Key role of PKC and
Ca2+ in EGF protection
of microtubules and intestinal barrier against oxidants
A.
Banan,
J. Z.
Fields,
Y.
Zhang, and
A.
Keshavarzian
Departments of Internal Medicine (Division of Digestive Diseases),
Pharmacology, and Molecular Biophysics and Physiology, Rush University
Medical Center, Chicago, Illinois 60612
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ABSTRACT |
Using monolayers of
human intestinal (Caco-2) cells, we showed that growth factors (GFs)
protect microtubules and barrier integrity against oxidative injury.
Studies in nongastrointestinal cell models suggest that protein kinase
C (PKC) signaling is key in GF-induced effects and that cytosolic
calcium concentration ([Ca2+]i) is essential
in cell integrity. We hypothesized that GF protection involves
activating PKC and maintaining normal
[Ca2+]i. Monolayers were pretreated with
epidermal growth factor (EGF) or PKC or Ca2+ modulators
before exposure to oxidants (H2O2 or HOCl).
Oxidants disrupted microtubules and barrier integrity, and EGF
protected from this damage. EGF caused rapid distribution of PKC-
,
PKC-
I, and PKC-
isoforms to cell membranes, enhancing PKC
activity of membrane fractions while reducing PKC activity of cytosolic
fractions. EGF enhanced 45Ca2+ efflux and
prevented oxidant-induced (sustained) rises in
[Ca2+]i. PKC inhibitors abolished and PKC
activators mimicked EGF protection. Oxidant damage was mimicked by and
potentiated by a Ca2+ ionophore (A-23187), exacerbated by
high-Ca2+ media, and prevented by calcium removal or
chelation or by Ca2+ channel antagonists. PKC activators
mimicked EGF on both 45Ca2+ efflux and
[Ca2+]i. Membrane Ca2+-ATPase
pump inhibitors prevented protection by EGF or PKC activators. In
conclusion, EGF protection of microtubules and the intestinal epithelial barrier requires activation of PKC signal transduction and
normalization of [Ca2+]i.
tubulin; growth factor; monolayer barrier permeability; Caco-2
cells
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INTRODUCTION |
THE
GASTROINTESTINAL (GI) mucosal epithelium is an essential
permeability barrier that normally restricts the passage of harmful proinflammatory molecules into the mucosa and systemic circulation (36). The loss of GI barrier integrity, in contrast, can
lead to the penetration of normally excluded luminal compounds (e.g., bacterial endotoxin) into the mucosa and can result in the initiation and/or perpetuation of mucosal inflammation and injury. This damage has
been implicated in a wide range of inflammatory illnesses including
inflammatory bowel disease (IBD) where high levels of injurious
oxidants are present (36, 41).
Accordingly, protection and maintenance of GI barrier function against
oxidant insult are critical in preventing the sustained inflammation in
these disorders. One protective strategy of tissues uses growth factors
such as epidermal growth factor (EGF) and transforming growth factor
(TGF)-
. These agents can protect the GI mucosal barrier against an
array of insults under both in vivo and in vitro conditions (10,
12, 14, 23, 32, 38, 43, 45, 57, 59, 69, 73). For example, we
previously demonstrated (10, 12, 14) that oxidants can
cause loss of barrier integrity of monolayers of human intestinal
(Caco-2) cells and that EGF or TGF-
prevents the development of
monolayer hyperpermeability. However, the intracellular mechanisms
underlying this protection remain unresolved.
We previously demonstrated (10, 14) that EGF and TGF-
protect the integrity of Caco-2 monolayers through the stabilization and remodeling of the microtubule cytoskeleton. We also showed a
critical role for microtubule integrity in the maintenance of intestinal barrier integrity under in vitro (10, 13, 14) as well as in vivo (7, 8) conditions. This stabilizing
effect appears to be a plausible mechanism for protection by growth
factors because an intact microtubule cytoskeleton is required for the maintenance of cellular integrity, structure and architecture, and
transport functions (9, 10, 11, 14, 17, 49, 72). Despite
the critical importance of the microtubule cytoskeleton in
intestinal barrier integrity, the intracellular signaling
mechanisms through which EGF stabilizes the microtubules and intestinal
barrier integrity remain elusive.
