Ethanol modulation of intestinal epithelial tight junction
barrier
Thomas Y.
Ma,
Don
Nguyen,
Vuong
Bui,
Hanh
Nguyen, and
Neil
Hoa
Division of Gastroenterology, Department of Medicine, Department of
Veterans Affairs Medical Center, Long Beach 90822; and
University of California Irvine, Irvine, California 92717
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ABSTRACT |
Previous studies
have shown that high concentrations of ethanol (
40%) cause
functional damage of the gastrointestinal epithelial barrier by direct
cytotoxic effect on the epithelial cells. The effects of lower
noncytotoxic doses of ethanol on epithelial barrier function are
unknown. A major function of gastrointestinal epithelial cells is to
provide a barrier against the hostile substances in the
gastrointestinal lumen. The apicolaterally located tight junctions (TJs) form a paracellular seal between the lateral membranes of adjacent cells and act as a paracellular barrier. In this study, we
investigated the effects of lower doses of ethanol on intestinal epithelial TJ barrier function using filter-grown Caco-2 intestinal epithelial monolayers. The Caco-2 TJ barrier function was assessed by
measuring epithelial resistance or paracellular permeability of the
filter-grown monolayers. Ethanol (0, 1, 2.5, 5, 7.5, and 10%) produced
a dose-related drop in Caco-2 epithelial resistance and increase in
paracellular permeability. Ethanol also produced a progressive
disruption of TJ protein (ZO-1) with separation of ZO-1 proteins from
the cellular junctions and formation of large gaps between the adjacent
cells. Ethanol, at the doses used (
10%), did not cause cytotoxicity
(lactate dehydrogenase release) to the Caco-2 cells. Ethanol produced a
disassembly and displacement of perijunctional actin and myosin
filaments from the perijunctional areas. On ethanol removal, actin and
myosin filaments rapidly reassembled at the cellular borders. Ethanol
stimulated the Caco-2 myosin light chain kinase (MLCK) activity but did
not affect the MLCK protein levels. Specific MLCK inhibitor ML-7
inhibited both ethanol increases in MLCK activity and TJ permeability
without affecting the MLCK protein levels. Consistent with these
findings, metabolic inhibitors sodium azide and 2,4-dinitrophenol
significantly prevented ethanol-induced increase in Caco-2 TJ
permeability, whereas cycloheximide or actinomycin D had no effect. The
results of this study indicate that ethanol at low noncytotoxic doses causes a functional and structural opening of the Caco-2 intestinal epithelial TJ barrier by activating MLCK.
paracellular permeability; myosin light chain kinase; actin
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INTRODUCTION |
ETHYL ALCOHOL CAUSES morphological and functional
damage of the gastrointestinal mucosal surface. The role of ethanol as
a causative agent of hemorrhagic gastritis and enteritis has been well
established (2, 15, 36, 38). It has been demonstrated by various
investigators that oral ingestion or direct endoscopic spraying of high
concentrations of ethanol (40-60%) causes extensive mucosal
injury (2, 15, 36, 38, 39). Shortly after ethanol exposure, rupture and
exfoliation of surface epithelial cells occur (38, 39). This is
followed by rupture of the mucosal microvessels and subsequent
intramucosal hemorrhage, platelet aggregation, and fibrin deposition.
Consistent with these findings, Szabo et al. (34, 35) demonstrated that
gastric instillation of high concentrations of ethanol produces a rapid
vascular leakage of Evans blue dye, indicating an increase in vascular
permeability. It has been proposed that, on mucosal penetration,
ethanol produces leakage of intravascular fluid resulting in
interstitial edema, vascular stasis, vascular thrombosis, and more
extensive vascular and mucosal injury (35).
A major function of gastrointestinal epithelial cells is to provide a
physical barrier between the hostile gastrointestinal lumen and the
subepithelial tissue. The apicolaterally located tight junctions (TJs)
form a paracellular seal or barrier between the lateral membranes of
the adjacent cells and act as structural barrier against the
paracellular penetration of water-soluble molecules (1, 21). The
disruption of the TJ barrier allows an increase in epithelial
penetration by the hydrophilic substances present in the gastric and
the intestinal lumen (11, 17, 22). The leaky TJ barrier allows
paracellular penetration of toxic luminal substances, which promote
gastrointestinal mucosal injury and inflammation (11, 20, 22).
