* Cellular Biochemistry and Biophysics Program, Memorial Sloan-Kettering Cancer Center, New York 10021; and Department
of Physiology, University of California at San Francisco, San Francisco, California 94120
Occludin, the putative tight junction integral membrane protein, is an attractive candidate for a protein that forms the actual sealing element of the tight junction. To study the role of occludin in the formation of the tight junction seal, synthetic peptides (OCC1 and OCC2) corresponding to the two putative extracellular domains of occludin were assayed for their ability to alter tight junctions in Xenopus kidney epithelial cell line A6. Transepithelial electrical resistance and paracellular tracer flux measurements indicated that the second extracellular domain peptide (OCC2) reversibly disrupted the transepithelial permeability barrier at concentrations of < 5 µM. Despite the increased paracellular permeability, there were no changes in gross epithelial cell morphology as determined by scanning EM. The OCC2 peptide decreased the amount of occludin present at the tight junction, as assessed by indirect immunofluorescence, as well as decreased total cellular content of occludin, as assessed by Western blot analysis. Pulse-labeling and metabolic chase analysis suggested that this decrease in occludin level could be attributed to an increase in turnover of cellular occludin rather than a decrease in occludin synthesis. The effect on occludin was specific because other tight junction components, ZO-1, ZO-2, cingulin, and the adherens junction protein E-cadherin, were unaltered by OCC2 treatment. Therefore, the peptide corresponding to the second extracellular domain of occludin perturbs the tight junction permeability barrier in a very specific manner. The correlation between a decrease in occludin levels and the perturbation of the tight junction permeability barrier provides evidence for a role of occludin in the formation of the tight junction seal.
The tight junction, also known as zonula occludens,
is the apical-most component of the junctional complex of epithelial and endothelial cells. It is a region
where the plasma membrane of adjacent cells forms a series of contacts that appears to completely occlude the extracellular space as observed by transmission EM. These
contact points of the tight junction correspond to a network of intramembrane fibrils, when studied by freezefracture EM, which completely circumscribes the apices of
cells (9).
Two main functions have been attributed to the tight
junction. First, the tight junction seals the intercellular
space and is responsible for the separation of apical and
basolateral fluid compartments of epithelia and endothelia. Macromolecules of radii The tight junction is regulated in response to various
physiological and tissue-specific needs. For example, the
tight junction permeability to nutrients has been shown to
be increased by intestinal luminal glucose after food intake, suggesting that the regulation of tight junction permeability has a role in absorption of nutrients in the intestine (4, 6). A number of hormones have been shown to
affect the tight junction, including transforming growth factors ( While the physiological significance of the tight junction
is well recognized, the molecular component(s) involved
in the formation of a functional tight junction barrier are
not yet established. Several cytoplasmic peripheral membrane proteins (ZO-1, ZO2, cingulin, 7H6, and rab 13) (1,
2, 11, 12, 20, 32, 36, 38, 39) and one integral membrane
protein (occludin) (15) have been found to localize at the
tight junction. Occludin was shown to localize to junctional fibrils by immunogold labeling of freeze-fracture replicas of tight junctions (14). The cytoplasmic tail of occludin is necessary for its localization to cell-cell contacts,
perhaps via binding to ZO-1 and ZO-2 (16). However,
how these proteins function in the formation of the tight
junction permeability barrier is unclear.
Occludin, being the only putative integral membrane
protein so far identified, is a candidate for the formation of
the functional intercellular seal of the tight junction. The
primary amino acid sequence of chick occludin predicts
four membrane-spanning regions and two 44-amino acid
extracellular loops. Both extracellular domains of occludin
consist solely of uncharged residues with the exception of
one or two charged residues adjacent to the transmembrane regions (3). The nonpolar nature of the extracellular
domains and the conservation of their sequences between
five species (human, mouse, dog, chick, and rat-kangaroo)
suggest that the extracellular domains have important functional roles, perhaps in the formation of the actual contact
seal of the tight junction. To test this hypothesis, we made
synthetic peptides corresponding to each of the putative
extracellular domains of chick occludin and assayed for
their ability to alter tight junction barrier function.
Cell Culture, Calcium Switch Assay, and Measurement
of Transepithelial Resistance
The A6.2 subclone of the Xenopus kidney epithelial A6 cell line (27) was
grown on Transwell filters (Costar Corp., Cambridge, MA) in 89% DME
(1 g/liter glucose) supplemented with 0.74 g/liter bicarbonate, 5.95 g/liter
Hepes (pH 7.4), and 5% FCS and maintained at 28°C and 1% CO2. For
the calcium switch assay (17, 19, 25), A6 cells were allowed to grow in normal growth medium to confluency and were subsequently changed to low
calcium medium for 18 h. At the end of the low calcium (< 50 µM) incubation, A6 cells were replenished with normal calcium media, and the formation of tight junctions was monitored by the generation of transepithelial electrical resistance (TER).1 TER was measured directly in normal
growth media in Transwell wells. A short 4-µA current pulse was passed
across the cell monolayer using a pair of calomel electrodes via KCl salt
bridges, and the voltage was measured by a conventional voltmeter across
the same cell monolayer using a pair of Ag/AgCl electrodes via KCl salt bridges. TER was calculated from the measured voltage and normalized by the area of the monolayer. The background TER of blank Transwell filters was subtracted from the TER of cell monolayers.
