Epidermal growth factor attenuates Clostridium
difficile toxin A- and B-induced damage of human
colonic mucosa
Martin
Riegler1,
Roland
Sedivy2,
Tacettin
Sogukoglu1,
Ignazio
Castagliuolo3,
Charalabos
Pothoulakis3,
Enrico
Cosentini1,
Georg
Bischof1,
Gerhard
Hamilton1,
Bela
Teleky1,
Wolfgang
Feil4,
J. Thomas
Lamont3, and
E.
Wenzl1
1 University Clinic of Surgery
and 2 Institute of Clinical
Pathology, University of Vienna, A-1090 Vienna;
4 Department of Surgery, Danube
Hospital, A-1220 Vienna, Austria;
and 3 Division of
Gastroenterology, Beth Israel Deaconess Medical Center, Harvard
Medical School, Boston, Massachusetts 02215
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ABSTRACT |
Epidermal growth
factor (EGF) exhibits a cytoprotective effect on gastrointestinal
epithelia via a receptor-mediated mechanism. We investigated the effect
of EGF on Clostridium difficile toxin A (TxA)- and toxin B (TxB)-induced damage of human colon.
Ussing-chambered colonic mucosa was exposed serosally to EGF before and
during luminal exposure to TxA and TxB. Resistance was calculated from potential difference and short-circuit current. Epithelial damage was
assessed by light microscopy and alteration of F-actin by fluoresceinated phalloidin. Luminal exposure of colonic strips to TxA
and TxB caused a time- and dose-dependent decrease in electrical resistance, necrosis and dehiscence of colonocytes, and disruption and
condensation of enterocyte F-actin. These effects were inhibited by
prior, but not simultaneous, serosal application of EGF (20 nM).
Administration of the tyrosine kinase inhibitor genistein (10
6 M) inhibited the
protective effects of EGF. We conclude that EGF protects against TxA
and TxB probably by stabilizing the cytoskeleton, the main target of
these toxins.
enterotoxins; genistein; human colonic epithelium
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INTRODUCTION |
EPIDERMAL GROWTH FACTOR (EGF), originally isolated from
mouse submaxillary gland (9), is mainly involved in the regulation of
gastrointestinal epithelial barrier function. In addition to modulating
epithelial maturation, proliferation, and differentiation (18, 30), EGF
also exhibits acute and chronic cytoprotective effects on
gastrointestinal epithelia (28, 35, 36, 38, 46). For example, EGF
stimulates healing of chronic gastric and duodenal ulcers in rats (28,
36) and inhibits gastric acid secretion in rats when applied
subcutaneously (35). Furthermore, luminal application of EGF increases
mucosal blood flow in ethanol-exposed rat gastric mucosa (23), and EGF
promotes epithelial restitution of intestinal cell monolayers (6, 7,
11) and of rabbit duodenum in vitro (49). Salivary glands, duodenum,
and kidney are the major sources of EGF in humans (27). Because EGF is cleaved to less active forms in gastric acid, duodenal and pancreatic EGF are suggested to be essential for protection of small and large
bowel epithelium (38). EGF mediates its biological effects via binding
to its receptor on the basolateral membrane of epithelial cells (50,
54), thus activating an intracellular signal transduction pathway (54)
that modulates cell growth and cell metabolism (52) and may also
regulate the cytoskeleton (56). Recently, it was shown that systemic
EGF attenuated epithelial damage in an experimental model of acute
colitis in rats (46).
Clostridium difficile, the causative
agent of antibiotic-associated enterocolitis in animals (1, 5) and in
humans (3, 4), produces two high-molecular-weight exotoxins: toxin A
and toxin B (40, 51). In addition to causing fluid secretion and intestinal inflammation through activation of inflammatory cells (26,
31) and nerves (8, 41) and release of mediators of inflammation (16,
42, 53), both toxins have direct effects on intestinal epithelial cells
(20, 21, 32, 45, 48). For example, toxin A damages villus tip
epithelium of guinea pig ileum in vitro (32), and both toxins A and B
impair epithelial barrier function of human colonic cancer T84 cell
monolayers in vitro (20, 21). We recently demonstrated that both toxins severely damage human colonic epithelium in vitro (48).