At high doses (
40%), ethanol causes cell death with rupture and
exfoliation of the surface epithelial cells, resulting in large open
wounds in the gastrointestinal mucosal surface (36, 38). The open
breach in the epithelial surface allows influx of the toxic luminal
contents into the mucosal tissue, further exacerbating the mucosal
injury. Whereas it has been demonstrated that high doses of ethanol
cause mucosal damage by direct cytotoxicity, the effects of low
noncytotoxic doses of ethanol on gastrointestinal TJ barrier are
unknown. In the present study, we examined the effects of lower doses
of ethanol on intestinal epithelial TJ barrier using the filter-grown
Caco-2 intestinal epithelial monolayer. The Caco-2 cells, initially
derived from a human colon carcinoma (30) when confluent and allowed to
mature on permeable inserts, form TJs and attain many of the
morphological and functional characteristics of small intestinal
enterocytes that make them suitable for use as a model to study small
intestinal epithelial barrier function (9, 10, 17, 20, 30, 31). The
results of this study demonstrate that ethanol at low noncytotoxic
doses (
10%) causes functional opening of the Caco-2 intestinal
epithelial TJ barrier. Additionally, some of the intracellular
processes involved in the ethanol opening of the intestinal epithelial
TJ barrier is elucidated.
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MATERIALS AND METHODS |
DMEM, trypsin, and fetal bovine serum were purchased from Life
Technologies (Gaithersburg, MD). Glutamine, penicillin, streptomycin, and PBS solution were purchased from Irvine Scientific (Santa Ana, CA).
Colchicine, cytochalasin B, and cycloheximide were purchased from Sigma
Chemical (Indianapolis, IN). Millicell-HA 0.4-µm permeable filters
(12 mm) were purchased from Millipore (Bedford, MA). Anti-ZO-1 antibody
and FITC-streptavidin were obtained from Zymed Laboratories (San
Francisco, CA), and fluorescein-conjugated rabbit anti-rat antibodies
were obtained from Boehringer Mannheim (Indianapolis, IN).
[14C]mannitol was
obtained from NEN Research Products (Wilmington, DE). All other
chemicals were of reagent grade.
Cell cultures.
Caco-2 cells were purchased from American Type Culture Collection
(Rockville, MD). The stock cultures were grown in a culture medium
composed of DMEM with 4.5 mg/ml glucose, 50 U/ml penicillin, 50 U/ml
streptomycin, 4 mmol/l glutamine, and 10% fetal bovine serum (10, 30).
Culture medium was changed every 2 days. The cells were subcultured by
partial digestion with 0.25% trypsin and 0.9 mmol/l EDTA in
Ca2+- and
Mg2+-free PBS solution. Caco-2
cells were detached from stock cultures by trypsin digestion, washed
once by centrifugation, resuspended, and subcultured in 14 ml medium in
culture flasks at a concentration of 1 × 105 cells/ml. Cultures were
examined on a regular basis under an inverted light microscope to
monitor growth and contamination. For growth on filters, high-density
Caco-2 cells (5 × 105 cells)
were plated on nitrocellulose-based Millicell-HA filters and monitored
regularly by measuring epithelial resistance.
Determination of epithelial monolayer resistance and paracellular
permeability.
The electrical resistance of the filter-grown intestinal monolayers was
measured with an epithelial voltohmmeter (World Precision Instruments,
Sarasota, FL) as previously reported (18, 27). For resistance
measurements, both apical and basolateral sides of the epithelium were
bathed with the same buffer solution. Electrical resistance was
measured until similar values were recorded on three consecutive
measurements. The resistances of monolayers in this study are reported
after subtraction of the resistance value of the filters alone. The
effect of ethanol on Caco-2 monolayer paracellular permeability was
examined with the established paracellular marker mannitol (23, 24).
For determination of mucosal-to-serosal flux rates of the paracellular
probe mannitol, only Caco-2-plated filters having epithelial resistance
of 400-450
· cm2 were
used. The filter-grown Caco-2 monolayers reached epithelial resistance
of 400-450
· cm2 by
3-4 wk after plating (10, 20). Unless specified otherwise, Krebs-PBS (pH 7.4) was used as the incubation solution during the
experiments. Buffered solution (300 µl) was added to the apical compartment, and 450 µl were added to the basolateral compartment to
ensure equal hydrostatic pressure as recommended by the manufacturer. Known concentrations of mannitol (10 µmol/l) and its radioactive tracer ([14C]mannitol)
were added to the apical solution. Low concentrations of mannitol were
used to ensure that negligible osmotic or concentration gradient was
introduced. Test reagent was added to both the apical and basolateral
compartments as indicated. All flux studies were carried out at
37°C. All of the experiments were repeated four to six times to
ensure reproducibility.