Peptide Synthesis and Treatment of Cells
Peptides OCC1 corresponding to amino acids 81-125 of chick occludin (44 amino acids [aa] = DYGYGLGGAYGTGLGGFYGSNYYGSGLSYSYGYGGYYGGVNQRT) and OCC2 corresponding to amino acids 184-
227 of chick occludin (44 aa = GVNPQAQMSSGYYYSPLLAMCSQAYGSTYLNQYIYHYCTVDPQE) are the entire first and second putative extracellular domains of chick occludin, respectively. Two different
peptide forms of the second extracellular domain were synthesized. The
first peptide, OCC2, was modified at the two cysteine residues of the extracellular domain by covalent linkage to acetamidomethyl (underlined)
to prevent formation of disulfide bond(s). An unmodified form of the second extracellular domain peptide without the protection groups at the two
cysteine residues, OCC2(U), was also synthesized. A scrambled peptide
OCC2(S) composed of a scrambled sequence of the same residues as
OCC2, including the acetamidomethyl modification at cysteines, was also
synthesized (44 aa = GACQVYDPYMSGNYPAQSLMYQNLQYLVSGIHTYPECSYATQSY). Peptides were prepared as 10 mM stock solutions in DMSO and were added to both sides of the Transwell bathing
wells. All peptides were synthesized by the Microchemistry Core Facility
at the Memorial Sloan-Kettering Cancer Center (New York).
Paracellular Tracers Flux Assay
Flux assays were performed on 6.5-mm Transwell (in 24-well cell culture
dishes). Four different paracellular tracers, [3H]mannitol (Amersham
Corp., Arlington Heights, IL), [14C]inulin (Amersham Corp.), neutral
dextran (mol wt 3,000) conjugated with Texas red (Molecular Probes, Eugene, OR), and neutral dextran (mol wt 40,000) conjugated with Texas
red (Molecular Probes), were used. At the beginning of the flux assay,
both sides of the bathing wells of Transwell filters were replaced with
fresh media without peptides (containing 5 mM unlabeled mannitol [mol
wt 184] and 1 mM unlabeled inulin [mol wt 5,200]). Each tracer was added to a final concentration of 3.6 nmol for [3H]mannitol, 0.36 nmol for
[14C]inulin, 25 µg/100 µl for dextran (mol wt 3,000), or 50 µg/100 µl for
dextran (mol wt 40,000) to the apical bathing wells that contained 100 µl
of media. The basal bathing well had no added tracers and contained 700 µl
of the same flux assay media as in the apical compartment. All flux assays
were performed at 25°C with gentle circular agitation. Cell monolayers
were first allowed to equilibrate for 30 min after the addition of tracers.
At each of the following 15-min time intervals, the entire Transwell filter cup was removed from the basal bathing well and transferred to a fresh
basal bathing well containing 700 µl of the flux assay media. Three 15-min
flux sampling intervals were taken, and the mean was used for the calculation of paracellular tracer flux that is taken as [amount of tracer in the
basal bathing media]/[time]. For [3H]mannitol and [14C]inulin flux, the entire 700 µl of the basal media was added to 5 ml of scintillation fluid, and
3H and 14C were counted. The concentration of [3H]mannitol and [14C]inulin in the basal bathing media was calculated from a titration curve of
known concentration of the same tracers. For dextran (3 kD and 40 kD),
the concentration was calculated from the amount of fluorescence emission at 610 nm (excitation at 587 nm) using a titration curve of known concentration of the same tracers.
Immunofluorescence Staining and Western Blotting
A6 cells were grown on Transwell filters and TER was measured before
and after peptide treatment. For immunofluorescence, cells were fixed
with 100% methanol at Metabolic Labeling, Immunoprecipitations,
and Fluorography
A6 cells were grown on 75-mm Transwell filters. To examine the turnover
of occludin, each monolayer was labeled with 0.8 mCi [35S]methionine in
methionine-free media (supplemented with 5% FBS) for 22 h, and protein
turnover was monitored by replacing labeling media with fresh media in
the presence or absence of 10 mM OCC2 for 12 h. Cells were extracted for
immunoprecipitation either immediately at the end of the labeling period
(t = 0) or at 12 h after the end of the labeling period (t = 12 h). For examination of occludin synthesis, cells were rinsed once with methionine-free media and incubated in methionine-free media for 30 min before metabolic labeling. Each Transwell cell monolayer was then briefly labeled for
2 h with 1.5 mCi [35S]methionine in methionine-free media (supplemented
with 5% dialyzed FBS) in the presence or absence of 5 µM OCC2 before
extraction for immunoprecipitation. Pulse-labeling experiments were also performed on cells pretreated with 5 µM OCC2 for 20 h. For immunoprecipitation, cells from each 75-mm Transwell cell monolayer were extracted with 2 ml 1% SDS containing 5 mM EDTA and protease inhibitors (5 mM
PMSF, 5 µg/ml pepstatin A, 1 µg/ml TLCK, 10 µg/ml leupeptin, 20 µg/ml
aprotinin, 50 µg/ml antipain, 2 mM benzamidine, 50 µg/ml soybean trypsin inhibitor, and 2.5 mM iodoactamide). Each sample was boiled for 10 min before the addition of Triton X-100, deoxycholate, NaCl, and Hepes
to a final concentration of 0.2% SDS, 1% Triton X-100, 0.5% deoxycholate, 0.15 M NaCl, and 20 mM Hepes (pH 7.4). Immunoprecipitation was
performed with protein A-Sepharose (Sigma Chemical Co., St. Louis,
MO) in the presence of rabbit anti-chick occludin antibodies (see above)
or preimmune serum. Immunoprecipitates were prepared for SDS-PAGE
as described above. Polyacrylamide gels were fixed with 50% methanol
and 10% acetic acid for 1 h and incubated in Amplify (Amersham Corp.)
for 45 min before being dried under vacuum. Dried gels were exposed to
Hyperfilm-MP (Amersham Corp.) at Scanning EM
A6 cells were plated on Thermanox coverslips (Nunc, Inc., Napaville,
IL) coated with polylysine and were allowed to grow to confluency. Cells
were either untreated or treated for 40 h with 10 µM OCC1 or 10 µM
OCC2. Subsequently, the cells were rinsed twice in PBS and fixed with
2.5% glutaraldehyde/20 mM Pipes/pH 7.5 at room temperature overnight. Coverslips were rinsed in 20 mM Pipes/pH 7.5, followed by dehydration in a graded series of alcohol (50%, 75%, 95%, through absolute
alcohol) and critical point dried in a DCP-1 critical point dryer (Denton
Vacuum Inc., Cherry Hill, NJ). The samples were sputter coated with
gold/palladium in a Hummer VI sputtering system (Technic Inc., Providence, RI). The samples were photographed using a JSM 35 scanning
electron microscope (JEOL USA, Peabody, MA). All EM was performed
by Nina Lampen in the Electron Microscopy Facility at the Memorial
Sloan-Kettering Cancer Center.