Electrophysiological studies show that toxin-induced damage is
paralleled by a decline of transepithelial resistance, indicating
impaired epithelial barrier function (20, 21, 32, 48). Interestingly,
epithelial cell damage by toxins is confined to the surface epithelium,
whereas the epithelium of the crypts remains intact (32, 48). This distribution of damage is probably related to the expression of toxin
receptors on villus cell brush borders with no receptors on crypt cells
(44).
Both toxins induce disruption of epithelial cell cytoskeletal F-actin,
thus impairing paracellular permeability (20, 21, 32, 48). Both toxins
are suggested to elicit their action on target cells via a
receptor-mediated mechanism involving release of intracellular calcium
(17, 44). The cellular target of C. difficile toxins is the small GTP-binding protein rho
(12, 24, 25), which regulates assembly of F-actin in fibroblasts in
vitro (10, 22, 29, 33, 34, 37, 47). In contrast, recent studies
indicated that EGF modulates F-actin assembly via activation of small
GTP-binding proteins in fibroblasts (34, 47) and that the EGF receptor
is associated with F-actin in vitro (56). According to the above
considerations, the aim of the present study was to investigate the
effects of EGF on toxin A- and toxin B-induced epithelial damage of
human colonic mucosa in vitro. Epithelial integrity was assessed by
light microscopy, morphometry, electrophysiology (15, 48), and
immunofluorescent microscopy of enterocyte F-actin as previously
described (48).
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METHODS |
Toxin preparation.
Toxins A and B were purified to homogeneity as previously described by
us (40, 44). Enterotoxic activity of toxin A was assayed in rat ileal
loops (8, 43, 53), and cytotoxic activity of both toxins was tested
against IMR-90 fibroblasts (40, 44). Purity of toxins A and B was
determined by sodium dodecyl sulfate-polyacrylamide gel electrophoresis
(SDS-PAGE) (40, 44). Concentrations of toxins A and B were expressed in
moles based on molecular masses of 308 kDa for toxin A (14) and 279 kDa
for toxin B (2).
Experimental design.
Human colonic mucosal sheets were first equilibrated in Ussing chamber
with buffer alone for 30 min. Thereafter, tissues were paired and
exposed to buffer alone or serosal buffer containing 20 nM EGF (no. E
1257, Sigma Chemie, Deisenhofen, Germany) for 60 min. Luminal solution
was then replaced with buffer containing 32 nM toxin A or 3.0 nM toxin
B for 3 h, while tissues remained incubated serosally with buffer or
buffer containing 20 nM EGF (n = 6, paired). In separate experiments, tissues were incubated with serosal
buffer or serosal buffer containing 10 nM EGF for 60 min before and
during 3 h of luminal exposure to 32 nM toxin A or 3.0 nM toxin B
(n = 3 per group). In some
experiments, after 60 min of baseline incubation, tissues were
incubated with luminal buffer containing 32 nM toxin A or 3.0 nM toxin
B and serosal buffer containing 20 nM EGF for 3 h
(n = 4 per group). Control tissues
were incubated for 4.5 h with buffer or serosal buffer containing 20 nM
EGF (n = 6, paired).
In an additional series of experiments, four human colonic explants
from a single individual were mounted in parallel and incubated with or
without serosal buffer containing EGF (20 nM) or EGF (20 nM)/genistein
(10
6 M; no. G 6776, Sigma
Chemie) 60 min before and during 3 h of luminal toxin A (32 nM) or
toxin B (3.0 nM) exposure; one tissue was incubated with buffer alone
for 4.5 h and served as control (n = 6, quadruplicate). Similar doses of EGF and genistein were previously
used to test the effects of these compounds in vitro (6, 7, 11, 49,
55).
In another series of experiments, two human colonic explants from a
single individual were mounted in parallel and incubated with serosal
buffer with or without 10
6
M genistein for 4.5 h (n = 5, paired).