Fluorescent labeling of cytoskeletal elements and TJ proteins.
Distribution of actin microfilaments was assessed using fluorescent
labeling techniques as previously described (20).
Monolayers grown on coverslips were fixed in 3.75% formaldehyde
solution in PBS for 20 min at room temperature and were permeabilized
in acetone at
20°C for 5 min and washed with 1 M PBS
solution. Then, 10 U of fluorescein-labeled phalloidin (Molecular
Probes, Eugene, OR) dissolved in 200 µl of PBS was placed on the
coverslips for 40 min. After PBS rinse, coverslips were mounted on a
slide with the cell side down in a 1:1 solution of PBS and glycerol.
The Caco-2 myosin filaments were labeled with anti-myosin antibody.
After fixation with 2.0% formaldehyde and permeabilization in acetone
as previously described, Caco-2 monolayers were labeled with 1:10
diluted anti-myosin antibody (Amersham, Arlington Heights, IL) in PBS.
This was followed by incubation with 1:10 fluorescein-conjugated rabbit
anti-mouse antibody (Amersham) in PBS. Coverslips were mounted in 60%
glycerol-PBS, 0.4% n-propyl gallate.
The tight junctional protein ZO-1 was labeled with anti-ZO-1 antibody
(26, 27). Epithelial monolayers grown on coverslips were fixed with
2.0% formaldehyde and permeabilized in acetone as described
previously. The Caco-2 monolayers were labeled with anti-ZO-1 antibody
diluted 1:20 with Tris-buffered saline solution; this was followed by
incubation with 1:30 diluted Tris-buffered saline solution containing
secondary anti-rabbit IgG biotinylated antibody (Zymed Laboratories)
and incubation with 1:20 diluted Tris-buffered saline solution
containing FITC-streptavidin (Zymed Laboratories). Coverslips were
mounted in 60% glycerol-Tris-buffered saline solution, 0.4%
n-propyl gallate, and viewed on a
Nikon epifluorescence microscope. All of the fluorescent labeling
experiments were repeated three to five times to ensure reproducibility.
In vitro myosin light chain kinase activity determination.
Caco-2 myosin light chain kinase (MLCK) activity was determined by
direct in vitro kinetic measurement of MLCK activity of the
immunoprecipitated MLCK. For MLCK activity studies, Caco-2 monolayers
were serum deprived overnight. After appropriate experimental treatment, cells were immediately rinsed with ice-cold Hanks' balanced
salt solution. Cells were then lysed using 0.8 ml lysis buffer (50 mM
HEPES, 100 mM NaCl, 2 mM EDTA, 1 µM pepstatin, 1 µg/ml leupeptin, 2 mM phenylmethysulfonyl fluoride, 2 mM sodium vanadate, 2 µg/ml
aprotinin, 40 mM para-nitrophenol phosphate di-cyclohexylammonium salt) and scraped, and lysates were placed in
Microfuge tubes (tube A) and
microcentrifuged 5 min to yield a clear lysate.
Anti-MLCK antibody (5 µl/200 µl lysis buffer) was added to a
separate Microfuge tube (tube B)
containing the protein A beads and incubated end-over-end for 1 h at
4°C. Then 100 µl of each cleared lysate (tube
A) were added to the microvial (tube
B) containing the pelleted protein A bead-MLCK
antibody complex and incubated end-over-end for 2 h at 4°C. The
microvial containing the immune mixture was microfuged, and the
supernatant was aspirated and washed sequentially with lysis buffer and
solution of 10 mM HEPES and 10 mM magnesium acetate at 4°C.
