barrier function and cortical cytoskeletal rearrangement: potential role of glycogen synthase kinase-3. FASEB J 16: 950-962, 2002.
35. Luft JH. Ruthenium red and violet. I. Chemistry, purification, methods of use for electron microscopy and mechanism of action. Anat Rec 171: 347-368, 1971.[ISI][Medline]
36. Luft JH. Ruthenium red and violet. II. Fine structural localization in animal tissues. Anat Rec 171: 369-416, 1971.[ISI][Medline]
37. Lyon M, Deakin JA, Mizuno K, Nakamura T, and Gallagher JT. Interaction of hepatocyte growth factor with heparan sulfate. Elucidation of the major heparan sulfate structural determinants. J Biol Chem 269: 11216-11223, 1994.
38. Matsumoto K, Tajima H, Hamanoue M, Kohno S, Kinoshita T, and Nakamura T. Identification and characterization of "injurin," an inducer of expression of the gene for hepatocyte growth factor. Proc Natl Acad Sci USA 89: 3800-3804, 1992.[Abstract]
39. Matter K and Balda MS. Signalling to and from tight junctions. Nat Rev Mol Cell Biol 4: 225-236, 2003.[CrossRef][ISI][Medline]
40. Miyazawa K, Shimomura T, Naka D, and Kitamura N. Proteolytic activation of hepatocyte growth factor in response to tissue injury. J Biol Chem 269: 8966-8970, 1994.
41. Miyazawa K, Tsubouchi H, Naka D, Takahashi K, Okigaki M, Arakaki N, Nakayama H, Hirono S, Sakiyama O, Takahashi K, Gohda E, Daikuhara Y, and Kitamura N. Molecular cloning and sequence analysis of cDNA for human hepatocyte growth factor. Biochem Biophys Res Commun 163: 967-973, 1989.[ISI][Medline]
42. Montesano R, Matsumoto K, Nakamura T, and Orci L. Identification of a fibroblast-derived epithelial morphogen as hepatocyte growth factor. Cell 67: 901-908, 1991.[ISI][Medline]
43. Montesano R, Schaller G, and Orci L. Induction of epithelial tubular morphogenesis in vitro by fibroblast-derived soluble factors. Cell 66: 697-711, 1991.[ISI][Medline]
44. Moore R, Carlson S, and Madara JL. Rapid barrier restitution in an in vitro model of intestinal epithelial injury. Lab Invest 60: 237-244, 1989.[ISI][Medline]
45. Mostov KE and Deitcher DL. Polymeric immunoglobulin receptor expressed in MDCK cells transcytoses IgA. Cell 46: 613-621, 1986.[ISI][Medline]
46. Nagafuchi A and Takeichi M. Cell binding function of E-cadherin is regulated by the cytoplasmic domain. EMBO J 7: 3679-3684, 1988.[Abstract]
47. Naidu YM, Rosen EM, Zitnick R, Goldberg I, Park M, Naujokas M, Polverini PJ, and Nickoloff BJ. Role of scatter factor in the pathogenesis of AIDS-related Kaposi sarcoma. Proc Natl Acad Sci USA 91: 5281-5285, 1994.[Abstract]
48. Naka D, Ishii T, Shimomura T, Hishida T, and Hara H. Heparin modulates the receptor-binding and mitogenic activity of hepatocyte growth factor on hepatocytes. Exp Cell Res 209: 317-324, 1993.[CrossRef][ISI][Medline]
49. Nakamura T, Nawa K, Ichihara A, Kaise N, and Nishino T. Purification and subunit structure of hepatocyte growth factor from rat platelets. FEBS Lett 224: 311-316, 1987.[CrossRef][ISI][Medline]
