Regulation of RPE Intercellular Junction Integrity and
Function by Hepatocyte Growth Factor
Manlin Jin,1 Ernesto Barron,2 Shikun He,1 Stephen J. Ryan,2,3,4 and David R. Hinton1,2,4
PURPOSE. To evaluate the effect of hepatocyte growth factor
(HGF) on the integrity and function of tight junctions and
adherens junctions in the retinal pigment epithelial (RPE)
monolayer.
METHODS. Fresh bovine eyes were dissected to obtain 2- to
3-mm2 explants of intact RPE with underlying choroid and
sclera. Explants were cultured with or without HGF (20 ng/
mL) for various periods (20 minutes to 72 hours). Junction
integrity was assessed by transmission and scanning electron
microscopy; localization, expression, and phosphorylation of
junction proteins; and measurement of transepithelial resistance (TER), diffusion of fluorescent labeling in the plasma
membrane, and the migration of RPE cells from the monolayer.
RESULTS. Untreated explants consisted of polarized cells with
apical microvilli and well-developed tight and adherens junctions. After HGF treatment, the explants showed loss of tight
and adherens junctions ultrastructurally, diffusion of fluorescent label from apical to lateral membrane domains, and increased chemotactic migration of RPE cells from the monolayer. Primary cultures of confluent RPE cells showed a
progressive decrease in TER. Western blot analysis showed
rapid tyrosine phosphorylation of ZO-1, occludin, and ␤-catenin within 20 minutes of stimulation. There was a marked loss
of ZO-1 protein within 1 hour of HGF treatment. After 6 hours
of treatment with HGF, occludin, claudin-1, and ␤-catenin were
redistributed from the membrane to the cytoplasm.
CONCLUSIONS. Treatment of RPE explants with HGF results in
rapid disassembly of tight and adherens junctions associated
with loss or redistribution of junctional proteins, decreased
TER, and increased migration of RPE cells from the monolayer.
(Invest Ophthalmol Vis Sci. 2002;43:2782–2790)
R
etinal pigment epithelial (RPE) cells function, in part, to
provide a permeability barrier between retina and choroid
and to enable vectorial transport between these layers.1 Their
highly specialized junctional complexes separate cell surface
membrane proteins and lipids into apical and basolateral membrane domains.1 The tight junction, which is the most apical
component of the junctional complex, represents the anatomic substrate of the outer blood–retinal barrier.1
From the Departments of 1Pathology and 3Ophthalmology, the
Beckman Macular Research Center, and the 4Doheny Eye Institute,
Keck School of Medicine of the University of Southern California Los
Angeles, California.
Supported in part by Grant EY02061 and Core Grant EY03040
from the National Eye Institute, a grant from the Arnold and Mabel
Beckman Foundation, and a grant to the Department of Ophthalmology from Research to Prevent Blindness.
Submitted for publication October 9, 2001; revised March 14,
2002; accepted April 25, 2002.
Commercial relationships policy: N.
The publication costs of this article were defrayed in part by page
charge payment. This article must therefore be marked “advertisement” in accordance with 18 U.S.C. §1734 solely to indicate this fact.
Corresponding author: David R. Hinton, Department of Pathology,
Keck School of Medicine of the University of Southern California, 2011
Zonal Ave, HMR 209, Los Angeles, CA 90033; dhinton@hsc.usc.edu.
2
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A number of tight junction–associated proteins have been
identified and cloned, including the cytoplasmic anchor proteins ZO-1, -2, and -3 and the transmembrane protein occludin.
Occludin has a domain that binds to both ZO-1 and -2, which
in turn bind to the cytoskeleton. The extracellular domain of
occludin binds to another occludin molecule on an adjacent
cell, forming the tight junction responsible for the permeability
barrier.2– 4 The claudin family of transmembrane proteins has
also been identified as a critical component of this tight junction barrier function.5,6
Adherens junctions bind to a continuous belt of actin filaments (the adhesion belt), where they function to hold neighboring cells together through the family of Ca2⫹-dependent
cell– cell adhesion molecules known as cadherins.7 There are
many members of the classic cadherin family, but E-cadherin in
epithelial tissue is the best studied member in the context of
stable adhesion. Both continued expression and functional
activity of E-cadherin are required for cells to remain tightly
associated in the epithelium.7
To exhibit functional adhesive activity, cadherins must form
complexes with cytoplasmic plaque proteins called catenins
and with the actin cytoskeleton. ␤-Catenin is a critical regulatory component of the adherens junction, and its tyrosine
phosphorylation correlates with diminished adhesion in response either to growth factors or to cell transformation.7
Cultured RPE cells can form an intact monolayer with defined
barrier functions, and they can express junctional proteins,
including ZO-1, occludin, cadherins, and catenins.8,9
The expression of E-cadherin by RPE has been controversial. In comparison with other epithelial cells, RPE cells have
been reported not to express E-cadherin and to express Ncadherin instead.10,11 Burke et al.8,12 have reported that Ecadherin is found in adult human and bovine RPE in situ and in
postconfluent RPE cell cultures and that high E/P-cadherin
expression is associated with decreased apical polarity of Na,Kadenosine triphosphatase (ATPase).
Little is currently known about the regulation of junctions
in the intact RPE monolayer. Serum inhibits the formation of
tight junctions in cultured RPE, as do interferon-␥ and tumor
necrosis factor-␣.13,14 In cultures of other cell types, barrier
function is regulated by a variety of factors, including hepatocyte growth factor (HGF), vascular endothelial growth factor,
and histamine.15–18 However, the effect of these growth factors on preformed cell junctions in an intact epithelial monolayer has not been determined.
Hepatocyte growth factor/scatter factor is a glycoprotein,
usually produced by mesenchymal cells, that induces the proliferation, survival, dissociation, motility, and invasiveness of
epithelial and endothelial cells.19 The receptor for HGF is the
membrane-spanning tyrosine kinase c-met. The HGF/c-met system is critical for epithelial–mesenchymal interactions and has
widespread effects in embryogenesis and regeneration.19,20
The scatter effect of HGF was first found in the Madin-Darby
canine kidney (MDCK) cell line. When MDCK epithelial cells
are grown as small colonies at low density on impermeant
supports and then exposed to HGF, the cells assume a fibroblastic morphology and scatter away from the colonies.21 HGF
decreases the expression of occludin and transendothelial reInvestigative Ophthalmology & Visual Science, August 2002, Vol. 43, No. 8
Copyright © Association for Research in Vision and Ophthalmology
IOVS, August 2002, Vol. 43, No. 8
HGF Regulation of RPE Junction Integrity and Function
sistance in vascular endothelial cells15 and alters the polarity of
MDCK monolayers in two-dimensional culture,16 suggesting
that HGF is a critical regulator of monolayer functions. In vivo,
HGF is involved in angiogenesis and the branching morphogenesis of epithelial cells in the development of many organs.22
In pathologic conditions, HGF is associated with several benign
and malignant proliferative disorders. Its expression is increased in a variety of cancers, and it is believed to be related
to the invasion and metastasis of tumor cells.23
We and others have shown that cultured RPE cells express
the HGF receptor c-met and that they respond to HGF by
proliferation and chemotactic migration.24 c-Met is also expressed on RPE in epiretinal membranes from patients with
proliferative vitreoretinopathy (PVR),25 and HGF is elevated in
PVR vitreous fluids,26 suggesting that HGF/c-met may be involved in the pathogenesis of PVR. We initiated a study to
examine the effects of HGF on the intact monolayer and to
elucidate the mechanism by which RPE cells that adhere tightly
to one another in the monolayer can separate and migrate to
participate in membrane formation. In the current study, HGF
dramatically altered junctional integrity and function in the
RPE monolayer, and HGF promoted the ability of RPE to separate and participate in membrane formation.