Protein A bead-MLCK antibody immunoprecipitated MLCK (immune mixture)
was then used in an in vitro kinase reaction in microfuge tubes to
determine the phosphorylation of MLC by the isolated MLCK. For this, 20 µl myosin light chain protein (2 mg/ml), 20 µl of 3 times hot mix
{150 µM ATP, 10 µl
[32P]ATP (5 µCi/reaction), 30 mM magnesium acetate, 30 mM HEPES} were
added, mixed with the immune mixture, and incubated for 30 min at
30°C. The MLCK-catalyzed phosphorylation reaction was terminated by
addition of 20 µl stop buffer solution (1 ml 2 M Tris buffer, pH 6.8, 2 ml 20% SDS, 4 ml glycerol, 3 ml water, 308 mg dithiothreitol, and
trace of bromphenol blue). Subsequently, the reaction mixture was
boiled for 3 min and microcentrifuged for 10 s, and then the supernatant (40-50 µl) was separated on 10% SDS-PAGE. The gel was fixed in 40% MeOH and 10% acetic acid overnight and stained with
Coomassie blue solution, dried, and then autoradiographed.
 |
RESULTS |
Ethanol modulation of Caco-2 TJ barrier.
The effect of ethanol on Caco-2 intestinal epithelial TJ barrier
function or TJ permeability was determined by measuring Caco-2 epithelial resistance and/or epithelial permeability to paracellular marker mannitol (23, 24). Addition of varying doses (0, 1, 3, 5, 7.5, 10%) of ethanol produced a dose-related drop in Caco-2 epithelial
resistance over the 60-min treatment period (Fig.
1). As shown in Fig.
2A,
ethanol (7.5%) produced a progressive decrease in Caco-2 epithelial
resistance over the 60-min treatment period. Ethanol also caused a
progressive increase in Caco-2 epithelial permeability to paracellular
marker mannitol over the 60-min period (Fig.
2B). Previous studies (20, 24) have
demonstrated an inverse relationship between intestinal epithelial
resistance and paracellular permeability. To verify such a
relationship, we compared ethanol-induced alteration in Caco-2
epithelial resistance and paracellular permeability (Fig.
2C). There was a linear relationship (r = 0.97) between ethanol-induced
decrease in Caco-2 epithelial resistance and increase in epithelial
permeability to mannitol, confirming an inverse relationship between
Caco-2 epithelial paracellular permeability and epithelial resistance.
[As in other studies (8, 20, 23, 25, 39), increase in
paracellular permeability or drop in epithelial resistance was used as
an indication of increase in TJ permeability.]

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Fig. 1.
Effect of varying concentrations of ethanol on Caco-2 epithelial
resistance. Filter-grown Caco-2 monolayers were incubated with varying
concentrations of ethanol for 60 min.
A: time course of ethanol effect on
Caco-2 epithelial resistance. B: dose
effect of ethanol on Caco-2 epithelial resistance after 60 min of
treatment. Data represent means ± SE of epithelial
resistance; n = 4.
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Fig. 2.
Comparison of ethanol modulation of Caco-2 epithelial resistance and
paracellular permeability. A: ethanol
(7.5%) modulation of Caco-2 epithelial resistance (means ± SE;
n = 4).
B: ethanol (7.5%) modulation of
Caco-2 epithelial permeability to paracellular marker mannitol (means ± SE; n = 4).
C: comparison of ethanol effect on
epithelial resistance vs. paracellular permeability
(r = 0.97).
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Next, we examined whether ethanol alteration of Caco-2 TJ barrier
function was reversible. After a 60-min treatment, ethanol was removed
and Caco-2 epithelial monolayers were incubated in Krebs buffer
solution for 3 h. Similar to above, ethanol treatment resulted in an
increase in Caco-2 paracellular permeability and a decrease in
epithelial resistance. After ethanol removal, Caco-2 epithelial
resistance (Fig.
3A) and
paracellular permeability (Fig. 3B)
rapidly returned to the baseline levels, indicating retightening of the
TJ barrier. These findings suggested that ethanol-induced increase in
TJ permeability resulted from a reversible functional change in the TJ
barrier rather than from a permanent cell injury or cell death. The
possible cytotoxic effect of ethanol on Caco-2 monolayers was also
determined by measuring lactate dehydrogenase (LDH) release using the
LDH assay kit from Sigma Chemical. Ethanol, at the doses used (
10%),
did not result in a significant increase in LDH release by the Caco-2
cells (Fig. 4). There was a significant
increase in LDH release at ethanol concentrations >12%. To examine
the possible "late LDH release" by the ethanol-treated cells, LDH
release was also measured for an additional 24 h after ethanol
treatment. There was no significant increase in late LDH release by the
ethanol (
10%)-treated cells compared with the control or untreated
cells (data not shown).

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Fig. 3.