Expression and Localization of Occludin
in A6 Cells Correlate with the Development of
Transepithelial Resistance
The Xenopus kidney epithelial A6 cell line formed monolayers that had a very high TER of ~8,000
TER Is Reduced by a Synthetic Peptide (OCC2)
Corresponding to the Second Extracellular Domain
of Occludin
The predicted topology of occludin from the chick occludin
amino acid sequence consists of two short extracellular domains, each one having 44 residues. Synthetic peptides
corresponding to either the entire first or second extracellular domain were made and assayed for their ability to affect tight junctions as assessed by measurements of TER.
Treatment of A6 cell monolayers with the second extracellular domain peptide (OCC2) caused a substantial reduction of TER from ~6,000
OCC2 was synthesized with a protection group, acetamidomethyl, covalently linked to the sulfhydryls of the
two cysteine residues of the peptide. When the peptide
was synthesized without the protection groups, OCC2(U)
(U stands for unmodified), no change in TER was observed at concentrations that worked maximally for the
protected peptide OCC2 (data not shown). The addition
of acetamidomethyl protection groups to the two cysteine
residues increased the hydrophilicity of OCC2 (as judged
by a mobility shift using HPLC; data not shown) and prevented intermolecular disulfide bond formation (as judged by formation of a ladder of higher molecular weight forms
using SDS-PAGE; data not shown). Either effect may
contribute to the enhanced effectiveness of the protected
peptide.
The magnitude of change in TER by treatment with
OCC2 depended on the growth state of the cells and the
dosage used. The time course and dose response of the
OCC2-induced decrease in TER were assayed on both
newly formed monolayers that were developing TER (TER ~1,000 OCC2 Increases Paracellular Permeability
The decrease in TER caused by the OCC2 peptide could
be attributed to either an increase in paracellular tight
junction permeability or transcellular plasma membrane
permeability to ions. To distinguish between the two possibilities, we assessed the flux of membrane-impermeant paracellular tracer molecules across A6 cell monolayers.
Four different paracellular flux tracers were used: mannitol (mol wt 184), inulin (mol wt 5,200), dextran 3K (mol wt
3,000), and dextran 40K (mol wt 40,000). Newly formed
A6 monolayers (starting TER of ~1,000
It was possible that OCC2 only altered the rates of
movement of these relatively small paracellular tracers
through the tight junction. To examine whether the functional tight junction barrier to macromolecules was disrupted by OCC2 peptide, the paracellular flux of dextran
40K, to which A6 cell monolayers are usually completely impermeable, was measured. Indeed, treatment of A6 cell
monolayers with OCC2 opened the paracellular barrier to
dextran 40K (Fig. 3 a). OCC1 treatment had no detectable
effect on the flux of dextran 40K. Therefore, in addition to
reducing TER, the tight junction permeability barrier to
macromolecules of 40 kD was also compromised by OCC2
treatment.
For individual Transwell monolayers, there was a close
correlation between tracer fluxes and the magnitude of the
drop in TER. For this analysis, data from the experiments
shown in Fig. 3 a was replotted for individual Transwell
monolayers in Fig. 3, b-e. This correlation held for all four
tracers: mannitol (Fig. 3 b), inulin (Fig. 3 c), dextran 3,000 (Fig. 3 d), and dextran 40,000 (Fig. 3 e). This correlation
suggested that the decrease in TER caused by OCC2 was
predominantly, if not exclusively, due to the increase in
paracellular permeability. Therefore, we conclude that the
peptide corresponding to the second extracellular domain of occludin perturbed the tight junction permeability barrier function of the A6 cell monolayers.
OCC2 Selectively Decreases the Level of Occludin
in A6 Cells
To examine the potential effects of OCC2 on the tight
junction at the molecular level, the localization and total
cellular content of various known tight junction proteins
were determined. A6 cell monolayers that were used in
paracellular tracer flux assays (see above) were immediately processed for indirect immunofluorescence microscopy. OCC2-treated monolayers (TER ~250
To confirm the depletion of occludin biochemically, we
performed Western blot analysis of various tight junction
proteins in A6 cell lysates that were treated for 24 h with
OCC1 (10 µM), OCC2 (10 µM), or DMSO solvent control
(0.1%). OCC2 selectively reduced total cellular occludin
levels but did not alter the levels of ZO-1, ZO-2, cingulin,
or E-cadherin (Fig. 5 a). Moreover, OCC1 had no effect
on the level of occludin. Total cellular occludin levels were
also unaltered by the scrambled peptide OCC2(S) and the
unmodified peptide OCC2(U) (Fig. 5 b). These results
suggested that the perturbation of the tight junction permeability barrier correlated with the selective reduction of
total cellular occludin levels but not the levels of other
junctional proteins, ZO-1, ZO-2, cingulin, and E-cadherin.