Dose-response studies on the effect of C. difficile toxins A and B on human colonic mucosa were
described previously (48). Doses of C. difficile toxins A and B used in this study caused comparable electrophysiological and morphological changes in human colonic mucosal strips in vitro (48).
Ussing chamber measurements.
In this study, a total of 62 individual specimens of histological
normal sigmoid colon was used. After removal of the seromuscular layer
by blunt dissection, one to five mucosal sheets from each specimen
measuring 4-10 cm2 were
mounted vertically in Ussing chambers (19) (1 cm2 surface area; Precision
Instruments, Lake Tahoe, CA) as previously described (48, 49). Luminal
and serosal sides were bathed at 37°C with a nutrient buffer
containing (in mM) 122.0 NaCl, 2.0 CaCl2, 1.3 MgSO4, 5.0 KCl, 20.0 glucose, and
25.0 NaHCO3 (pH 7.51 when gassed
with 95% O2-5%
CO2; temperature 37°C). The
top level of fluids in both luminal and serosal reservoirs was
identical. Potential difference (PD) and short-circuit current
(Isc) were continuously recorded every 10 min. Luminal and serosal solutions were
connected via Ag-AgCl electrodes and Ringer-agar bridges to a voltmeter
(voltage-current clamp model VCC600, Physiologic Instruments).
Resistance (R) was calculated using
Ohm's law from the open-circuit PD and the
Isc. PD values
were given in millivolts, Isc in
microamperes per square centimeter, and
R in ohms times square centimeter. PD
values were corrected for the junction potentials (<0.1 mV) between
luminal and serosal solutions.
Morphometry.
After Ussing chamber experiments were performed, tissues were processed
for light microscopy. Mucosal damage was assessed by a histopathologist
(R. Sedivy), who performed morphometry on coded, paraffin-embedded,
hematoxylin and eosin-stained slides as previously described (15, 48,
49). The histopathologist was not informed about the experimental
conditions to which the tissues had been exposed. The surface area of
each tissue mounted into chambers was 1 × 1 cm. Morphometry was
performed on nine vertical sections from different locations, each of
which represented the total height and length (1 cm) of the mucosal
preparation. Thus a total length of 8-9 cm of mucosal surface was
examined for each tissue. We used a Leitz Diaplan research microscope
(objective magnification, ×4, ×6.3, ×16, ×25,
×40; ocular magnification, ×12.5; Wild Leitz, Heerbrugg,
Switzerland), a Panasonic color charge-coupled device video camera
(model WV-CD 130/G, Matshushita), an analog-to-digital monitor screen
(model PVM-1271Q, superfine pitch, Sony), a personal computer
(IBM-PS/2, model PS2; IBM, Armonk, NY), extended by insertion of a
frame grabber (ITI PCVision plus board; Imaging Technology, Woburn,
MA), and a graphic tablet (Summasketch plus 12" × 12" MM
1201; Summagraphics, Fairfield, CT). Downloading MIPSY (the Micro-based
image processing system) real-time morphometry was performed, and the
extent of epithelial damage was measured in micrometers and expressed
as percent of total mucosal surface.
Histological criteria for epithelial damage were as follows: reduced
staining intensity of epithelial cells, kariopyknosis, kariolysis and
kariorrhexis, cell disruption, formation of subepithelial blebs, and
lifting off of cells from the basal lamina (15, 48, 49).
Fluorescent microscopy.
Fluorescent staining of F-actin was performed using fluorescein
isothiocyanate-labeled phalloidin (Molecular Probes) on fresh-frozen tissue. Sections (4 µm) were fixed in 4% paraformaldehyde (pH 8.0),
washed in phosphate-buffered saline solution (PBS; pH 8.0), and
incubated for 45 min. Subsequently, slides were mounted with glycerol-PBS (1:9) and examined and photographed using a Zeiss Axiophot
fluorescence microscope (48).
Statistics.
All data were expressed as means ± SE. Statistical analyses were
performed by the Student's t-test for
paired and unpaired observations. Probabilities were regarded as
significant if they reached a 95% level of confidence
(P < 0.05).
 |
RESULTS |
Effect of EGF on toxin-induced electrophysiological changes.