MATERIALS
AND
METHODS
RPE Explant Culture and Treatment
Fresh bovine eyes were obtained from Shamrock Meats, Inc. (Vernon,
CA). Rat eyes were enucleated from albino rats (Charles River, Boston,
MA). All procedures adhered to the ARVO Statement for the Use of
Animals in Ophthalmic and Vision Research.
The eyes were immersed in Hanks’ balanced salt solution (HBSS;
Irvine Scientific, Santa Ana, CA) containing 1% penicillin and streptomycin for 10 minutes. Under a dissecting microscope in sterile conditions, the vitreous humor and neural retina were carefully removed
without disturbing the RPE layer. The eyes were then cut into 2- to
3-mm2 explants, each with an intact RPE layer, choroid, and sclera. The
explants were immediately placed in high-glucose Dulbecco’s modified Eagle’s medium (DMEM; Irvine Scientific) supplemented with
penicillin G (100 U/mL), L-glutamine (0.292 mg/mL), and 1% fetal
bovine serum (FBS; Irvine Scientific). Explants were cultured with or
without HGF (20 ng/mL; R&D Systems, Minneapolis, MN) for various
amounts of time, up to 72 hours. We have shown that these explants
remain viable and can respond to growth factor stimulation for at least
72 hours.27
Electron Microscopy
Explants were fixed in half-strength Karnovsky fixative (2.5% glutaraldehyde and 2% paraformaldehyde in 0.1 M phosphate buffer [pH
7.2–7.4]) at 4°C for 48 hours. Specimens prepared for transmission
electron microscopy (TEM) were postfixed in 1% osmium tetroxide for
2 hours, dehydrated in a series of graded alcohols, embedded with an
epoxy resin kit (PolyBed 812; Polyscience, Warrington, PA), thin sectioned, stained with uranyl acetate, and examined with a transmission
electron microscope (model EM10B; Carl Zeiss, Thornwood, NY).
Specimens prepared for scanning electron microscopy (SEM) were
postfixed in 1% osmium tetroxide, dehydrated, critical point dried,
coated with gold palladium, and examined with a scanning electron
microscope (model S-570; Hitachi, Tokyo, Japan). For TEM and SEM, at
least three control- and three HGF-treated explants were examined. For
TEM, at least 10 sections were examined for each explant.
Fluorescent Lipid Labeling of the
Plasma Membrane
The diffusion barrier between the apical and basolateral domains was
evaluated by using the fluorescent probe FM1-43 (Molecular Probes,
2783
Inc., Eugene, OR). FM1-43 inserts into the outer leaflet of the lipid
bilayer, but it cannot flip-flop to the inner layer; thus, it cannot permeate an intact tight junction.28,29 When incubation periods are short,
FM1-43 is a valuable marker of an intact tight junction.30,31 After
removal of the retina, FM1-43 (2 ␮M) was added to the culture medium
bathing the bovine eyecup. After a 5-minute incubation, the eyecup
was washed with cold phosphate-buffered saline (PBS), dissected, and
embedded in optimal cutting temperature (OCT) compound (TissueTek, Torrance, CA). Eight-micrometer serial sections were prepared
and evaluated with a confocal microscope (model LSM510; Zeiss) with
a 40⫻ oil-immersion objective. Three independent experiments were
performed.
Transepithelial Resistance
Fresh bovine RPE cells were isolated, as previously described32 and
plated at a density of 2 ⫻ 105 cells/cm2 on a 24-mm permeable
membrane insert (12-mm diameter, 0.4-mm pore size, Transwell;
Costar, Cambridge, MA) coated with laminin (5 ␮g/cm2; Becton Dickinson Laboratory, Franklin Lakes, NJ) in DMEM with 10% FBS. After the
RPE cells became confluent, the medium was changed to DMEM with
1% FBS until the transepithelial resistance (TER) stabilized. At this stage
of culture, the RPE cells expressed prominent membrane-associated
ZO-1 and occludin (results not shown). The TER was monitored with
an epithelial voltohmeter (World Precision Instruments, Sarasota, FL)
at regular intervals and was corrected for background resistance contributed by the blank filter and culture medium. Three independent
experiments were performed. Results are expressed as mean ohms 䡠
cm2 ⫾ SD and compared by Student’s t-test. Significance was set at P ⬍
0.05.
Migration Assay
Migration of RPE cells from the monolayer was detected by Boyden
chamber assay, as previously described,33 modified for explant culture.
Briefly, 24-well cell culture inserts (Fisher Scientific, Fair Lawn, NJ)
were coated with fibronectin (5 ␮g/cm2). Explants were placed RPE
side down in the upper compartment. Platelet-derived growth factor
(PDGF, 20 ng/mL, R&D Systems) was added to the lower compartment
as a chemotaxis stimulus. After 12 hours of incubation, the inserts were
washed three times with PBS, fixed with pure cold methanol for 10
minutes, and counterstained with hematoxylin for 20 minutes. Cells on
the top surface of the insert membrane were removed by wiping with
a cotton swab. The number of migrating cells was determined by
phase-contrast microscopy at 320⫻ high-power field. Four randomly
chosen fields were counted per insert. Three independent experiments were performed. Comparisons were made by Student’s t-test
with significance set at P ⬍ 0.05.
Immunofluorescence Microscopy
The expression and localization of the tight junction proteins were
examined by qualitative analysis of immunofluorescence staining. Sections from at least five explants were used in all cases. Explants were
fixed in 2% paraformaldehyde in PBS for 15 minutes at room temperature, permeabilized in 0.2% Triton X-100 in PBS for 15 minutes, and
incubated with the primary antibody for 60 minutes at 4°C and the
secondary antibody for 30 minutes at 4°C. Primary antibodies were
diluted in PBS: anti-ZO-1 (1:100; Chemicon, Temecula, CA), anti-occludin (1:100; Zymed Laboratory Inc, San Francisco, CA), anti-claudin-1
(1:100, Cat. No. 51-9000; Zymed Laboratory Inc.), anti-E-cadherin
(clone 34, 1:100; Transduction Laboratories, Lexington, KY), and anti␤-catenin (1:100; Transduction Laboratories). The secondary antibodies were rhodamine-conjugated goat-anti-rabbit (1:400; Chemicon) and
FITC-conjugated goat-anti-mouse (1:400; Chemicon). The slides were
washed in PBS, mounted, and examined with a confocal microscope
(Carl Zeiss) with a 40⫻ oil-immersion objective.