Reversibility of ethanol modulation of Caco-2 epithelial resistance and
paracellular permeability. Filter-grown Caco-2 monolayers were treated
with ethanol (7.5%) for 60 min; subsequently, ethanol was removed and
Caco-2 monolayers were incubated with Krebs-PBS solution for 3 h.
Effect of ethanol removal on Caco-2 epithelial resistance
(A) and Caco-2 permeability to
mannitol (B)
(n = 4).
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Fig. 4.
Cytotoxicity of ethanol as measured by lactate dehydrogenase (LDH)
release. Filter-grown Caco-2 monolayers were treated with various
concentrations of ethanol for 60 min; subsequently, incubation solution
was collected and LDH release was measured. Values are means ± SE;
n = 4. Total LDH release (determined
by treatment with 1 times Triton X-100) was 1,438 U/l. Experiments were
repeated in quadruplicate to ensure reproducibility.
* P < 0.001.
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Ethanol modulation of Caco-2 ZO-1 tight junctional proteins.
In the following studies, the modulatory effect of ethanol on Caco-2 TJ
was examined by immunofluorescent labeling of tight junctional proteins
ZO-1. In the control monolayers, ZO-1 proteins were present in the
cellular periphery and appeared as a continuous band localized at the
intercellular borders (Fig.
5A).
Ethanol treatment produced a progressive disruption of ZO-1 proteins
and displacement of ZO-1 proteins away from the cellular borders with formation of large paracellular openings between the adjacent cells
(Fig. 5B). On ethanol removal, ZO-1
proteins rapidly reassembled at the apical cellular borders with
reclosure of the paracellular gaps (Fig.
5D). These findings visually
correlated with ethanol-induced functional "opening" (increased
in TJ permeability) and subsequent reclosure of the TJ barrier,
confirming a structural-functional relationship.

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Fig. 5.
Effect of ethanol on immunofluorescent localization of Caco-2
ZO-1 proteins (en face views). Caco-2 ZO-1 proteins were labeled with
immunofluorescent antibody labeling technique as described in
MATERIALS AND METHODS.
A: untreated or control monolayer.
B: Caco-2 cells treated with ethanol
for 60 min. C: Caco-2 cells
pretreated with sodium azide (30 mM) and then treated with ethanol for
60 min. D: Caco-2 cells treated with
ethanol for 60 min, followed by removal of ethanol and incubation in
cell culture medium for 2 h (original magnification ×200).
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Ethanol perturbation of Caco-2 actin microfilaments.
Previous studies have suggested a central role for perijunctional actin
and myosin filaments in modulation of intestinal epithelial TJs (20,
22, 23). In the following studies, the possible involvement of actin
microfilaments in ethanol modulation of Caco-2 TJ barrier was examined.
In the mature Caco-2 monolayers, F-actin filaments were present as
brightly staining structures localized mainly at the perijunctional
area (Fig.
6A) as
visualized by immunofluorescent labeling. Ethanol treatment resulted in
breakage of the F-actin filaments with clumping and displacement of the
perijunctional actin filaments from the cellular borders (Fig.
6B). This was associated with a
pulling apart of the adjacent cells and formation of large paracellular
gaps. On removal of ethanol (Fig.
6D), actin filaments rapidly
reassembled at the perijunctional location correlating with the
functional changes in Caco-2 TJ barrier. (It should be noted that ZO-1
and F-actin labeling studies were also performed on the Caco-2 cells
grown on permeable inserts with similar results, suggesting that
ethanol-induced morphological changes in these structures were similar
between the two systems.)

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Fig. 6.
Effect of ethanol on fluorescent localization of F-actin filaments (en
face views). Caco-2 F-actin filaments were labeled with
fluorescein-conjugated phalloidin as described in
MATERIALS AND METHODS.
A: untreated or control monolayer.
B: Caco-2 monolayers treated with
ethanol for 60 min. C: Caco-2 cells
pretreated with sodium azide and then treated with ethanol for 60 min.
D: Caco-2 cells treated with ethanol
for 60 min, followed by removal of ethanol and incubation in cell
culture medium for 2 h (original magnification ×200).
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Ethanol modulation of Caco-2 myosin filaments and MLCK.