The effects of EGF on toxin-induced changes on colonic
electrophysiology were assessed by comparing PD,
Isc, and
electrical resistance (R). As
expected (48), luminal exposure of tissues to 32 nM toxin A or 3.0 nM
toxin B for 3 h caused a time-dependent decrease of PD,
Isc, and
R as compared with buffer alone
(P < 0.05, Table
1). Exposure of tissues to EGF (20 nM) 60 min before and during 3 h of luminal toxin A exposure completely
prevented all colonic electrophysiological changes
(P < 0.05; Table 1, Fig.
1). Under the same experimental conditions,
EGF (20 nM) partially, but significantly, reduced toxin B-mediated PD
and R decline
(P < 0.05), whereas
Isc remained the
same (Table 1). However, 10 nM EGF failed to inhibit any of the
toxin-mediated electrophysiological responses (Table 1). These results
indicate that EGF is more potent in preventing toxin A-induced
impairment of epithelial barrier function when compared with toxin B
(Table 1, Fig. 1).
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Table 1.
Effect of epidermal growth factor and Clostridium difficile toxins A
and B on electrophysiology of human colonic mucosa
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Fig. 1.
Time course of 20 nM epidermal growth factor (EGF) on 32 nM toxin A
(A)- and 3.0 nM toxin B-induced
(B) decline of colonic resistance in
vitro. Human colonic mucosal sheets were incubated with or without
serosal buffer containing 20 nM EGF 60 min before and during 3 h of
luminal toxin A (32 nM) or toxin B (3.0 nM) exposure. Resistance was
calculated from potential difference and short-circuit current. Values
represent means ± SE of 6 paired experiments.
* P < 0.05, P < 0.01 vs. toxin.
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The time course effect of 20 nM serosal EGF on toxin A (32 nM)- or
toxin B (3.0 nM)-induced decline of transepithelial resistance is shown
in Fig. 1. When compared with tissues treated with EGF before and
during toxin A or toxin B exposure, decline of resistance of
buffer-treated, toxin-exposed tissues became statistically significant
after 60 and 120 min for 32 nM toxin A and 3.0 nM toxin B, respectively
(P < 0.05, n = 6, paired) (Fig. 1).
Coadministration of serosal EGF (20 nM) with luminal toxin A or toxin B
exposure did not have an effect in toxin-induced decline of
electrophysiological parameters (Table 2).
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Table 2.
Effect of simultaneous epidermal growth factor and Clostridium
difficile toxins A and B on electrophysiology of human colonic mucosa
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Effect of genistein on EGF-mediated protective effects.
The EGF receptor is a tyrosine kinase that is autophosphorylated upon
ligand binding (54). Thus we next investigated the effect of the
tyrosine kinase inhibitor genistein on the protective effects of EGF on
toxin A- and toxin B-induced electrophysiological changes. As shown in
Fig. 2, coadministration of serosal 20 nM EGF with 10
6 M genistein
(EGF/genistein) 60 min before and during 3 h of luminal toxin A (32 nM)
or toxin B (3.0 nM) exposure caused an ~40%
R decline when compared with
buffer-treated controls (P < 0.01, n = 6). However, this
R decline was statistically
indistinguishable from the R decline
seen in EGF/genistein-untreated, toxin-exposed tissues (Fig. 2). In
contrast, serosal 20 nM EGF before and during luminal toxin A or toxin
B exposure inhibited the R decrease
seen after toxin A and attenuated toxin B-induced
R decline (Fig. 2).

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Fig. 2.
Effect of EGF on C. difficile toxin A-
and toxin B-induced changes of colonic resistance in vitro. Human
colonic mucosal sheets were incubated with or without serosal buffer
containing 20 nM EGF (EGF + toxin) or EGF (20 nM)/genistein (Gen;
10 6 M) 60 min before and
during 3 h of luminal toxin A (32 nM) or toxin B (3.0 nM) exposure.