Immunoprecipitation and Western Blot Analysis
Western Blot Analysis. The explant was homogenized in 100
␮L modified RIPA buffer (50 mM Tris-HCl (pH 7.4), 1% Triton X-100,
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Jin et al.
IOVS, August 2002, Vol. 43, No. 8
FIGURE 1. HGF-induced disassembly
of junctions detected by electron microscopy. Scanning electron micrograph of the control RPE explant (A)
shows polygonal cells with well-developed apical microvilli. After treatment
with HGF for 60 minutes, the RPE explant cells (B) showed retraction of
apical membrane, loss of apical microvilli, and focal areas of RPE separation, appearing as a hole in the center
of the image. Transmission electron
micrograph of control RPE cells (C)
shows well-developed tight and adherens junctions, and abundant, long apical microvilli (mv). High magnification
shows an intact junctional complex
(E). After treatment with HGF for 60
minutes, the RPE cell showed fewer
apical microvilli (D). High magnification shows loss of tight and adherens
junction structure, a reduced junctionassociated actin belt, and focal separation between cells in the region of the
junction (F). Arrows: tight junction;
arrowheads: adherens junction. Bar
applies to (A) and (B). Magnification:
(C–F) ⫻2000.
0.2% sodium dodecyl sulfate [SDS], 1 mM dithiothreitol, 2 mM EGTA,
4 mM EDTA, 2 mM sodium orthovanadate, and 100 mM NaCl) with
fresh protease inhibitors, including 0.2 mM phenylmethylsulfonyl fluoride (PMSF) and 10 ␮g/mL each of aprotinin, pepstatin A, and soybean trypsin inhibitor (Sigma, St. Louis, MO). Samples were incubated
at 4°C for 30 minutes to solubilize proteins, and insoluble materials
were pelleted by centrifugation at 14,000g for 10 minutes. Protein
concentration was determined with a protein assay (Bio-Rad, Richmond, CA), and equal total protein content was loaded into 7.5%
SDS-polyacrylamide gels (SDS-PAGE; Bio-Rad). Proteins were transferred to polyvinylidene difluoride membranes (Millipore, Bedford,
MA), blocked by 1% bovine serum albumin for 60 minutes, and incubated with the primary antibodies of anti-ZO-1, anti-occludin, anticlaudin-1, anti-E-cadherin, and anti-␤-catenin, followed by horseradish
peroxidase– conjugated secondary antibody. The membranes were
then washed and developed with the addition of a chemiluminescence
detection kit (Amersham Pharmacia Biotech, Piscataway, NJ).
Triton X-100 Extraction Assay. To make soluble or insoluble
pools, RPE explants were lysed in 0.5% Triton X-100 containing 50 mM
Tris-HCl, 2 mM sodium orthovanadate, 50 mM NaCl, and fresh protease
cocktail (described earlier) plus phosphatase inhibitor cocktail (Sigma)
and then centrifuged at 14,000g for 10 minutes. This supernatant was
considered the Triton X-100 –soluble pool. The pellet was solubilized
in 1% SDS and referred to as the Triton X-100 –insoluble pool.
Immunoprecipitation. RPE explants were treated with HGF
for various periods and lysed in the buffer used in Western blot
experiments for 30 minutes. The insoluble materials were removed by
centrifugation at 14,000g, and the supernatants were collected. Equal
amounts of protein were clarified and incubated with anti-occludin,
anti-ZO-1, or anti-␤-catenin antibody overnight at 4°C with the rotor
on. The immune complexes were collected using anti-rabbit agarose
beads for 2 hours and centrifuged at 10,000g for 2 minutes. After
extensive washing with the same buffer, Laemmli sample buffer (BioRad) was added to the residual agarose with immune complexes and
boiled at 95°C for 5 minutes. These samples were loaded onto SDS
polyacrylamide gels and Western blotting followed as described above
using anti-phosphotyrosine primary antibody (Sigma). In a parallel
experiment, blots (n ⫽ 3) used to detect the phosphorylation state of
ZO-1 were imaged, placed in protein stripping buffer (100 mM 2-mercaptoethanol, 2% SDS, 65 mM Tris-HCl [pH 6.8]), and then reprobed
with anti-ZO-1 antibody.
Densitometric and Statistical Analysis
Densitometry was performed with a scanner (Hewlett-Packard, Palo
Alto, CA) and quantitated by the an image analysis software program
(Scion Corp., Frederick, MD). For quantitative data, each assay was
repeated at least three times and compared by Student’s t-test. The
significance was set at P ⬍ 0.05.
RESULTS
HGF-Induced Disassembly of Tight
and Adherens Junctions
SEM of the control explants (n ⫽ 3) revealed that the RPE cells
had uniform polygonal shape and abundant apical microvilli
(Fig. 1A). Dramatic changes in the ultrastructural appearance
of the RPE cells were observed after 60 minutes of HGF treatment of the explants (n ⫽ 3). The RPE cells showed a loss of
apical microvilli and retraction of the apical membrane, allowing us to see the lateral membranes and areas in which RPE
cells separate from each other (Fig. 1B).
TEM (three explants, 10 sections each) revealed that the
control RPE explant cells had well-developed apical RPE tight
and adherens junctions and a junction-related actin filament
belt. The apical surface of the cells had abundant, long microvilli (Fig. 1C, 1E). After treatment with HGF for 60 minutes,
TEM sections (five explants, 10 sections each) revealed that the
RPE cells had lost their defined tight and adherens junction
structures. The RPE cells showed a reduced junction-associated
actin belt and fewer apical microvilli (Fig. 1D, 1F).
Tight junctions contribute to the establishment of epithelial
cell surface polarity and inhibit the diffusion of lipids in the
outer leaflet of the plasma membrane between apical and
basolateral membrane domains.29 To confirm the HGF-induced
disruption of barrier function, the fluorescent lipid probe
FM1-43 was used to pulse label the apical plasma membranes.
The control cells showed only apical membrane labeling of FM
1-43 (Fig. 2A). After HGF treatment of the RPE for 60 minutes,
pulse label of the apical membrane resulted in prominent
labeling of both apical and lateral membranes (Fig. 2B), indi-
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HGF Regulation of RPE Junction Integrity and Function
FIGURE 2. Diffusion of fluorescent label from apical to basolateral
domains after treatment of RPE with HGF. A bovine eyecup was
pulse-labeled by FM1-43 for 5 minutes, embedded in OCT, cut into
8-␮m serial sections, and evaluated by confocal microscopy. The control cells showed label limited to the apical membrane (A). After HGF
treatment for 60 minutes, pulse labeling with FM1-43 resulted in
diffusion of the label to the lateral membranes (B), indicating loss of
the barrier between apical and basolateral domains. Arrows: apical and
basal surfaces of the RPE monolayer. Magnification, ⫻400.
cating loss of the diffusion barrier between these domains.
Three independent experiments were performed.