50. Nakamura T, Nishizawa T, Hagiya M, Seki T, Shimonishi M, Sugimura A, Tashiro K, and Shimizu S. Molecular cloning and expression of human hepatocyte growth factor. Nature 342: 440-443, 1989.[CrossRef][ISI][Medline]
51. Naldini L, Vigna E, Narsimhan RP, Gaudino G, Zarnegar R, Michalopoulos GK, and Comoglio PM. Hepatocyte growth factor (HGF) stimulates the tyrosine kinase activity of the receptor encoded by the proto-oncogene c-MET. Oncogene 6: 501-504, 1991.[ISI][Medline]
52. Naldini L, Weidner KM, Vigna E, Gaudino G, Bardelli A, Ponzetto C, Narsimhan RP, Hartmann G, Zarnegar R, and Michalopoulos GK. Scatter factor and hepatocyte growth factor are indistinguishable ligands for the MET receptor. EMBO J 10: 2867-2878, 1991.[Abstract]
53. Nusrat A, Parkos CA, Bacarra AE, Godowski PJ, Delp-Archer C, Rosen EM, and Madara JL. Hepatocyte growth factor/scatter factor effects on epithelia. Regulation of intercellular junctions in transformed and nontransformed cell lines, basolateral polarization of c-met receptor in transformed and natural intestinal epithelia, and induction of rapid wound repair in a transformed model epithelium. J Clin Invest 93: 2056-2065, 1994.[ISI][Medline]
54. Park M, Dean M, Kaul K, Braun MJ, and Vande Woude G. Sequence of MET protooncogene cDNA has features characteristic of the tyrosine kinase family of growth-factor receptors. Proc Natl Acad Sci USA 84: 6379-6383, 1987.[Abstract]
55. Pasdar M, Li Z, Marreli M, Nguyen BT, Park M, and Wong K. Inhibition of junction assembly in cultured epithelial cells by hepatocyte growth factor/scatter factor is concomitant with increased stability and altered phosphorylation of the soluble junctional molecules. Cell Growth Differ 8: 451-462, 1997.[Abstract]
56. Pollack AL, Runyan RB, and Mostov KE. Morphogenetic mechanisms of epithelial tubulogenesis: MDCK cell polarity is transiently rearranged without loss of cell-cell contact during scatter factor/hepatocyte growth factor-induced tubulogenesis. Dev Biol 204: 64-79, 1998.[CrossRef][ISI][Medline]
57. Prat M, Crepaldi T, Gandino L, Giordano S, Longati P, and Comoglio P. C-terminal truncated forms of Met, the hepatocyte growth factor receptor. Mol Cell Biol 11: 5954-5962, 1991.[ISI][Medline]
58. Prat M, Crepaldi T, Pennacchietti S, Bussolino F, and Comoglio PM. Agonistic monoclonal antibodies against the Met receptor dissect the biological responses to HGF. J Cell Sci 111: 237-247, 1998.
59. Prat M, Narsimhan RP, Crepaldi T, Nicotra MR, Natali PG, and Comoglio PM. The receptor encoded by the human c-MET oncogene is expressed in hepatocytes, epithelial cells and solid tumors. Int J Cancer 49: 323-328, 1991.[ISI][Medline]
60. Rahimi N, Saulnier R, Nakamura T, Park M, and Elliott B. Role of hepatocyte growth factor in breast cancer: a novel mitogenic factor secreted by adipocytes. DNA Cell Biol 13: 1189-1197, 1994.[ISI][Medline]
61. Reuss L. Tight junction permeability to ions and water. In: Tight Junctions, edited by M C. Boca Raton, FL: CRC, 1991, p. 49-66.