To determine whether the decreased occludin levels by
OCC2 treatment were due to an inhibition of occludin
protein synthesis or an enhancement of occludin turnover,
the effect of OCC2 on occludin synthesis and turnover was
tested. The synthesis of occludin was examined by briefly
metabolically labeling cellular proteins with [35S]methionine (2.5-h pulse) followed by immunoprecipitation of occludin. Incorporation of [35S]methionine into occludin was
not altered by either 2 or 22 h of OCC2 treatment relative
to untreated cells. Therefore, it is unlikely that OCC2 decreased total cellular occludin levels by inhibiting occludin
synthesis (Fig. 5 c).
The effect of OCC2 on occludin turnover was determined by analyzing the disappearance of [35S]methioninelabeled occludin from steady-state labeled cells (22 h). OCC2
peptide was added to the chase medium and, within 12 h
of chase, the amount of labeled occludin was noticeably
lower than that of untreated cells, suggesting that OCC2
caused occludin to turn over faster (Fig. 5 d). Therefore,
the depletion of cellular occludin by OCC2 was attributed
to an increase in occludin turnover.
TER Recovery after OCC2 Removal
If OCC2 decreased TER and occludin levels by specifically promoting occludin turnover rather than nonspecific
toxicity, A6 cells would be expected to remain healthy and
to be capable of reforming the tight junction permeability
barrier after the removal of OCC2. Therefore, we tested
the ability of A6 cell monolayers to recover TER after the
removal of OCC2 peptide. Newly formed monolayers
were treated with 5 µM OCC2 for 24 h, and TER dropped from ~2,000
Cell Morphology after OCC2 Treatment
The OCC2 peptide could act by disrupting the tight junction extensively, causing adjacent cells to separate from
each other. Alternatively, OCC2 might selectively perturb
the tight junction sealing element, leaving the overall tight
junction morphology relatively intact. Scanning EM was
used to examine the effect of OCC2 on A6 cell contacts.
Examination of gross cell morphology by scanning EM did
not reveal any detectable difference between cell monolayers that were treated for 30 h with DMSO (0.1%),
OCC1 (10 µM), or OCC2 (10 µM) (Fig. 7). All monolayers appeared intact and where characterized by a high
density of microvilli at cell boundaries. In addition, in preliminary experiments, no changes in the tight junction, as
observed by transmission EM of thin-sections, were detected (unpublished observations). However, a much
more extensive study would be necessary to determine the
effects of OCC2 at the ultrastructural level.
A synthetic peptide (OCC2) corresponding to the entire
second extracellular domain of chick occludin was able to
consistently and significantly decrease TER when added
to Xenopus kidney epithelial A6 cell monolayers. The decrease in TER was attributed to a disruption of the tight
junction permeability barrier because it was associated with
an increase in paracellular flux of membrane-impermeant
tracers. The effect of OCC2 on the tight junction permeability barrier correlated with a selective depletion of total
cellular levels and junctional localization of occludin. On
the other hand, the total cellular levels and localization of
cytoplasmic components of the tight (ZO-1, ZO-2, and
cingulin) and adherens (E-cadherens) junctions were not
affected. Furthermore, cell morphology, as observed by
scanning EM, was not altered. These results suggested that
the second extracellular domain peptide of occludin (OCC2) acted specifically to perturb the permeability barrier function of the tight junction. The correlation of the physiological effects of OCC2 with the selective reduction of occludin provides evidence for a role for occludin in the
formation of a functional tight junction seal.
The effect of OCC2 did not appear to be caused by general cell toxicity or perturbation of the plasma membrane.
First of all, the perturbation of epithelial permeability by
OCC2 occurred with only a maximal effective concentration
of 5 µM. Secondly, the slow time course of the effect of
OCC2, developing over 24 h of incubation, suggested that
the peptide did not act by perturbing plasma membrane
integrity, an effect that would be expected to be immediate. Thirdly, incubation of OCC2 for up to 5 d did not cause changes in overall epithelial cell morphology, suggesting that OCC2 was not detrimental to the cells. In fact,
if the observed increase in the flux of 40-kD dextran had
been caused by leaking through the plasma membrane, the
cells would not be expected to survive. Moreover, cells
that were treated with OCC2 excluded the vital dye, trypan blue, indicating that they remained intact and alive
(data not shown). Fourthly, the effect of OCC2 was reversible after removal of the peptide, indicating that
OCC2 did not impair the cells permanently but only temporarily perturbed the tight junction permeability barrier.
In conclusion, OCC2 did not appear to be toxic to the cells
or disruptive to the integrity of the plasma membrane in a
nonspecific manner.
The effect of OCC2 appeared to be due to specific perturbation of the tight junction permeability barrier. First
of all, the effect was specific to the amino acid sequence of
OCC2 because only OCC2, but not OCC2(S), the scrambled sequence of the same amino acid composition as
OCC2, was able to decrease the TER of cell monolayers.
Secondly, the decrease in TER caused by OCC2 was associated with an increase in the flux of membrane-impermeant tracers, demonstrating that OCC2 affected the
paracellular tight junction barrier. Thirdly, OCC2 did not
cause observable changes in epithelial cell morphology,
suggesting that the perturbation of the tight junction barrier was not an indirect effect resulting from some gross
morphological alteration of the cells. Furthermore, OCC2
did not disrupt the entire epithelial junctional complex, because neither the cellular levels nor the localization of
various known tight (ZO-1, ZO-2, and cingulin) and adherens (E-cadherin) junction proteins were altered. Therefore, the physiological effects of OCC2 appeared to be due
to specific perturbation of the permeability seal of the
tight junction.