HGF-Induced Decrease in TER
The loss of tight junctions is associated with a decrease in
TER.12 The TER of cultured bovine RPE cells was measured at
30 minutes and 1, 3, 24, 48, and 72 hours after treatment with
HGF (Fig. 3). The TER of control RPE cells remained stable (180
⍀/cm2) throughout the experiments. A reduction of TER was
first seen after 60 minutes of treatment with HGF (P ⬍ 0.05).
Continuous culture of RPE cells with HGF for longer periods
resulted in further reduction of TER, with a maximum effect
after 24 hours of treatment (P ⬍ 0.01) that was maintained to
72 hours (P ⬍ 0.01).
HGF-Induced Migration of RPE Cells from the
Explant Monolayer
The adherens junction plays a critical role in intercellular
adhesion and functions to hold neighboring cells together, thus
preventing easy dissociation of RPE from the monolayer.7 We
modified the Boyden chamber assay to test the migration of
RPE cells from the monolayer in response to a chemotactic
FIGURE 3. HGF-induced decrease of TER. The effect of HGF on TER of
bovine RPE cells was measured 30 minutes and 1, 3, 24, 48, and 72
hours after stimulation. The control cells showed stable TER throughout the experiment. However, HGF treatment induced a significant
decrease of TER beginning at 60 minutes (P ⬍ 0.05), with a further
decrease at 180 minutes (P ⬍ 0.01) and a maximum effect at 24 hours
(P ⬍ 0.01) that was maintained to 72 hours (P ⬍ 0.01). Data are the
mean ⫾ SD of results in three independent experiments.
2785
FIGURE 4. HGF-induced increase in RPE cell migration from an intact
RPE explant in a modified Boyden chamber assay. The control explant
had very few migrating cells. After treatment with HGF for 6 hours,
there was significantly increased cell migration from the monolayer in
response to PDGF (P ⬍ 0.001). Data are the mean ⫾ SD of three
independent experiments.
agent after HGF treatment. To maximize the migration of RPE
cells from the control explant, we used optimal conditions, as
previously published,26 including the use of PDGF as a chemotaxis stimulus and the use of fibronectin as a coating for the
intervening porous membrane. Under those conditions, very
few cells migrated from the control explant. After 6 hours of
HGF treatment, the RPE explant showed a sixfold increase in
the number of migrating cells (Fig. 4, P ⬍ 0.001).
Immunofluorescent Staining of Tight
Junction Proteins
The expression and localization of the tight junction proteins
were examined by qualitative assessment of immunofluorescent staining. Results shown are representative of five independent experiments. The control RPE explant cells showed a
clear membrane-associated staining pattern for ZO-1, and the
cells had a uniform polygonal shape (Fig. 5A). After 60 minutes
of HGF treatment, the RPE explant cells showed decreased
continuous membrane staining (Fig. 5B). Empty spaces surrounded by three or more abnormally shaped RPE cells were
occasionally observed, suggesting that an individual cell had
been displaced from the monolayer. Quantitative analysis revealed that such empty spaces represented 2% to 3% of the RPE
cell population after 60 minutes of HGF treatment. After treatment with HGF for 6 hours, ZO-1 staining was much weaker
(Fig. 5C). The control RPE explant cells showed a membraneassociated staining pattern for occludin similar to that seen
with ZO-1, but the pattern with occludin was more granular
(Fig. 5D). After 60 minutes of HGF treatment, occludin staining
was still intact (Fig. 5E). After 6 hours of HGF treatment,
occludin showed a loss of association with the membrane,
appearing instead in a diffuse, cytoplasmic pattern (Fig. 5F).
The control RPE explant cells showed a membrane-associated
pattern for claudin-1 staining (Fig. 5G). After 60 minutes of
HGF treatment, claudin-1 staining was still intact (Fig. 5H).
After 6 hours of HGF treatment, claudin-1 staining showed a
loss of linear membrane-associated pattern, with the development of punctate aggregates (Fig. 5I).
Immunofluorescent Staining of Adherens
Junction Proteins
The expression and localization of adherens junction proteins
are shown Figure 6. Results are representative of five independent experiments. The control RPE explant cells showed a
membrane-associated pattern of ␤-catenin staining (Fig. 6A).
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Jin et al.
FIGURE 5. Immunofluorescent staining of tight junction proteins in
RPE explants. (A) The control RPE explant cells showed clear membrane-associated staining of ZO-1. (B) After 60 minutes of HGF treatment, the explant started to lose ZO-1 immunoreactivity. Occasional
empty spaces were observed, surrounded by three or more abnormally
shaped RPE cells, suggesting loss of an individual cell from the monolayer (arrows). Subsequent micrographs were taken to avoid areas of
cell loss, so that the intact monolayer could be observed. (C) After 6
hours of treatment, the ZO-1 staining was much weaker. Occludin and
claudin-1 staining patterns were similar. The control RPE explant cells
(D, G) showed clear membrane-associated staining. After 60 minutes of
treatment with HGF (E, H), the RPE explant cells showed intact,
well-maintained, membrane-associated staining. After 6 hours of treatment, the explant showed loss of membrane-associated staining of
occludin and diffuse staining in the cytoplasm (F), and claudin-1
staining showed loss of linear membrane-associated pattern and development of punctate aggregates (I). Magnification, ⫻400.
After 60 minutes of HGF treatment, ␤-catenin staining was
intact (Fig. 6B). After 6 hours of HGF treatment, there was loss
of membrane ␤-catenin staining, which instead appeared as
IOVS, August 2002, Vol. 43, No. 8
FIGURE 7. Western blot analysis of tight and adherens junction proteins after treatment of RPE explants with HGF. HGF induced a significant loss of ZO-1 protein after treatment for 60 minutes and 6 hours
(P ⬍ 0.01) and of occludin protein (P ⬍ 0.05) after 6 hours. There was
no significant change in claudin-1, ␤-catenin, or E-cadherin protein.
Blots were quantified by densitometric analysis. Data are representative of results in three separate experiments. Primary data are shown in
Table 1.
diffuse cytoplasmic staining (Fig. 6C). The control explants
showed a membrane-associated localization for E-cadherin (Fig.
6D). After 60 minutes of HGF treatment, E-cadherin staining
was intact (Fig. 6E). After 6 hours of treatment, E-cadherin
staining was weaker with cytoplasmic aggregation (Fig. 6F).
Western Blot Analysis of Junctional Proteins
Quantitation of junctional proteins was determined by Western
blot analysis. ZO-1 was identified as a single 220-kDa band,
occludin as two bands of approximately 60 to 65 kDa, claudin-1 as a single 22-kDa band, E-cadherin as a 120-kDa band,
and ␤-catenin as a 95-kDa band. As shown in Figure 7, the
amount of ZO-1, occludin, claudin-1, E-cadherin, and ␤-catenin
was not significantly changed after treatment with HGF for
either 20 minutes or 40 minutes. The amount of ZO-1 protein
detected by the Western blot analysis was decreased by 50%
(P ⬍ 0.01) and by 80% (P ⬍ 0.01), after treatment with HGF for
60 minutes and 6 hours, respectively. After 6 hours of HGF
treatment, the amount of occludin protein was also decreased
by 25% (P ⬍ 0.05). There was no significant change in the
amount of claudin-1, ␤-catenin, and E-cadherin protein after
HGF treatment. All experiments were repeated three times;
densitometric data are shown as a percentage of the control in
Table 1.