62. Rosario M and Birchmeier W. How to make tubes: signaling by the Met receptor tyrosine kinase. Trends Cell Biol 13: 328-335, 2003.[CrossRef][ISI][Medline]
63. Rosen EM, Jaken S, Carley W, Luckett PM, Setter E, Bhargava M, and Goldberg ID. Regulation of motility in bovine brain endothelial cells. J Cell Physiol 146: 325-335, 1991.[ISI][Medline]
64. Rosen EM, Meromsky L, Setter E, Vinter DW, and Goldberg ID. Purified scatter factor stimulates epithelial and vascular endothelial cell migration. Proc Soc Exp Biol Med 195: 34-43, 1990.[Abstract]
65. Sakata H, Stahl SJ, Taylor WG, Rosenberg JM, Sakaguchi K, Wingfield PT, and Rubin JS. Heparin binding and oligomerization of hepatocyte growth factor/scatter factor isoforms. Heparan sulfate glycosaminoglycan requirement for Met binding and signaling. J Biol Chem 272: 9457-9463, 1997.
66. Schmassmann A, Stettler C, Poulsom R, Tarasova N, Hirschi C, Flogerzi B, Matsumoto K, Nakamura T, and Halter F. Roles of hepatocyte growth factor and its receptor Met during gastric ulcer healing in rats. Gastroenterology 113: 1858-1872, 1997.[ISI][Medline]
67. Schmidt C, Bladt F, Goedecke S, Brinkmann V, Zschiesche W, Sharpe M, Gherardi E, and Birchmeier C. Scatter factor/hepatocyte growth factor is essential for liver development. Nature 373: 699-702, 1995.[CrossRef][ISI][Medline]
68. Sergeant N, Lyon M, Rudland PS, Fernig DG, and Delehedde M. Stimulation of DNA synthesis and cell proliferation of human mammary myoepithelial-like cells by hepatocyte growth factor/scatter factor depends on heparan sulfate proteoglycans and sustained phosphorylation of mitogen-activated protein kinases p42/44. J Biol Chem 275: 17094-17099, 2000.
69. Shiota G, Rhoads DB, Wang TC, Nakamura T, and Schmidt EV. Hepatocyte growth factor inhibits growth of hepatocellular carcinoma cells. Proc Natl Acad Sci USA 89: 373-377, 1992.[Abstract]
70. Silvagno F, Follenzi A, Arese M, Prat M, Giraudo E, Gaudino G, Camussi G, Comoglio PM, and Bussolino F. In vivo activation of met tyrosine kinase by heterodimeric hepatocyte growth factor molecule promotes angiogenesis. Arterioscler Thromb Vasc Biol 15: 1857-1865, 1995.
71. Sonnenberg E, Meyer D, Weidner KM, and Birchmeier C. Scatter factor/hepatocyte growth factor and its receptor, the c-met tyrosine kinase, can mediate a signal exchange between mesenchyme and epithelia during mouse development. J Cell Biol 123: 223-235, 1993.[Abstract]
72. Sonnenberg E, Weidner KM, and Birchmeier C. Expression of the met-receptor and its ligand, HGF-SF during mouse embryogenesis. EXS 65: 381-394, 1993.[Medline]
73. Soriano JV, Pepper MS, Nakamura T, Orci L, and Montesano R. Hepatocyte growth factor stimulates extensive development of branching duct-like structures by cloned mammary gland epithelial cells. J Cell Sci 108: 413-430, 1995.