Controls were incubated with buffer alone for 4.5 h. Resistance was
calculated from potential difference and short-circuit current. Values
for resistance were obtained after 3 h of luminal toxin A or toxin B
exposure. Bars represent means ± SE of 6 experiments performed in
quadruplicate. * P < 0.01, P < 0.05 vs. controls.
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The time course effect of serosal EGF/genistein on colonic resistance
is shown in Fig. 3. Coadministration of
serosal EGF/genistein induced a R
decrease that became statistically significant after 60 and 120 min of
luminal exposure to toxins A (32 nM) and B (3.0 nM), respectively, when
compared with EGF-preincubated, toxin-exposed tissues
(P < 0.05, n = 6).

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Fig. 3.
Time course of EGF and EGF/genistein on 32 nM toxin A-
(A) and 3.0 nM toxin B-induced
(B) decline of colonic resistance in
vitro. Human colonic mucosal sheets were incubated with or without
serosal buffer containing 20 nM EGF or EGF (20 nM)/genistein
(10 6 M) 60 min before and
during 3 h of luminal toxin A (32 nM) or toxin B (3.0 nM) exposure.
Resistance was calculated from potential difference and short-circuit
current. Values represent means ± SE of 6 paired experiments.
* P < 0.05, P < 0.01 vs. EGF + toxin.
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Incubation of tissues with either buffer alone or serosal buffer
containing 20 nM EGF for 4.5 h did not alter basal PD,
Isc, and
R (Table 1). Incubation of tissues
with serosal genistein (10
6
M) for 4.5 h caused a statistically significant decrease of
Isc, whereas
R remained unchanged.
(Isc baseline vs.
4.5 h for controls: 80 ± 12 vs. 82 ± 11 µA/cm2;
10
6 M genistein: 87 ± 9.2 vs. 58 ± 8.6 µA/cm2;
P < 0.01;
n = 5, paired).
Histology.
No tissue strip used in these experiments showed histological criteria
of malignancy. Tissues incubated with buffer (Fig. 4A),
serosal buffer containing 20 nM EGF or
10
6 M genistein (data not
shown) showed normal colonic architecture (n = 6, paired). As shown by
morphometry (Fig. 5), incubation of tissues
with buffer alone or serosal buffer containing 20 nM EGF or
10
6 M genistein (data not
shown) for 4.5 h did not impair epithelial integrity.

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Fig. 4.
Morphological effect of EGF on toxin A- and toxin B-induced epithelial
damage on human colonic mucosa in vitro. Human colonic mucosal sheets
were incubated with luminal buffer alone for 4.5 h
(A). Tissues were incubated with
luminal buffer containing 32 nM toxin A
(B), 3.0 nM toxin B for 3 h
(C), or incubated with serosal 20 nM
EGF 60 min before and during toxin A
(D) and toxin B
(E) exposure. At end of experiments,
tissues were fixed, stained (hematoxylin and eosin), and processed for
light microscopy. Toxins A and B caused disruptions of superficial
epithelium (B and
C); in contrast, only minor
epithelial defects are detected in EGF-treated tissues
(D and
E). Magnification, ×200.
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Fig. 5.
Morphometric analysis of colonic epithelial cell damage. Human colonic
mucosal sheets were incubated with or without serosal buffer containing
10 or 20 nM EGF 60 min before and during 3 h of luminal toxin A (32 nM)
or toxin B (3.0 nM) exposure. Toxin-unexposed tissues were incubated
with buffer alone or serosal buffer containing 20 nM EGF for 4.5 h.
Mucosal sheets were fixed in Formalin and processed for light
microscopy, and morphometric analysis was performed as described in
METHODS. Results are expressed as
means ± SE; n, number of
experiments. P < 0.01, * P < 0.05 vs. toxin alone,
paired experiments. ** P < 0.05 toxin A vs. toxin B EGF treated.
P < 0.05 toxin unexposed vs. EGF treated and toxin exposed.