HGF-Induced Redistribution
of Junctional Proteins
FIGURE 6. Immunofluorescent staining of adherens junction proteins
in RPE explants. (A) The control RPE explant cells showed membraneassociated staining of ␤-catenin. (B) After treatment with HGF for 60
minutes, the explant showed intact ␤-catenin staining. (C) After treatment for 6 hours, the explant showed loss of membrane-associated
␤-catenin staining; however, diffuse cytoplasmic staining became apparent. (D) The control RPE explant cells showed a membrane-associated localization of E-cadherin. (E) After treatment with HGF for 60
minutes, E-cadherin staining was still intact. (F) After treatment for 6
hours, there was weaker staining of E-cadherin compared with control,
and there was cytoplasm aggregation. Magnification, ⫻400.
As shown by immunofluorescent staining, junctional proteins
appeared to lose their association with plasma membranes.
Triton X-100 extraction assay was used to identify the redistribution of junctional proteins from a cytoskeleton-associated
membrane region to the cytoplasm based on changes in solubility.34,35 The RPE explants were fractionated in 0.5% Triton
X-100 and divided into soluble (S) and insoluble (I) pools (Fig.
8A). After treatment with HGF for 1 hour and 6 hours, a
significant decrease of ZO-1 protein was detected in the insoluble pool (P ⬍ 0.01). After treatment with HGF for 6 hours,
there were significant decreases of occludin (P ⬍ 0.01), claudin-1 (P ⬍ 0.05), ␤-catenin (P ⬍ 0.01), and E-cadherin (P ⬍
HGF Regulation of RPE Junction Integrity and Function
IOVS, August 2002, Vol. 43, No. 8
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TABLE 1. Densitometry Data for Western Blot of Junctional Proteins
ZO-1
Occludin
Claudin-1
␤-Catenin
E-Cadherin
Control
HGF (20 min)
HGF (40 min)
HGF (1 h)
HGF (6 h)
100
100
100
100
100
95.1 ⫾ 2.8
93.6 ⫾ 11.4
97.4 ⫾ 9.5
100.8 ⫾ 8.6
96.7 ⫾ 4.6
90.1 ⫾ 8.6
95.8 ⫾ 5.7
97.3 ⫾ 3.1
97.4 ⫾ 5.0
91.4 ⫾ 4.0
55.1 ⫾ 16.8*
94.2 ⫾ 3.9
92.8 ⫾ 9.9
95.7 ⫾ 2.1
87.7 ⫾ 9.1
20.2 ⫾ 10.9†
75.6 ⫾ 9.5*
90.2 ⫾ 11.2
91.3 ⫾ 5.5
86.5 ⫾ 7.1
Densitometric analysis after treatment with HGF revealed a significant decrease in the amount of ZO-1 protein after HGF treatment for 1 hour
and 6 hours, and amount of occludin protein after HGF treatment for 6 hours. Data are representative of mean results in three separate experiments
and are shown as normalized percentage to control.
* P ⬍ 0.05.
† P ⬍ 0.01.
0.05) protein in the insoluble pool. Blots were quantified by
densitometric analysis, and measurements were expressed as a
percentage of total density of both fractions (I/S⫹I) (Fig. 8B).
HGF-Induced Increase in Tyrosine
Phosphorylation of ZO-1, Occludin,
and ␤-Catenin
Because junctional protein function and solubility are highly
regulated by phosphorylation, HGF-induced phosphorylation
may represent the mechanism by which the tight junctions are
disassembled. RPE explants were treated with HGF for various
amounts of time and extracted with a buffer containing both
1% Triton X-100 and 0.2% SDS. Cell extracts were then immunoprecipitated with anti-ZO-1, anti-occludin, or anti-␤-catenin
antibody. Western blots of the immunoprecipitates were
probed with anti-phosphotyrosine antibody. There was a significant increase in the phosphotyrosine content of phosphorylated ZO-1, occludin, and ␤-catenin after as little as 20 minutes
of HGF treatment (Fig. 9A). Blots were quantified by densitometric analysis, and data are shown as a percentage of the
control in Table 2. In a parallel experiment repeated three
times, the immunoprecipitation blot was stripped and reprobed with ZO-1 antibody to confirm the concurrent decrease
in ZO-1 protein in the sample after 60 minutes of HGF treatment (Fig. 9B). No tyrosine phosphorylation of claudin-1 and
E-cadherin was detected (data not shown).
FIGURE 8. Analysis of junctional protein solubility in Triton-X after
treatment with HGF. Triton X-100 soluble (S) and insoluble (I) fractions
were separated by 7.5% SDS-PAGE and transferred to membranes. (A)
Blots probed with antibodies specific for ZO-1, occludin, claudin-1,
E-cadherin and ␤-catenin. Data are representative of results in three
separate experiments. After treatment with HGF for 1 hour and 6
hours, a significant decrease of ZO-1 protein in the insoluble pool (P ⬍
0.01) was detected. After treatment for 6 hours, a significant decrease
of occludin (P ⬍ 0.01), claudin-1 (P ⬍ 0.05), ␤-catenin (P ⬍ 0.01), and
E-cadherin (P ⬍ 0.05) protein was seen in the insoluble pool. (B)
Densitometric quantification of blots, with measurements expressed as
a percentage of total density of both fractions (I/S⫹I). *P ⬍ 0.05 and
**P ⬍ 0.01 versus control.
FIGURE 9. Effect of HGF on the tyrosine phosphorylation of junctional
proteins in RPE explants. (A) The tyrosine phosphorylation of ZO-1,
occludin, and ␤-catenin showed a significant increase after treatment
with HGF for as long as 60 minutes. Data are representative of results
in three separate experiments. Data from densitometric analysis of
blots are shown in Table 2. (B) In a parallel experiment, blots used to
detect the tyrosine phosphorylation (PY) state of ZO-1 were placed in
protein stripping buffer and then reprobed with anti-ZO-1 antibody,
confirming that increased tyrosine phosphorylation of ZO-1 was associated with a decreased amount of ZO-1 protein.
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Jin et al.
IOVS, August 2002, Vol. 43, No. 8
TABLE 2. Densitometry Data for PY Content of Junctional Proteins
Control HGF (20 min) HGF (40 min)
ZO-1
Occludin
␤-Catenin
100
100
100
188.8 ⫾ 30.2*
193.9 ⫾ 12.2†
179.7 ⫾ 27.6*
184.0 ⫾ 23.6*
189.7 ⫾ 14.5*
245.8 ⫾ 33.9*
HGF (1 h)
225.5 ⫾ 24.0†
250.3 ⫾ 22.3†
221.9 ⫾ 21.8†
Densitometric analysis after treatment with HGF revealed a significant increase in tyrosine phosphorylation of ZO-1, occludin and
␤-catenin after HGF treatment for up to 60 minutes. Data are representative of mean results in three separate experiments and shown as
normalized percentage to control.