In small intestinal epithelial cells, a peripheral band of F-actin and
myosin filaments encircle the cells near the apical junctions. As shown
in Figs. 6 and 7, perijunctional belt of
actin and myosin filaments also encircle the Caco-2 cells at the apical borders (en face views). In the following experiments, the effect of
ethanol on perijunctional myosin filaments was determined. In the
Caco-2 intestinal monolayers, myosin filaments were present near the
apical junctions forming a continuous peripheral belt encircling the
cells (Fig. 7A). Ethanol treatment
produced a disassembly and displacement of perijunctional myosin
filaments internally with visible separation of myosin filaments from
the adjacent cells (Fig. 7B). On
removal of ethanol, myosin filaments rapidly reassembled at the
cellular borders (Fig. 7D),
correlating with the functional changes in Caco-2 TJ barrier.

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Fig. 7.
Effect of ethanol on immunofluorescent localization of Caco-2 myosin
filaments (en face views). Caco-2 myosin filaments were labeled using
immunofluorescent antibody labeling technique as described in
MATERIALS AND METHODS.
A: untreated or control monolayer.
B: Caco-2 cells treated with ethanol
for 60 min. C: Caco-2 cells
pretreated with sodium azide and then treated with ethanol for 60 min. D: Caco-2 cells treated with
ethanol for 60 min, followed by removal of ethanol and incubation in
cell culture medium for 2 h (original magnification ×200).
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The MLCK plays an integral role in activating contraction of actin and
myosin filaments (13, 14). In the following studies, the possible role
of MLCK in ethanol modulation of Caco-2 TJ barrier was examined. The
modulatory effect of ethanol on Caco-2 MLCK activity was determined by
direct in vitro kinetic measurements of MLC phosphorylation by the
immunoprecipitated Caco-2 MLCK. The Caco-2 MLCK immunoprecipitated from
ethanol-treated cells greatly increased MLC phosphorylation compared
with the untreated cells (control), indicating ethanol activation of
Caco-2 MLCK. In contrast, ethanol did not effect Caco-2 MLCK protein
level (Fig. 8). Specific MLCK inhibitor
1-(5-iodonaphthalene-1-sulfonyl)-1H-hexahydro-1,4-diazepine (ML-7)
inhibited the ethanol-induced increase in MLCK activity without
affecting the MLCK protein levels. ML-7 also inhibited the
ethanol-induced drop in Caco-2 epithelial resistance (Fig. 9), suggesting that MLCK activation was
necessary for the increase in Caco-2 TJ permeability.

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Fig. 8.
Immunoblot analysis of ethanol effect on Caco-2 myosin light chain
kinase (MLCK) activity. A: Western
blot analysis of Caco-2 MLCK following treatment with either regular
medium, ethanol (7.5%), ethanol (7.5%) and ML-7 (10 µM), or ML-7
(10 µM) alone for 10 min. B:
immunoblot of phosphorylated MLC following in vitro phosphorylation by
immunopreciptated Caco-2 MLCK. Phosphorylated MLC was separated by
SDS-PAGE, stained with Coomassie blue solution, and autoradiographed as
described in MATERIALS AND METHODS.
C: densitometry measurements of
phosphorylated MLC bands expressed in pixels
(n = 4).
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Fig. 9.
Effect of MLCK inhibitor ML-7 on ethanol modulation of Caco-2
epithelial resistance. Caco-2 monolayers were pretreated with ML-7 (10 µM) for 10 min before ethanol treatment for 60 min. ML-7
significantly inhibited (P < 0.001)
ethanol-induced drop (control) in Caco-2 epithelial resistance
(n = 4).
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Role of metabolic energy and protein synthesis on ethanol modulation
of Caco-2 permeability and perijunctional structures.
In smooth muscle cells, the contraction of actin-myosin filaments
requires metabolic energy generated by MLCK-induced
activation of myosin-Mg2+-ATPase
(26). In the following studies, the possible roles of protein
synthesis, transcription, and metabolic energy in ethanol-induced increase in Caco-2 TJ permeability were examined. Metabolic inhibitors sodium azide (Fig.
10A)
and 2,4-dinitrophenol (Fig. 10B)
prevented the ethanol-induced drop in Caco-2 epithelial resistance. The pretreatment of Caco-2 monolayers with cycloheximide (70 µM) and actinomycin D (1 µg/ml) at the doses previously shown to inhibit Caco-2 protein synthesis and transcription did not affect the ethanol-induced drop in epithelial resistance (Fig. 10,
C and
D). These findings suggest that the
ethanol-induced increase in Caco-2 TJ permeability was dependent on
metabolic energy but not new protein synthesis.

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Fig. 10.