74. Stevenson BR, Siliciano JD, Mooseker MS, and Goodenough DA. Identification of ZO-1: a high molecular weight polypeptide associated with the tight junction (zonula occludens) in a variety of epithelia. J Cell Biol 103: 755-766, 1986.[Abstract]
75. Stoker M and Gherardi E. Factors affecting epithelial interactions. Ciba Found Symp 125: 217-239, 1987.[ISI][Medline]
76. Stoker M, Gherardi E, Perryman M, and Gray J. Scatter factor is a fibroblast-derived modulator of epithelial cell mobility. Nature 327: 239-242, 1987.[CrossRef][ISI][Medline]
77. Stoker M and Perryman M. An epithelial scatter factor released by embryo fibroblasts. J Cell Sci 77: 209-223, 1985.[Abstract]
78. Tajima H, Matsumoto K, and Nakamura T. Hepatocyte growth factor has potent anti-proliferative activity in various tumor cell lines. FEBS Lett 291: 229-232, 1991.[CrossRef][ISI][Medline]
79. Tajima H, Matsumoto K, and Nakamura T. Regulation of cell growth and motility by hepatocyte growth factor and receptor expression in various cell species. Exp Cell Res 202: 423-431, 1992.[ISI][Medline]
80. Terano A, Sakata-Horie K, Shimada T, Hiraishi H, Yoshiura K, Yoneda M, Takahashi M, and Fujimori T. The role of cellular migration in the repair process of gastric epithelial cells. Life Sci 69: 3083-3089, 2001.[CrossRef][ISI][Medline]
81. Ueda T, Takeyama Y, Toyokawa A, Kishida S, Yamamoto M, and Saitoh Y. Significant elevation of serum human hepatocyte growth factor levels in patients with acute pancreatitis. Pancreas 12: 76-83, 1996.[ISI][Medline]
82. Uehara Y, Minowa O, Mori C, Shiota K, Kuno J, Noda T, and Kitamura N. Placental defect and embryonic lethality in mice lacking hepatocyte growth factor/scatter factor. Nature 373: 702-705, 1995.[CrossRef][ISI][Medline]
83. Wanson JC, Drochmans P, Mosselmans R, and Ronveaux MF. Adult rat hepatocytes in primary monolayer culture. Ultrastructural characteristics of intercellular contacts and cell membrane differentiations. J Cell Biol 74: 858-877, 1977.[Abstract]
84. Weidner KM, Arakaki N, Hartmann G, Vandekerckhove J, Weingart S, Rieder H, Fonatsch C, Tsubouchi H, Hishida T, Daikuhara Y, and Birchmeier W. Evidence for the identity of human scatter factor and human hepatocyte growth factor. Proc Natl Acad Sci USA 88: 7001-7005, 1991.[Abstract]
85. Weidner KM, Behrens J, Vandekerckhove J, and Birchmeier W. Scatter factor: molecular characteristics and effect on the invasiveness of epithelial cells. J Cell Biol 111: 2097-2108, 1990.[Abstract]
86. Weidner KM, Sachs M, and Birchmeier W. The Met receptor tyrosine kinase transduces motility, proliferation, and morphogenic signals of scatter factor/hepatocyte growth factor in epithelial cells. J Cell Biol 121: 145-154, 1993.[Abstract]
87. Woolf AS, Kolatsi-Joannou M, Hardman P, Andermarcher E, Moorby C, Fine LG, Jat PS, Noble MD, and Gherardi E. Roles of hepatocyte growth factor/scatter factor and the met receptor in the early development of the metanephros. J Cell Biol 128: 171-184, 1995.[Abstract]
88. Yanagita K, Matsumoto K, Sekiguchi K, Ishibashi H, Niho Y, and Nakamura T. Hepatocyte growth factor may act as a pulmotrophic factor on lung regeneration after acute lung injury. J Biol Chem 268: 21212-21217, 1993.
89. Zarnegar R, DeFrances MC, Oliver L, and Michalopoulos G. Identification and partial characterization of receptor binding sites for HGF on rat hepatocytes. Biochem Biophys Res Commun 173: 1179-1185, 1990.[ISI][Medline]
90. Zarnegar R and Michalopoulos G. Purification and biological characterization of human hepatopoietin A, a polypeptide growth factor for hepatocytes. Cancer Res 49: 3314-3320, 1989.[Abstract]
91. Zegers MM, O'Brien LE, Yu W, Datta A, and Mostov KE. Epithelial polarity and tubulogenesis in vitro. Trends Cell Biol 13: 169-176, 2003.[CrossRef][ISI][Medline]
92. Zioncheck TF, Richardson L, Liu J, Chang L, King KL, Bennett GL, Fugedi P, Chamow SM, Schwall RH, and Stack RJ. Sulfated oligosaccharides promote hepatocyte growth factor association and govern its mitogenic activity. J Biol Chem 270: 16871-16878, 1995.