Although OCC2 caused cell monolayers to become permeable to macromolecular tracers, including dextran 40K,
the TERs of treated monolayers remained at significant
levels (~250 OCC2 had specific reproducible effects on occludin, the
protein from which it is derived. OCC2 selectively depleted the cellular level and junctional localization of occludin without influencing the levels of other junctional
proteins examined. Control peptides that did not perturb
the tight junction permeability barrier, including a scrambled version of OCC2, OCC2(S), did not alter occludin
levels or localization. In addition, the amount of reduction in occludin levels at various time points after OCC2 treatment closely correlated with the magnitude of drop of TER
(data not shown). Moreover, after the removal of OCC2,
occludin expression and localization recovered completely
to normal levels. In fact, the time course of both the depletion and recovery of occludin correlated with the time
course of the physiological effects of OCC2. Thus, the selective depletion of occludin from the tight junction appeared to be a biochemical effect consistently associated with the physiological action of the peptide. Therefore, we
propose that the depletion of occludin by OCC2 is the
mechanism by which OCC2 perturbed the tight junction
permeability barrier.
OCC2 behaves as if it is a competitive inhibitor of occludin function that competes with endogenous occludin for
its receptor(s) or binding protein(s). Two possible mechanisms of action of OCC2 could account for its perturbation
of the tight junction barrier. First, OCC2 could bind an occludin receptor by intercalating into the tight junction and
directly interfering with the normal function of a functional seal. Secondly, OCC2, by binding to occludin receptor(s), could cause the release of occludin from its normal
stabilized interactions in the tight junction, which subsequently leads to gradual disassembly of tight junction sealing elements. Our results are consistent with the second
mechanism because the effect of OCC2 correlated with
the depletion of occludin, suggesting that disassembly of
sealing elements is the mechanism of OCC2 action.
Our results strongly implicate occludin in the formation
of the tight junction permeability barrier. Both the specificity of the OCC2 peptide sequence and the correlation of
occludin levels and TER support this contention. This conclusion is consistent with the findings that occludin is
present at the tight junction contact points and the intramembrane fibrils (14, 15), the region defined as the occluding barrier where paracellular tracers do not permeate
through. The simplest model to explain how occludin might
participate in sealing the tight junction is that occludin polymerizes in the plane of the plasma membrane and completely circumscribes the apex of cells. However, we do
not known whether occludin is the only component of the
fibrils or some other unidentified protein(s) also participate(s) in the formation of these fibrils. We also do not
know how occludin forms the cell-cell contact at the tight
junction. Does it bind to another occludin from the adjacent cell in a homophilic interaction or to an unidentified protein in a heterophilic manner? Interestingly, preliminary data indicate that OCC2 peptide-coupled beads can
form a large aggregate, which might suggest that OCC2
binds to itself, perhaps reflecting either a homophilic binding event or polymerization of occludin (Wong, V., unpublished data). Regardless of the mechanism, our findings
provide strong evidence for the functional role of occludin in the tight junction permeability barrier.
The fact that occludin can be depleted from the tight
junction without affecting other known tight junction proteins suggests that its incorporation into the tight junction
can be regulated separately. This can provide a potential
mechanism to regulate the permeability barrier function
of the tight junction without affecting the rest of the tight
junction structure. Instead of assembling and disassembling the whole tight junction complex, the tight junction
permeability barrier could be regulated by recruiting and
removing occludin to and from the tight junction. This
could also help to explain the dynamic nature of the tight junction, which must rapidly open and close during physiological processes. Further experiments will be required to
determine whether the incorporation of occludin into the
tight junction is regulated physiologically.
The OCC2 peptide offers the possibility to selectively
and transiently eliminate the tight junction permeability
barrier without disrupting the general architecture of cells.
OCC2 treatment is the most specific means identified so
far to selectively perturb the paracellular barrier of the
tight junction. The transient perturbation of the barrier
function of the tight junction by OCC2 could be potentially useful in medical therapeutics such as the facilitation
of drug delivery across the blood-brain barrier. The OCC2
sequence should be considered as just a starting point for
potential pharmaceutical development. It may be possible
to identify other occludin sequences or small molecules that
can optimally perturb occludin and the physiological properties of the tight junction.
15 Å cannot flow through the
tight junction (23). However, depending on the properties
of each individual epithelium, small ions can pass through
the tight junction to varying degrees, as indicated by differences in transepithelial electrical resistance in various
epithelial and endothelial cells (5
cm2 to > 5,000
cm2)
(13, 29). Therefore, the tight junction is crucial for the formation of blood-tissue barriers, such as the blood-brain
barrier and the blood-retinal barrier, which are absolutely
essential for the normal functioning of the organism (30,
34). Second, the tight junction functions as a diffusion barrier to plasma membrane lipids and proteins, which helps
to define apical and basolateral membrane domains of
these polarized epithelial and endothelial cells (18, 20).
Therefore, the tight junction is crucial for the epithelium
to generate chemical and electrical gradients across the
cell monolayer that is necessary for vectorial transport
processes such as absorption and secretion.
and
) (8, 35) and glucocorticoids (37). Disruption of the tight junction has been found to occur in many
diseases including hepatitis, Celiac Spruce, Crohn's disease, and gastritis, in which the tight junction intramembrane fibrils of the respective epithelia manifest discontinues and poor organizations (21, 24, 28, 33). On the other
hand, increases in tight junctional depth in intestinal epithelium in Blind Loop syndrome correlate with a decrease
in permeability to nutrients (31). In addition, the tight
junction is regulated in various physiological processes, such as leukocyte transmigration across an endothelium
(26) and intestinal cell division (5) and extrusion (7), to ensure minimal disruption of the tight junction barrier.
Therefore, the tight junction appears to function as more
than just a paracellular seal, and instead is regulated in
physiologically important processes.