Effect of inhibitors of metabolic energy, protein synthesis, and
transcription on ethanol modulation of Caco-2 epithelial resistance.
Caco-2 monolayers were pretreated with metabolic inhibitors sodium
azide (A, 30 mM) and 2,4-dinitrophenol
(B, 1.0 mM); results for protein
synthesis inhibitor cycloheximide (C, 70 µM), and
transcription inhibitor actinomycin D
(D) for 10 min before and during
treatment with ethanol for 60 min are also shown. Values represent
Caco-2 epithelial resistance (means ± SE;
n = 4). , Ethanol-treated cells
(control). , Cells treated with selected agents before and during
ethanol treatment. Sodium azide (A)
and 2,4-dinitrophenol (B)
significantly inhibited ethanol-induced drop in epithelial resistance
(P < 0.001).
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To further examine the structural-functional relationship, the effect
of sodium azide, cycloheximide, and actinomycin D on ethanol modulation
of perijuctional structures was determined. Consistent with the above
findings, cycloheximide and actinomycin D did not have a significant
effect on ethanol modulation of actin or myosin filaments (data not
shown). In contrast, sodium azide and 2,4-dinitrophenol almost
completely inhibited ethanol-disruption of actin and myosin filaments
(Figs. 6C and
7C). Cycloheximide and actinomycin D
also did not have a significant effect on ethanol modulation of ZO-1
proteins, whereas sodium azide and 2,4-dinitrophenol prevented ethanol
disruption of ZO-1 proteins and formation of paracellular
openings (Fig. 5C). These findings
confirm the requirement of metabolic energy in the ethanol modulation
of actin and myosin filaments and the ZO-1 tight junctional proteins.
 |
DISCUSSION |
An important function of gastrointestinal epithelia is to provide a
barrier against the mucosal penetration of toxic substances and
antigens such as H+, bile acids,
proteolytic enzymes (e.g., pepsin, trypsin, and chymotrypsin), bacteria
and bacterial by-products, and food additives present in the gastric
and the intestinal lumen. The gastrointestinal epithelial TJs act as a
structural barrier against the paracellular permeation of luminal
compounds (1, 21). The disruption of the TJ barrier allows increased
epithelial penetration of normally excluded luminal substances that may
promote mucosal injury.
The results of the present study indicate that ethanol at low
noncytotoxic doses produces a functional and morphological opening of
the Caco-2 intestinal epithelial TJ barrier. The ethanol disruption of
the Caco-2 tight junctional proteins (ZO-1) and the increase in the TJ
permeability were accompanied by structural disturbance and
displacement of perijunctional actin and myosin filaments. The
functional retightening or reclosure of the Caco-2 TJ barrier following
ethanol removal paralleled the structural reassembly of perijunctional
actin and myosin filaments and the TJ proteins, as well as the
morphological reclosure of the paracellular gaps, demonstrating a
structural-functional relationship. Additionally, our data suggest that
ethanol increase in Caco-2 TJ permeability was mediated by the
activation of Caco-2 MLCK, so that the ethanol increase in TJ
permeability correlated with an increase in MLCK activity and
inhibition of MLCK activity with prevention of ethanol-induced increase
in TJ permeability. Consistent with these findings, inhibition of
metabolic energy (which is required for MLCK activation of actin-myosin
contraction) prevented both ethanol increases in TJ permeability and
the alteration of the perijunctional structures.
Ethanol produces many morphological and functional disturbances of the
gastrointestinal epithelium (2, 12, 15, 36, 38, 40). In previous
studies, it was demonstrated that high doses of ethanol (
40%) caused
a direct cytotoxic injury of the epithelial cells at the
gastrointestinal surface with a resultant rupture and exfoliation of
the epithelial layers and formation of large open wounds in the
epithelial surface (2, 15, 36, 38, 39). The effect of lower
noncytotoxic doses of ethanol on gastrointestinal epithelial barrier is
not well understood. The small intestinal luminal levels of ethanol
routinely reach concentrations of 2-10% following moderate
consumption of ethanol (50 ml ethanol in 20% solution) (3, 28). It had
been demonstrated that the peak serum levels of ethanol were
100-150 times higher than the peak small intestinal levels
following moderate ethanol consumption, suggesting that the peak serum
ethanol levels of 100 mg/dl correspond to a small intestinal level of
10-15% (28). This is the first study to demonstrate that ethanol
at lower clinically achievable doses produces a functionally reversible
opening of the intestinal epithelial TJ barrier. The low doses of
ethanol (
10%) used in this study did not cause permanent cell damage or cell death. Thus ethanol-induced increase in TJ permeability was due
to a reversible change in the TJ barrier and not from cell death or
formation of large open wounds in the epithelial surface as seen with
the higher doses of ethanol (2, 15, 36, 38, 39).