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The effect of serosal EGF (20 nM) on toxin A (32 nM)- or toxin B (3.0 nM)-induced mucosal damage is shown in Fig. 4,
B-E. Both toxins caused damage of the superficial epithelium while the
epithelium of the crypts remained intact (Fig. 4,
B and
C). Moreover, epithelial cell damage
had a patchy distribution as intact epithelium was adjacent to damaged
areas (Fig. 4, B and C). In contrast, tissues incubated
with 20 nM EGF 60 min before and during 3 h of luminal exposure to
toxin A (32 nM) (Fig. 4D) or toxin B
(3.0 nM) (Fig. 4E) retained an
almost confluent epithelium, with only few areas showing damage and
detachment of individual epithelial cells. The amount of epithelial
discontinuities was less in toxin A-exposed (Fig.
4D) than in toxin B-exposed tissues (Fig. 4E) after pretreatment with
EGF.
Serosal exposure of tissues to EGF (20 nM)/genistein
(10
6 M) 60 min before and
during 3 h of luminal toxin A or B exposure blocked the protective
effect of EGF on toxin A- or toxin B-induced epithelial damage and
caused morphological changes that did not differ from untreated,
toxin-exposed tissues (data not shown).
The dose-dependent effect of EGF on toxin A- or toxin B-induced
morphological damage expressed as the percent of damaged mucosa is
shown in Fig. 5. Incubation of tissues with 20 nM of serosal EGF 60 min
before and during 3 h of luminal toxin A or B exposure significantly
reduced the extent of damaged surface
(P < 0.05). Furthermore, EGF was
more potent in reducing toxin A-induced damage when compared with toxin
B (P < 0.05 toxin A vs. toxin B,
n = 6 per group). Serosal application
of EGF (20 nM) at the same time with luminal exposure to toxins A or B
did not alter toxin-mediated epithelial cell damage
(n = 4). Furthermore, 10 nM serosal
EGF did not influence toxin A- or toxin B-mediated epithelial cell damage (Fig. 5).
Serosal exposure of tissues to EGF (20 nM)/genistein
(10
6 M) 60 min before and
during 3 h of luminal toxin A or toxin B exposure caused morphological
damage that was statistically indistinguishable from the damage seen in
EGF/genistein-untreated, toxin A- or toxin B-exposed tissues
(EGF/genistein-toxin A vs. toxin A alone: 27 ± 4 vs. 24 ± 5%;
EGF/genistein-toxin B vs. toxin B alone: 28 ± 6 vs. 26 ± 4%;
n = 6, paired).
Distribution of F-actin.
Recent studies demonstrated that both toxins caused a marked decrease
in fluorescent staining for F-actin in monolayers of the T84 colonic
adenocarcinoma cell line (20, 21), guinea pig ileal enterocytes (32),
and human colonic epithelium in vitro (48). We therefore assessed the
effect of EGF on toxin A- and toxin B-induced changes of F-actin
distribution. In keeping with our recent study (48), control cells had
a polygonal shape with F-actin distributed in the peripheral
actinomyosin ring associated with the cell membrane (Fig.
6A). Tissues treated with 32 nM
toxin A (Fig. 6B) or 3.0 nM toxin B
(Fig. 6C) showed disorganization, disruption, and condensation of intracellular F-actin, confirming our
previous observations (48). In contrast, F-actin distribution of
tissues incubated with serosal 20 nM EGF 60 min before and during 3 h
of luminal toxin A (32 nM) exposure (Fig.
6D) or toxin B (3.0 nM) exposure
(Fig. 6E) was similar to
buffer-exposed controls, and in the majority of cells F-actin ring
retained a polygonal shape. Furthermore, the F-actin ring showed dotted
condensations of increased staining activity, and no condensations of
F-actin were detected in the cytoplasm of epithelial cells (Fig. 6,
D and
E). Very few epithelial cells showed
interruptions and disorganization of intracellular F-actin ring.
Incubation of tissues with serosal buffer containing 20 nM EGF alone
did not alter F-actin distribution (data not shown).
 |
DISCUSSION |
The main finding of this study was that EGF attenuated
C. difficile toxin A- and toxin
B-induced electrophysiological changes and epithelial damage of human
colon in vitro. Furthermore, administration of the tyrosine
kinase inhibitor genistein blocked the effect of EGF.