Materials and Methods
20°C for 30 min and dried with 100% acetone at
20°C for 5 min. Filters were blocked with immunofluorescence staining
buffer (1% nonfat dry milk in 0.5% Triton X-100, 5 mM EDTA, 0.15 M
NaCl, and 20 mM Hepes, pH 7.0) before incubation with primary antibodies. Rabbit anti-occludin antibodies were raised against a glutathione-
S-transferase fusion protein of the cytoplasmic domain of chick occludin
(255-510 aa). Rabbit anti-ZO-1 (No. 10153) and -ZO-2 (No. 9989) (22)
antibodies were gifts from D. Goodenough (Harvard Medical School,
Cambridge, MA). Rabbit anti-cingulin antibodies were gifts from S. Citi
(University of Padua, Italy). Monoclonal mouse anti-Xenopus E-cadherin
antibodies (5D3) were raised against the extracellular domain of E-cadherin (10). FITC-conjugated secondary antibodies were obtained from Molecular Probes. For Western blot analysis, cells were rinsed twice in
PBS and extracted directly in SDS-PAGE sample buffer (50 mM TrisHPO4, pH 6.8, 2.5 mM EDTA, 15% sucrose, 2% SDS, and 50 mM DTT)
containing protease inhibitors (5 mM PMSF, 5 µg/ml pepstatin A, 1 µg/ml
N-
-p-tosyl-L-lysine-chloromethyl ketone (TLCK), 10 µg/ml leupeptin, 20 µg/ml aprotinin, 50 µg/ml antipain, 2 mM benzamidine, 50 µg/ml soybean
trypsin inhibitor, and 2.5 mM iodoactamide). Samples were boiled for 10 min, and cooled to room temperature before the addition of iodoacetic
acid to a final concentration of 125 mM, and then SDS-PAGE was performed. Western blots for occludin, ZO-1, ZO-2, cingulin, and E-cadherin were done using the same primary antibodies as those for immunofluorescence stainings. Secondary antibodies conjugated with HRP (Bio
Rad Laboratories, Hercules, CA) were developed by enhanced chemiluminescence (Amersham Corp.).
80°C.
Results
cm2 and were
impermeable to macromolecules of mol wt
40 kD. After induction of synchronized intercellular junction formation
by a calcium switch assay (see Materials and Methods for
details), occludin localization at cell boundaries correlated
with the formation of tight junctions as monitored by measurements of TER (Fig. 1 a). Western blot analysis of A6
total cell lysates using polyclonal antibodies raised against
the cytoplasmic domain of chick occludin showed a single
band of mol wt ~60 kD corresponding to Xenopus occludin. A minor band of lower molecular weight is detected
only occasionally, and therefore is presumed to be a degradation product of occludin. The expression levels of occludin during tight junction formation correlated roughly
with the increase in TER (Fig. 1 b). The increase in occludin expression plateaued as TER reached maximal steadystate levels (not shown). By Western blot analysis, ZO-1
and cingulin levels also increased during tight junction development (Fig. 1 b), but their expression levels changed
less dramatically and tended to plateau earlier than the
TER or occludin levels. The time course of occludin localization and expression was consistent with the hypothesis
that occludin participates in the formation of the tight
junction.
Fig. 1.
Expression and junctional localization of occludin correlated with the development of tight junctions in Xenopus A6
kidney epithelial cells. A6 cells were allowed to grow to confluency in Transwell filters in normal medium and were subsequently
changed to low calcium medium for 18 h. Then the medium was
replenished with normal calcium (t = 0), and the formation of tight
junctions was monitored by measuring TER at t = 0 h (TER = cm2), t = 15 h (TER ~0
cm2), t = 3 d (TER ~100
cm2),
and t = 5 d (TER > 1,000
cm2). (a) Indirect immunofluorescence of occludin in A6.2 cells at t = 0, 15 h, 3 d, and 5 d. (b)
Western blots of A6 cell lysate for occludin, cingulin, and ZO-1 at t
= 0, 15 h, 3 d, and 5 d.
[View Larger Version of this Image (29K GIF file)]
cm2 to ~900
cm2 (Fig. 2 a). In
contrast, the first extracellular domain peptide (OCC1) did
not alter TER as compared with the DMSO only control (Fig. 2 a). Additionally, a control peptide containing a
scrambled amino acid sequence from the second extracellular domain, OCC2(S), had no effect on TER (Fig. 2 a).
Therefore, the peptide corresponding to the second extracellular domain of chick occludin (OCC2) specifically reduced TER in Xenopus kidney epithelial A6 cell monolayers.
Fig. 2.
A synthetic peptide (OCC2) corresponding to the entire second extracellular domain of chick occludin decreased
TER of A6 cell monolayers. (a) Effect of various synthetic peptides on TER. OCC1 (corresponding to the entire first extracellular domain of chick occludin), OCC2 (corresponding to the entire
second extracellular domain of chick occludin), OCC2(S) (corresponding to the scrambled sequence of the entire second extracellular domain of chick occludin), and DMSO solvent control
were used. Newly confluent A6 cell monolayers (starting TER
~1,000 cm2) that were still developing TER were used. Cell
monolayers were treated with a final concentration of 5 µM
OCC1, 5 µM OCC2, 5 µM OCC2(S), or DMSO (0.05%) for 66 h,
and peptides were replenished every 24 h. At the end of the
66-h peptide incubation, TER for control monolayers reached
~5,000-6,000
cm2. n = 6 for each condition. (b) Time course of
effect of OCC2 peptide on TER of A6 cells that were still developing TER. Cell monolayers that had attained TER ~750
cm2
were treated with a final concentration of 5 µM OCC1 (n = 4) or
5 µM OCC2 (n = 5) at t = 0. Peptides were replenished at 30 h.