* P ⬍ 0.05.
† P ⬍ 0.01.
DISCUSSION
The RPE monolayer constitutes the outer blood–retinal barrier;
thus, its integrity is critical to the normal functioning of the
neural retina. This monolayer is characterized by well-organized intercellular tight and adherens junctions between RPE
cells.1 The RPE explant is a good model for investigating
disruption of these junctions. Compared with other cell culture monolayer models, such as confluent MDCK cells or vascular endothelial cells, the junctions of RPE explants are preformed and are closer to the normal physiological condition.
With this model, we provide evidence that HGF treatment of
the RPE monolayer induces disassembly of tight and adherens
junctions, both morphologically and physiologically.
This effect appeared to be mediated primarily by the action
of HGF on the cytoplasmic membrane plaque protein ZO-1.
ZO-1 is a peripheral membrane protein localized to the tight
junction complex in epithelial and endothelial cells. Anchoring
of ZO-1 with the underlying cytoskeleton is required for localization of occludin and claudin in the tight junction. ZO-1, -2,
and -3 contain three PDZ domains, one SH3 domain, and one
guanylyl kinase-like domain (GuK). Through its GuK domains,
ZO-1 binds directly to the carboxyl termini of claudins and
occludin and may function as an adaptor at the cytoplasmic
surface of the tight junction to recruit other proteins, including
cytoskeletal and signaling molecules.4 These components can
form a huge macromolecular complex at the cytoplasmic surface of tight junctions and may be involved in the regulation of
endothelial and epithelial cell polarization, proliferation, and
differentiation.4 As an adaptor, ZO-1 is a critical regulatory
protein between occludin or claudin and the cytoskeleton or
signaling molecules. Loss of ZO-1 may break down this linkage
and lead to the rapid morphologic and physiological changes
induced by HGF. The most notable early change in this study
was the rapid loss of ZO-1 protein after treatment with HGF for
60 minutes. The loss of ZO-1 protein preceded the redistribution of occludin and claudin-1 from the tight junction. The time
frame of this redistribution was consistent with the detected
morphologic and physiological changes. These changes, therefore, suggest that HGF may first target and induce degradation
of ZO-1 protein and that this degradation is followed by the
redistribution of occludin and claudin and the physiological
disruption of the tight junctions. The loss of junctional proteins
has been shown to play a critical role in the disassembly of
tight junctions in other cells. Agents such as histamine induce
a rapid decrease in ZO-1 protein, leading to an increase of
paracellular permeability in retinal endothelial cells.17 Phosphorylation of ZO-1 is also a critical regulator in the assembly
and disassembly of tight junctions.36,37 The tyrosine-phosphorylated form of ZO-1 may cause a dismantling of tight junctions.38,39 Our study found that HGF induced a marked increase
in tyrosine phosphorylation, but there was no detectable phosphorylation of serine and threonine on immunoprecipitated
ZO-1 (data not shown). The hyperphosphorylation of ZO-1
may contribute to its degradation, similar to the degradation of
p27kip1 and p57kip2.40 Proteasome inhibitors have been shown
to stabilize the HGF-induced disassembly of junctions in MDCK
cells, consistent with the notion that protein degradation plays
a part in junction breakdown.41
Occludin is a critical paracellular component of tight junction barrier function. Expression of occludin is high in vessels
with tight barriers, such as those of the brain,42 overexpression
of occludin by transfection increases TER in kidney cells,43 and
microinjection of COOH-terminal truncated occludin into oocytes causes a dominant negative effect on micromolecular
tissue barrier formation.44 In our experiment, treatment with
HGF altered the function of the tight junctions in the RPE
monolayer. This altered function may be caused either by the
rapid phosphorylation and redistribution of occludin from
membrane to cytoplasm or by a slower loss of occludin protein
content. The phosphorylation status of occludin has been
shown to be a critical regulator of its distribution in tight
junctions and cytoplasm. Phospho-amino acid analysis suggests
that occludin is phosphorylated predominantly on serine-threonine residues in MDCK cells.45 However, more recent studies
have shown that tyrosine phosphorylation of occludin is important for tight junction reassembly.32,46 In MDCK cells, occludin is heavily tyrosine phosphorylated, with a time course
similar to that of the formation of tight junctions after the
application of a calcium switch.46 In vascular endothelial cells,
HGF has been shown to induce increased tyrosine and serinethreonine phosphorylation of occludin associated with decreased TER.15
In our studies, HGF stimulation was associated with rapid
tyrosine phosphorylation of occludin. Preliminary immunoblot
experiments using anti-phosphoserine antibody (Sigma) demonstrated weak serine phosphorylation of occludin, but this
did not appear to be modulated by stimulation with HGF
(results not shown). Further characterization of serine-threonine phosphorylation sites on RPE-derived occludin and the
modulation of these sites by HGF by gel shift assay37,45 or
improved phosphoserine-threonine antibodies would be of interest. In RPE, the effect of HGF on occludin may relate in part
to the confluence of the monolayer. It has been reported that
subconfluent RPE cultured on type I collagen and treated with
HGF for 1 week maintain an epithelial phenotype and show
increased occludin expression localized to the cell membrane.47 Although occludin has been shown to be essential in
the barrier function of tight junctions, other studies indicate
that one or more other proteins are involved in the formation
of tight junctions in the absence of occludin. Occludin-deficient embryonic stem cells differentiate into polarized epithelial cells with well-developed tight junction strands.48
Claudin-1 and -2 have recently been identified as transmembrane proteins localizing at tight junctions of epithelium, and
more than 20 claudin-like proteins have been identified.4 We
showed that RPE explants express claudin-1 and that HGF
induces similar changes in claudin-1 and occludin, including
loss of the distinct membrane-associated pattern and redistribution away from the junctional area.
Loss of tight junctions correlates with the loss of polarity in
the RPE monolayer, as shown in SEM by the retraction of apical
microvilli and separation between the cells. The mechanism of
the changes observed in apical microvilli is unknown, but it
could be related to changes in expression or phosphorylation
of ezrin. Ezrin is a member of the ezrin-radixin-moesin family,
which localizes to the microvilli of RPE cells in vivo and
regulates their morphology.49 Ezrin has been shown to be a
substrate of HGF receptor and plays a critical role in HGFinduced morphogenesis in kidney cells.50
IOVS, August 2002, Vol. 43, No. 8
HGF Regulation of RPE Junction Integrity and Function
Intercellular adhesion, which plays a key role in maintaining
tissue integrity and morphology of RPE cells, is facilitated
primarily by the adherens junction. Because the adherens junction is critical for cell– cell adhesion, it has been extensively
investigated in cancer biology. Both in vitro and in vivo studies
have shown that a loss or a reduction of E-cadherin or ␤-catenin expression is associated with invasion and metastasis in
various cancers.51–53 In our study, HGF induced disassembly of
the adherens junctions, a rapid increase in tyrosine phosphorylation of ␤-catenin (20 minutes), and a redistribution of ␤-catenin and E-cadherin from membrane to cytoplasm (6 hours).