In keeping with our recent in vitro studies (20, 21, 32, 48), toxins A
and B induced a time-dependent decline of transepithelial resistance,
indicating toxin-induced impairment of epithelial barrier integrity
(Fig. 1). In contrast, incubation of tissues with serosal EGF before
and during luminal toxin exposure inhibited toxin A-induced decline of
resistance and attenuated toxin B-induced decrease in tissue resistance
(Table 1, Figs. 1 and 2). As also shown in Table 1, EGF prevented toxin
A- and toxin B-induced decline of
Isc. These data
indicate that EGF-treated, toxin-exposed tissues retained the ability
for active transepithelial electrogenic transport, whereas decline in
PD and R appears to reflect a slight increase in paracellular conductance (20, 21).
Our results show that EGF was more potent in reducing toxin A- than
toxin B-induced epithelial cell damage, although at the doses used both
toxins caused similar electrophysiological (Fig. 2) and morphological
changes (Fig. 5). The reason(s) for the different potency of EGF in the
prevention of toxin A's vs. toxin B's intestinal effects is unclear,
although they may reflect differences in the molecular pathways used by
these toxins in the human colonocyte.
Coadministration of EGF together with the tyrosine kinase inhibitor
genistein blocked the preventive effect of EGF on toxin A- and toxin
B-induced electrophysiological (Figs. 2 and 3) and morphological
changes. It is well accepted that upon binding to EGF, the EGF receptor
undergoes autophosphorylation, which in turn activates a tyrosine
kinase-dependent signal transduction pathway (54). Studies in
fibroblasts in vitro also indicate that tyrosine kinases are involved
in EGF-mediated modulation of the F-actin cytoskeleton (10, 33, 47).
Taken together, our data indicate that the mechanism of the preventive
effects of EGF in toxin-induced damage involves binding of EGF to its receptor and a tyrosine kinase-dependent signal transduction pathway.
An important finding of this study was that EGF prevented toxin A- and
toxin B-induced disruption of cytoskeletal F-actin (Fig.
6). It is well documented that both toxins
act on F-actin of the cytoskeleton, thus causing cell rounding and
detachment of cells (20, 21, 32, 48). Although toxin B disrupts F-actin of nonepithelial cells (12, 13, 40) and toxin A affects F-actin of
guinea pig ileal epithelial cells (32) in vitro, both toxins cause
disorganization of F-actin of human colonic epithelial cells in vitro
(20, 21, 48). According to the results of recent biochemical and cell
culture studies, the mechanism of C. difficile toxins involves inactivation of the small
GTP-binding protein rho (12, 24, 25). Several in vitro studies also showed that rho is involved in F-actin assembly and organization in
fibroblasts in vitro (10, 22, 29, 33, 34, 37, 47). These results
indicate that rho may be involved in interactions between the
cytoskeleton and the extracellular matrix proteins required for cell
adhesion (10, 22, 29, 33, 34, 37, 47). Although the exact mechanism is
not completely understood, rho was demonstrated to be involved in the
activation of focal adhesion kinase 125 (37, 47), which in turn
promotes binding of F-actin to the intracellular domain of integrin
receptors via the cytoskeletal glycoproteins talin, tensin, vinculin,
and
-actinin (22, 33). These results indicate that rho stimulates
cellular adhesion of cultured fibroblasts to an underlying matrix.