(c) Dose dependency of OCC2 peptide on TER in A6 cell monolayers that were still developing TER. A6.2 cells were allowed to
grow to confluency in normal medium and were subsequently
changed to low calcium medium for 18 h. The low calcium medium was replaced with normal calcium medium containing a final concentration of 0.2, 0.5, 2, and 5 µM OCC2. TER were measured after 4 d when control cell monolayers developed TER of
~3,000
cm2. n = 3 for all concentrates tested. (d) Time course
of OCC2 peptide on TER of steady-state A6 cell monolayers that
were confluent for ~2 wk (TER ~8,000
cm2). Cells were
treated with a final concentration of 5 µM OCC2 at t = 0. Untreated monolayers were done in parallel as control. Peptides
were replenished at 22 and 76 h. n = 3 for all conditions. (e) Dose
dependency of OCC2 peptide on TER in steady-state A6 monolayers (TER ~6,000
cm2). Cell monolayers were treated with a
final concentration of 0.5, 1, 2, 5, and 10 µM OCC2. TER was
measured at 40 h after peptide addition. The TER of each individual monolayer is plotted. Each concentration of OCC2 was
done on duplicate monolayers. All error bars represent standard
error.
[View Larger Version of this Image (38K GIF file)]
cm2) and steady-state monolayers that had
completely formed maximal TER (TER ~8,000-10,000
cm2). Newly formed monolayers were studied either by
plating A6 cells at confluent density or by inducing junction formation with the low calcium switch assay (see Materials and Methods); both procedures yielded the same
results. For newly formed monolayers that were still developing TER (see figure legends for details), OCC2 treatment resulted in a ~10-fold lower TER of ~250
cm2 as
compared with ~2,500
cm2 in 2 d (Figure 2 b). Treatment with the first extracellular domain peptide, OCC1,
had no effect (Fig. 2 b). The ability of OCC2 to decrease
TER in newly formed monolayers was dose dependent
with a maximal inhibition at the final concentration of 5 µM
OCC2, resulting in a ~10-fold lower TER of ~400
cm2
as compared with ~3,500
cm2 (Fig. 2 c). For steady-state
monolayers that had been confluent for >10 d and attained a maximal TER (~10,000
cm2), OCC2 decreased
TER from ~10,000
cm2 to ~2,000
cm2, whereas untreated monolayers retained a stable maximal TER (Fig. 2 d). However, the effect of OCC2 on steady-state monolayers took much longer than it did for newly formed
monolayers. It took 5 d to attain an approximately fivefold
decrease in TER in steady-state monolayers as compared
with 2 d for newly formed monolayers. The effect of
OCC2 on steady-state monolayers was also dose dependent with a maximal inhibition at 5 µM when TER decreased from ~6,000
cm2 to ~800
cm2 (Fig. 2 e). This
agreed well with the dose required for newly formed
monolayers, suggesting that OCC2 peptide acted in similar manner in both growth states. (It is important to point out
that the OCC2 peptide is somewhat water insoluble, and
therefore its actual effective concentration in solution is
unclear.) In conclusion, the second extracellular domain
peptide of chick occludin (OCC2) decreased the TER of
Xenopus A6 cell monolayers in a time- and dose-dependent manner, and the magnitude and time course of the effect depended on the growth state of the cells. Additional
experiments carried out using the calcium switch method,
in which peptides were added before TER development,
showed that OCC2 also inhibited the generation of TER
(data not shown).
cm2) were
treated with 5 µM OCC1 or OCC2 for 36 h. At the end of the 36 h of peptide treatment (when control TER developed to ~2,500
cm2), paracellular tracer flux assays were
performed. As before, treatment of monolayers with
OCC2 resulted in a ~10-fold reduction in TER from
~2,500
cm2 to ~250
cm2. In the same monolayers,
OCC2 caused a ~10-fold increase in the flux of paracellular tracers (Fig. 3 a). The flux of mannitol (hydrodynamic radius ~4 Å), inulin (hydrodynamic radius ~10-14 Å), and dextran 3K all increased ~10-fold after OCC2
treatment. Therefore, the decrease in TER caused by
OCC2 peptide was associated with an increase in paracellular permeability of the tight junction.
Fig. 3.
OCC2 increased the paracellular flux of membraneimpermeant tracer molecules. (a) Effects of OCC2 on the flux of
[3H]mannitol, [14C]inulin, Texas red-conjugated neutral dextran
(mol wt 3,000), and Texas red-conjugated neutral dextran (mol
wt 40,000). OCC1 was used as control peptide. A6 cell monolayers were allowed to grow until TER reached ~1,200 cm2. Cell
monolayers were then treated with a final concentration of 5 µl
OCC1 or OCC2 for 36 h. TER was measured and tracers flux assays were performed as described in Materials and Methods. For all four tracers, n = 8 and error bars represent SEM. (b-e) The relationship between tracer flux and TER changes induced by
OCC2 treatment. Absolute flux values for individual A6 cell
monolayers were plotted against TER of the same monolayer.
(b) [3H]mannitol, (c) [14C]inulin, (d) neutral dextran (mol wt
3,000) conjugated with Texas red, and (e) neutral dextran (mol wt
40,000) conjugated with Texas red.
[View Larger Version of this Image (34K GIF file)]
cm2) had
substantially less occludin present at cell boundaries as compared with OCC1-treated monolayers (TER ~2,500
cm2) (Fig. 4, a and b). On the other hand, ZO-1, ZO-2,
cingulin, and E-cadherin distributions were not changed
detectable by OCC2 treatment (Fig. 4, c-j). Therefore, it
appeared that OCC2 selectively depleted occludin from
the tight junction of A6 cells.
Fig. 4.
OCC2 reduced junctional stainings of occludin but not
ZO-1, cingulin, ZO-2, and E-cadherin. A6 cell monolayers from
the paracellular tracer flux assays described in Fig. 3 were processed for indirect immunofluorescence microscopy at the end of
the flux assays. OCC1-treated monolayers had TER of ~2,500
cm2, and OCC2-treated monolayers had TER of ~250
cm2.