The function of E-cadherin is mediated intracellularly by its
interaction with cytoplasm catenins. At the cell surface, Ecadherin forms an adherens junction that is intracellularly
linked to the actin cytoskeleton by the catenins. Data from
other cell types have shown that HGF-induced phosphorylation of ␤-catenin may cause disassociation of ␤-catenin from
the E-cadherin complex to the cytoplasm, leading to abnormal
function of the adherens junction.54 Disassembly of adherens
junctions leads to the loss of adhesion between RPE cells in the
monolayer and greatly increases the ability of RPE cells to
migrate from the monolayer in response to fibronectin and the
chemotactic agent PDGF. Both fibronectin and PDGF are prominently expressed in PVR epiretinal membranes.55,56 This work
suggests that prior exposure to HGF could dramatically facilitate the ability of PDGF and fibronectin to induce migration of
RPE cells from the monolayer.
We have demonstrated that HGF induces disassembly of
tight and adherens junctions in an intact RPE monolayer and
that this disassembly is associated with a decreased TER and an
increased ability to migrate from the monolayer. The effect of
HGF is likely to be targeted primarily at the ZO-1 protein. These
results, in conjunction with studies showing increased HGF
expression in PVR, suggest that HGF may play a critical role in
the pathogenesis of PVR.
12.
13.
14.
15.
16.
17.
18.
19.
20.
21.
22.
Acknowledgments
The authors thank Laurie Labree for statistical consultation and Chris
Spee for technical support.
23.
24.
References
1. Thumann G, Hinton DR. Cell biology of the retinal pigment epithelium. In: Ryan SJ, ed. Retina. Vol 1. 3rd ed. St. Louis: Mosby;
2001:104 –121.
2. Stevenson BR, Kean BH. The tight junction: morphology to molecules. Annu Rev Cell Dev Biol. 1998;14:89 –109.
3. Kniesel U, Wolburg H. Tight junctions of the blood-brain barrier.
Cell Mol Neurobiol. 2000;20:57–76.
4. Tsukita S, Furuse M, Itoh M. Mutifunctional strands in tight junctions. Nat Rev Mol Cell Biol. 2001;4:285–293.
5. Furuse M, Fujita K, Hiiragi T, Fujimoto K, Tsukita S. Claudin-1 and
-2: novel integral membrane proteins localizing at tight junctions
with no sequence similarity to occludin. J Cell Biol. 1998;141:
1539 –1550.
6. Inai T, Kobayashi J, Shibata Y. Claudin-1 contributes to the epithelial barrier function in MDCK cells. Eur J Cell Biol. 1999;78:849 –
855.
7. Gumbiner BM. Cell adhesion: the molecular basis of tissue architecture and morphogenesis. Cell. 1996;84:345–357.
8. Burke JM, Cao F, Irving PE, Skumatz CM. Expression of E-cadherin
by human retinal pigment epithelium: delayed expression in vitro.
Invest Ophthalmol Vis Sci. 1999;40:2963–2970.
9. Ban Y, Rizzolo LJ. A culture model of development reveals multiple
properties of RPE tight junctions. Mol Vis. 1997;3:18.
10. Grunwald GB. Cadherin cell adhesion molecules in retinal development and pathology. Prog Retinal Eye Res. 1996;15:363–392.
11. McKay BS, Irving PE, Skumatz CMB, Burke JM. Cell-cell adhesion
molecules and the development of an epithelial phenotype in
25.
26.
27.
28.
29.
30.
31.
32.
2789
cultured human pigment epithelial cells. Exp Eye Res. 1997;65:
661– 671.
Burke JM, Cao F, Irving PE. High levels of E-/P-cadherin: correlation
with decreased apical polarity of Na/K ATPase in bovine RPE cells
in situ. Invest Ophthalmol Vis Sci. 2000;41:1945–1952.
Chang CW, Ye L, Defoe DM, Caldwell RB. Serum inhibits tight
junction formation in cultured pigment epithelial cells. Invest
Ophthalmol Vis Sci. 1997;38:1082–1093.
Zech JC, Pouvreau I, Cotinet A, Goureau O, Le Varlet B, de Kozak
Y. Effect of cytokines and nitric oxide on tight junctions in cultured rat retinal pigment epithelium. Invest Ophthalmol Vis Sci.
1998;39:1600 –1608.
Jiang WG, Martin TA, Matsumoto K, Nakamura T, 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. 1999;181:319 –329.
Balkovetz DF, Pollack AL, Mostov KE. Hepatocyte growth factor
alters the polarity of Madin-Darby canine kidney cell monolayers.
J Biol Chem. 1997;272:3471–3477.
Gardner TW, Lesher T, Khin S, Vu C, Barber AJ, Brennan WA Jr.
Histamine reduces ZO-1 tight-junction protein expression in cultured retinal microvascular endothelial cells. Biochem J. 1996;320:
717–721.
Antonetti DA, Barber AJ, Khin S, Lieth E, Tarbell JM, Gardner TW.
Vascular permeability in experimental diabetes is associated with
reduced endothelial occludin content: vascular endothelial growth
factor decreases occludin in retinal endothelial cells. Penn State
Retina Research Group. Diabetes. 1998;47:1953–1959.
Matsumoto K, Nakamura T. Emerging multipotent aspects of hepatocyte growth factor. J Biochem (Tokyo). 1996;119:591– 600.
Zanegar R, Michalopoulos GK. The many faces of hepatocyte
growth factor: from hepatopoiesis to hematopoiesis. J Cell Biol.
1995;129:1177–1180.
Weidner KM, Behrens J, Vandekerckhove J, Birchmeier W. Scatter
factor: molecular characteristics and effect on the invasiveness of
epithelial cells. J Cell Biol. 1990;111:2097–2108.
Crepald T, Pollack AL, Prat M, Zborek A, Mostov K, Comoglio PM.
Targeting of the SF/HGF receptor to the basolateral domain of
polarized epithelial cells. J Cell Biol. 1994;125:313–320.
Hiscox S, Jiang WG. Hepatocyte growth factor/scatter factor disrupts epithelial tumour cell– cell adhesion: involvement of ␤-catenin. Anticancer Res. 1999;19:509 –517.
He PM, He S, Garner JA, Ryan SJ, Hinton DR. Retinal pigment
epithelial cells secrete and respond to hepatocyte growth factor.
Biochem Biophys Res Commun. 1998;249:253–257.
Lashkari K, Rahimi N, Kazlauskas A. Hepatocyte growth factor
receptor in human RPE cells: implications in proliferative vitreoretinopathy. Invest Ophthalmol Vis Sci. 1999;40:149 –156.
Mitamura Y, Takeuchi S, Matsuda A, Tagawa Y, Mizue Y, Nishihira
J. Hepatocyte growth factor levels in the vitreous of patients with
proliferative vitreoretinopathy. Am J Ophthalmol. 2000;129:678 –
680.