The intestinal epithelial TJs are the apical-most structures, which
encircle the cells at the lateral borders in a beltlike manner. The TJs
make homotypic contact across the intercellular space between the
adjacent cell (1). The lateral contacts, which may be visualized by
electron microscopy and freeze-fracture analysis, act as structural
barrier against the paracellular permeation of luminal substances (1,
21, 24). An apicolateral ring or belt of actin and myosin filaments
also encircles the intestinal epithelial cells near the TJs (22). The
proximity of the "perijunctional actomyosin ring" to the apical
TJs suggested a possible interdependent relationship (22). Indeed, a
correlation between disturbance of perijunctional actin and myosin
filaments and an increase in TJ permeability have been previously
demonstrated (20, 22, 23). The treatment of intestinal epithelial cells
with actin-depolymerizing agents (cytochalasins) caused a condensation
and disruption of perijunctional actin microfilaments and structural
and functional opening of the intercellular TJ complexes (20, 23). Some
studies have also shown that an increase in intestinal epithelial TJ
permeability is associated with an increase in MLC phosphorylation (7,
8, 25, 29, 32). Based on these findings, it was suggested that MLC
phosphorylation might be an important step in the propagation of a
series of intracellular processes resulting in the opening of the TJ
barrier (29, 39). It had been hypothesized that MLC phosphorylation
induces contraction of perijunctional actin and myosin filaments, which
in turn produces tension on the apical surface and the TJs with
subsequent opening of the TJs between the adjacent cells (29, 32, 39).
Our results suggested that the ethanol increase in Caco-2 TJ
permeability was due to an increase in Caco-2 MLCK activity. These
findings provide a direct evidence for the involvement of MLCK in the
modulation of the TJ permeability. It may be extrapolated that the
increase in MLC phosphorylation associated with increase in TJ
permeability seen in other studies (8, 26, 39) may also be due to an
increase in MLCK activity.
Recent studies suggest that altered intestinal epithelial TJ
permeability may be an important etiologic factor in a number of
diseases, including Crohn's disease, nonsteroidal anti-inflammatory drug-associated enteritis, and diarrheal syndromes caused by
Clostridia difficile,
Vibrio cholerae, and enteropathogenic
Escherichia coli (4, 5, 8, 11, 16,
32). It had been proposed in these diseases that the
increase in intestinal epithelial TJ permeability allows the
paracellular penetration of the toxic luminal substances, culminating
in intestinal inflammation and mucosal injury (4, 11, 16, 32). The
ethanol consumption in these clinical conditions may accentuate the
increase in TJ permeability, allowing greater mucosal penetration of
the luminal substances and further exacerbating intestinal
inflammation. Consistent with this possibility, in our preliminary
studies, combined treatment of ethanol and indomethacin had an
agonistic effect on ethanol-induced increase in Caco-2 TJ permeability
(data not shown), suggesting that the combination of these two drugs
was more damaging to the TJ barrier than either drug individually.
In summary, the results of this study indicate that ethanol at low
noncytotoxic doses causes an opening of the Caco-2 TJ barrier. The
ethanol opening of Caco-2 TJ barrier appeared to be mediated by
activation of MLCK and subsequent modulation of perijunctional actin
and myosin filaments. These findings demonstrate a mechanism by which
low doses of ethanol may alter intestinal epithelial TJ barrier and
possibly promote mucosal inflammation.
 |
ACKNOWLEDGEMENTS |
We thank Susan Mills for excellent technical assistance.
 |
FOOTNOTES |
This study was supported by Veterans Affairs Merit Review and Minority
Research Training Initiative grants from the Department of Veterans
Affairs (T. Y. Ma).
The costs of publication of this
article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement"
in accordance with 18 U.S.C. §1734 solely to indicate this fact.
Address for reprint requests and other correspondence: T. Y. Ma,
Gastroenterology Section, DVA Medical Center, 5901 E. Seventh St., Long
Beach, CA 90822 (E-mail: ma.thomas_y{at}long-beach.va.gov).
Received 2 June 1998; accepted in final form 5 January 1999.
 |
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