Microinjection of rho into toxin A- or toxin B-exposed fibroblasts
inhibited cell rounding and F-actin disorganization (12). In contrast, microinjection of C. difficile toxin
B-ribosylated rho into toxin-unexposed fibroblasts caused
disorganization of the cytoskeleton and cell rounding (25). Thus
C. difficile toxin-induced
inactivation of rho may represent the molecular switch that initiates
disruption, disorganization, and clumping of F-actin (12, 24, 25). A growing body of evidence indicates that EGF is capable of modulating actin assembly (10, 34, 47). EGF was demonstrated to stimulate F-actin
polymerization during stress fiber formation and focal adhesion
assembly when added to fibroblasts in vitro (47). This effect could be
blocked by Clostridium botulinum
exoenzyme C3-induced ADP-ribosylation of rho, which is known to
inactivate rho, thus indicating that EGF mediated its effect via
activation of rho (47). Furthermore, the EGF receptor was demonstrated
to directly bind the actin cytoskeleton (56). Because there exist no
data on the role of rho in epithelial cells, the action of EGF in our system remains speculative. Several intracellular targets are suggested
to enable EGF to inhibit toxin-induced disorganization of the
cytoskeleton: EGF could activate rho, prevent toxin-induced inactivation of rho, or stimulate actin polymerization via rho or via
direct binding of the activated EGF receptor to F-actin, thus resulting
in stabilization of the cytoskeleton (10, 22, 29, 33, 34, 37, 47, 56).
Net activation of rho would in turn promote the assembly of integrin
adhesion complexes and stabilize the cytoskeleton (22). Furthermore,
EGF was shown to induce expression of integrin receptors that in turn
could mediate increased adhesion of epithelial cells to the basal
lamina (7). Concerning time dependency of rho and EGF-induced
cytoskeletal alterations, Ridley and Hall (47) found that actin
polymerization could be observed by 30 min after microinjection of rho
into cultured fibroblasts, whereas EGF-induced actin polymerization
required up to 1 h (47). In keeping with these observations, we report here that EGF treatment had to be initiated 1 h before toxin exposure to prevent C. difficile toxin-induced
disruption of enterocyte F-actin. Taken together, EGF probably
stabilizes the actin cytoskeleton by enabling epithelial cells to
withstand the cytotoxic action of C. difficile toxins A and B.

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Fig. 6.
Effect of EGF on toxin A- and toxin B-induced disruption of cellular
F-actin. Fluorescent photomicrograph of human colonic mucosal sheet
incubated with buffer (A), 32 nM
toxin A (B), and 3.0 nM toxin B
(C) or with serosal 20 nM EGF 60 min
before and during toxin A (D) and
toxin B (E) exposure. Samples were
fixed, fresh frozen, and stained for F-actin as described in
METHODS. Note disruption of F-actin in
toxin-exposed tissues (B and
C), as opposed to ringlike
distribution of F-actin seen in controls
(A) and EGF-treated toxin-exposed
tissues (D and
E). Magnification, ×300.
|
|
Because we used mucosal preparations containing epithelial and
nonepithelial cells, we cannot exclude the possibility that the
protective effects seen are also mediated through EGF-induced activation of lamina propria cells. It is also possible that EGF attenuated toxin-induced biological effects via direct binding to the
toxin or via inhibition of toxin binding to its receptor. However,
binding studies showed that EGF does not bind to radiolabeled toxin A,
nor does it inhibit toxin A binding to intestinal membranes (C. Pothoulakis, unpublished observations).
Taken together, our results show that EGF attenuated
C. difficile toxin A- and B-induced
damage of human colonic epithelium in vitro. Although the exact
mechanism remains unclear, EGF probably elicits its effect by
stabilizing intracellular F-actin, the main intracellular target of
C. difficile toxins. Further studies
are required to elicit the molecular events by which EGF maintains intestinal epithelial barrier function.
 |
ACKNOWLEDGEMENTS |
We thank Ingrid Hammer for expert technical support.
 |
FOOTNOTES |
This study was supported by grants from the Jubiläumsfonds der
Österreichischen Nationalbank, the Anton
Dreher-Gedächtnisschenkung des Medizinischen Dekanats der
Universität Wien, National Institute of Diabetes and Digestive
and Kidney Diseases Grants DK-34583 and DK-47343, and the Crohn's and
Colitis Foundation of America.
Address for reprint requests: C. Pothoulakis, Beth Israel Deaconess
Medical Center, Div. of Gastroenterology, 330 Brookline Ave., Boston,
MA 02215.
Received 26 August 1996; accepted in final form 15 July 1997.
 |
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