Casella AM, Taba KE, Kimura H, et al. Retinal pigment epithelial
cells are heterogeneous in their expression of MHC-II after stimulation with interferon-gamma. Exp Eye Res. 1996;68:423– 430.
Schote U, Seelig J. Interaction of the neuronal marker dye FM1– 43
with lipid membranes: thermodynamics and lipid ordering. Biochim Biophys Acta. 1998;9:135–146.
Kay AR, Alfonso A, Alford S, et al. Imaging synaptic activity in
intact brain and slices with FM1– 43 in C elegans, lamprey, and rat.
Neuron. 1999;24:809 – 817.
Dragsten PR, Handler JS, Blumenthal R. Fluorescent membrane
probes and the mechanism of maintenance of cellular asymmetry
in epithelia. Fed Proc. 1982;41:48 –53.
van Meer G, Simons K. The function of tight junctions in maintaining differences in lipid composition between the apical and
basolateral cell surface domains of MDCK cells. EMBO J. 1986;5:
1455–1464.
Yoo JS, Sakamoto T, Spee C, et al. cis-Hydroxyproline inhibits
proliferation, collagen synthesis, attachment, and migration of
cultured bovine retinal epithelial cells. Invest Ophthalmol Vis Sci.
1997;38:520 –528.
2790
Jin et al.
33. Jin ML, He S, Wörpel V, Ryan SJ, Hinton DR. Promotion of adhesion and migration of RPE cells to provisional extracellular matrices by TNF-␣. Invest Ophthalmol Vis Sci. 2000;41:4324 – 4332.
34. Klingler C, Kniesel U, Bamforth SD, Wolburg H, Engelhardt B,
Risau W. Disruption of epithelial tight junctions is prevented by
cyclic nucleotide-dependent protein kinase inhibitors. Histochem
Cell Biol. 2000;113:349 –361.
35. Potempa S, Ridley AJ. Activation of both MAPKinase and phosphatidylinositide 3-kinase by Ras is required for hepatocyte growth
factor/scatter factor-induced adherens junction disassembly. Mol
Biol Cell. 1998;9:2185–2200.
36. Staddon JM, Herrenknecht K, Smales C, Rubin LL. Evidence that
tyrosine phosphorylation may increase tight junction permeability.
J Cell Sci. 1995;108:609 – 619.
37. Antonetti DA, Barber AJ, Hollinger LA, Wolpert EB, Gardner TW.
Vascular endothelial growth factor induces rapid phosphorylation
of tight junction proteins occludin and zonula occluden 1: a
potential mechanism for vascular permeability in diabetic retinopathy and tumors. J Biol Chem 1999;274:23463–23467.
38. Van Itallie CM, Balda MS, Anderson JM. Epidermal growth factor
induces tyrosine phosphorylation and reorganization of the tight
junction protein ZO-1 in A431 cells. J Cell Sci. 1995;108:1735–1742.
39. Collares-Buzato CB, Jepson MA, Simmons NL, Hirst BH. Increased
tyrosine phosphorylation causes redistribution of adherens junction and tight junction proteins and perturbs paracellular barrier
function in MDCK epithelia. Eur J Cell Biol. 1998;76:85–92.
40. Muller D, Bouchard C, Rudolph B, et al. Cdk2-dependent phosphorylation of p27 facilitates its Myc-induced release from cyclin
E/cdk2 complexes. Oncogene. 1997;15:2561–2576.
41. Tsukamoto T, Nigam SK. Cell-cell dissociation upon epithelial cell
scattering requires a step mediated by the proteasome. J Biol
Chem. 1999;274:24579 –24584.
42. Hirase T, Staddon JM, Saitou M, et al. Occludin as a possible
determinant of tight junction permeability in endothelial cells.
J Cell Sci. 1997;110:1603–1613.
43. McCarthy KM, Skare IB, Stankewich MC, et al. Occludin is a
functional component of the tight junction. J Cell Sci. 1996;109:
2287–2298.
44. Chen YH, Merzdorf C, Paul DL, Goodenough DA. COOH terminus
of occludin is required for tight junction barrier function in early
xenopus embryos. J Cell Biol. 1997;138:891– 899.
IOVS, August 2002, Vol. 43, No. 8
45. Sakakibara A, Furuse M, Saitou M, Ando-Akatsuka Y, Tsukita S.
Possible involvement of phosphorylation of occludin in tight junction formation. J Cell Biol. 1997;137:1393–1401.
46. Tsukamoto T, Nigam SK. Role of tyrosine phosphorylation in the
reassembly of occludin and other tight junction proteins. Am J
Physiol. 1999;276:F737–F750.
47. Yanagihara N, Miura Y, Moriwaki M, et al. Hepatocyte growth
factor promotes epithelial morphogenesis and occludin linkage to
the cytoskeleton in cultured retinal pigment epithelial cells.
Graefes Arch Clin Exp Ophthalmol. 2001;239:619 – 627.
48. Saitou M, Fujimoto K, Doi Y, et al. Occludin-deficient embryonic
stem cells can differentiate into polarized epithelial cells bearing
tight junctions. J Cell Biol. 1998;141:397– 408.
49. Crepaldi T, Gautreau A, Comoglio PM, Louvard D, Arpin M. Ezrin
is an effector of hepatocyte growth factor-mediated migration and
morphogenesis in epithelial cells. J Cell Biol. 1997;138:423– 434.
50. Bonilha VL, Finnemann SC, Rodriguez-Boulan E. Ezrin promotes
morphogenesis of apical microvilli and basal infoldings in retinal
pigment epithelium. J Cell Biol. 1999;147:1533–1548.
51. Hiscox S, Jiang WG. Expression of E-cadherin, ␣, ␤, and ␥-catenin
in human colorectal cancer. Anticancer Res. 1997;17:1349 –1354.
52. Jiang WG, Singhrao SK, Hiscox S, et al. Regulation of desmosomal
cell adhesion in human tumour cells by polyunsaturated fatty acid.
Clin Exp Metastasis. 1997;15:593– 602.
53. Bringuier PP, Umbas R, Schaafsma HE, Karthus HF, Debruyne FM,
Schelken JA. Decreased E-cadherin immunoreactivity correlates
with poor survival in patients with bladder tumors. Cancer Res.
1993;53:3241–3245.
54. Hiscox S, Jiang WG. Association of the HGF/SF receptor, c-met,
with the cell-surface adhesion molecule, E-cadherin, and catenins
in human tumor cells. Biochem Biophys Res Commun. 1999;261:
406 – 411.
55. Robbins SG, Mixon RN, Wilson DJ, et al. Platelet-derived growth
factor ligands and receptors immunolocalized in proliferative retinal diseases. Invest Ophthalmol Vis Sci. 1994;35:3649 –3663.
56. Casaroli Marano RP, Vilaro S. The role of fibronectin, laminin,
vitronectin and their receptors on cellular adhesion in proliferative vitreoretinopathy. Invest Ophthalmol Vis Sci. 1994;35:
2791–2803.