OCC1-treated (a, c, e, g, and i) and OCC2-treated (b, d, f, h, and
j) monolayers were immunostained in parallel for occludin (a and
b), ZO-1 (c and d), cingulin (e and f), ZO-2 (g and h), and E-cadherin (i and j).
[View Larger Version of this Image (60K GIF file)]
Fig. 5.
OCC2 specifically
decreased total cellular occludin levels. (a) Western
blots of occludin, cingulin,
ZO-1, ZO-2, and E-cadherin of total cell lysates from
monolayers that were treated
with OCC1, OCC2, or
DMSO solvent control. A6
cells were allowed to grow
until TER reached ~1,000
cm2, and monolayers were
treated with 10 µM of OCC1,
10 µM OCC2, or 0.1%
DMSO for 24 h. (b) Only the
peptide that decreased TER also caused a decrease in occludin levels. Western blot of
occludin in A6 total cell lysates of monolayers that
were treated with OCC1,
OCC2(U) (unmodified),
OCC2, or OCC2(S) (scrambled). A6 cells were allowed
to grow to confluency in normal medium and were subsequently changed to low calcium medium for 18 h. A6 cells were then replenished with normal calcium media containing peptides at a final concentration of 5 µM. OCC2(U), unmodified OCC2, and OCC2(S), scrambled sequence of OCC2.
Peptides were replenished every 24 h, and cells were extracted for analysis at 4 d after initial peptide treatment. (c) Occludin synthesis
was not reduced by OCC2 treatment. A6 cells that were either untreated or treated for either 2 or 22 h with a final concentration of 5 µM OCC2 were subsequently labeled for 2.5 h with [35S]methionine followed by immunoprecipitation (IP) of occludin. (d) Turnover of
occludin was enhanced by OCC2 treatment. A6 cells were metabolically labeled 20 h with [35S]methionine. At the end of the labeling
period (t = 0), fresh media (without [35S]methionine) containing 10 µM OCC2 was added for 12 h followed by immunoprecipitation
(IP) of occludin. Untreated A6 cells were used in parallel as a control.
[View Larger Version of this Image (36K GIF file)]
cm2 to ~180
cm2. OCC2-containing medium was then removed from the cells and replaced with
fresh OCC2-free medium. After OCC2 removal, the TER
slowly increased and recovered to the initial pre-OCC2
treatment value in 48 h (Fig. 6 a). In fact, for the OCC2
peptide to maintain a continuous effect on TER, it is necessary to replenish OCC2 every 48 h because cells that
were treated with only one dose of OCC2 recovered TER
readily. Immunofluorescence analysis of occludin showed
that the recovery of TER correlated with the reappearance of occludin at the tight junction (Fig. 6 b). The reversibility of the effect of OCC2 on both TER and occludin localization suggested that OCC2 only transiently altered the
ability of A6 cells to form tight junctions. Furthermore,
the correlation of TER recovery with occludin reappearance at the tight junction again provided evidence for a
role of occludin in the formation of the tight junction permeability barrier.
Fig. 6.
The effects of OCC2 on TER and occludin accumulation were reversible. (a) Reversibility of TER after OCC2 removal. A6 cell monolayers that had TER of ~1,700 cm2 were
treated at t = 0 with a final concentration of 5 µM OCC1 or
OCC2. At t = 24 h, peptides were either replenished (OCC1 and OCC2) or removed (OCC2 Recovery) from the cells. OCC1 (n = 6), OCC2 (n = 6), and OCC2 recovery (n = 3). (b) Recovery of
junctional stainings of occludin after OCC2 removal. A6 cell
monolayers that had TER ~1,000
cm2 were treated at t = 0 with a final concentration of 5 µM OCC1 or OCC2. At t = 24 h,
peptides were either replenished (OCC1 and OCC2) or removed
(OCC2 recovery) from the cells. At t = 60 h, cells were processed
for indirect immunofluorescence microscopy of occludin. OCC1
(TER ~2,200
cm2), OCC2 (TER ~250
cm2), and OCC2 recovery (TER ~2,300
cm2).
[View Larger Version of this Image (32K GIF file)]
Fig. 7.
OCC2 did not cause morphological changes in A6 cell monolayers as observed by scanning EM. Confluent A6 cells grown on
polylysine-coated coverslips were treated 24 h with a final concentration of 10 µM OCC1 (a and d), 10 µM OCC2 (b and e), and 0.1%
DMSO (c and f). Cells were then processed for scanning EM. A6 cells were ~7-10 µM in diam.
[View Larger Version of this Image (176K GIF file)]
Discussion
cm2). This suggests that the effect of OCC2
on cells within the monolayer was not homogeneous. It is
expected that regions of the monolayer responsible for
high tracer fluxes would also be areas of low electrical resistance. Consistent with this explanation are the nonhomogeneous patterns of residual occludin staining after
OCC2 treatment (Fig. 4, a and b). Therefore, it is likely
that a fraction of the cells in the monolayers were partially
resistant to perturbation of the permeability barrier by
OCC2 treatment.
Received for publication 30 July 1996 and in revised form 19 September 1996.
This work was partially supported by Cancer Center Support grant NCI-P30-CA-08748 and American Heart Association grant-in-aid 92006540. V. Wong was partially supported by National Institutes of Health predoctoral grants, NRSA-DK07265-130031 and NRSA-DK07265-130031, and a graduate opportunity fellowship from the University of California at San Francisco.We thank members of the Gumbiner laboratory for reading the manuscript and for helpful discussions. We also thank Scott Geramanos for excellent technical work on the peptide chemistry.
aa, amino acid; TER, transepithelial electrical resistance.