It has been proposed that Ca2+ is also important in
maintaining mucosal barrier integrity and that high intracellular
levels of this cation play a major role in promoting injury by various noxious agents including oxidants (11, 42, 75, 82, 83). Other studies using nonintestinal cellular models have suggested that
protein kinase C (PKC) signaling is a key transduction pathway stimulated by growth factors (6, 18, 60, 68, 88, 92). In
view of these considerations, we hypothesized that EGF protects the
microtubule cytoskeleton and intestinal epithelial barrier by enhancing
PKC signaling that, in turn, normalizes intracellular Ca2+
concentration ([Ca2+]i). The findings
reported herein support this hypothesis. Thus the objectives of this
study were to explore the interrelationships among PKC signal
activation, microtubule integrity, epithelial barrier integrity,
calcium homeostasis, and growth factor-mediated protection against
oxidant insult under in vitro conditions.
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MATERIALS AND METHODS |
Cell culture.
Caco-2 cells (a human intestinal cell line) were obtained from American
Type Culture Collection (Manassas, VA). The utility and
characterization of this cell line were described previously (25,
31, 54). Experiments were performed in DMEM with and without
fetal bovine serum.
Experimental design.
In the first series of experiments, isotonic saline (control) or
oxidant (H2O2 or HOCl; 0.1, 0.5, and 5 mM) was
incubated with postconfluent monolayers of Caco-2 cells for 30 min. In
the second series of experiments, monolayers were pretreated with EGF
(human recombinant EGF, 1, 5, and 10 ng/ml) or saline for 10 min before
exposure of monolayers to oxidant. All reagents were applied on the
apical side of monolayers in DMEM unless otherwise indicated. We
measured the effects of various agents alone or in combination on
Caco-2 barrier integrity, microtubule stability, PKC signal, and
calcium homeostasis. The concentrations of oxidants or EGF (10 ng/ml)
used have been shown to be effective in our laboratory (10, 12,
14) as well as others (16, 62, 63, 89). EGF and all
other chemicals were purchased from Sigma Chemical (St. Louis, MO).
To further investigate the potential importance of the PKC signaling
pathway in growth factor-mediated protection, in a third series of
experiments monolayers were preincubated (10 min) with either a PKC
activator or an inhibitor and then incubated with EGF before exposure
to oxidant. PKC activators included a synthetic diacylglycerol
(1-oleoyl-2-acetyl-sn-glycerol, OAG; 1, 50, and 100 µM) or
a phorbol ester (12-O-tetradecanoylphorbol 13-acetate, TPA;
1, 30, and 60 nM) or its inactive analog (4
-phorbol
12,13-didecanoate, 4
-PDD; 20 nM) (11). PKC inhibitors
included chelerythrine (1 µM) or bisindolylmalemide V (GF-109203X; 10 nM) or its inactive analog iGF-109203X. Controls were treated with
vehicle (0.01% DMSO or 0.2% ethanol). We confirmed that these doses
of PKC inhibitors were not toxic to cells.
In the fourth series of experiments, we investigated the role of
alterations in [Ca2+]i on growth
factor-mediated protection. Additional outcomes examined were
[Ca2+]i and Ca2+ efflux.
Monolayers were preloaded with the appropriate Ca2+ probe
(fluo 3-AM or 45Ca2+), then preincubated for 10 min with EGF, OAG, or TPA, and finally exposed to oxidant for 30 min.
Where indicated, monolayers were preincubated with either a
membrane-bound Ca2+-ATPase pump inhibitor (vanadate or
quercetine; 10 µM, 30 min) or a PKC inhibitor (as above) before the
EGF, OAG, or TPA. Vehicle solution was 0.01% DMSO or 0.2% ethanol.
In the fifth series of experiments, we investigated the effects on
monolayers of perturbations in extracellular and intracellular Ca2+. Caco-2 monolayers were preincubated (for 15 min
unless otherwise indicated) with one of the following: 1) a
Ca2+ ionophore (A-23187, 10 µM), 2) an
antagonist of voltage-operated Ca2+ channels (VOCC;
verapamil or nifedipine, 1 µM), 3) an antagonist of
store-operated Ca2+ channels (SOCC; La3+, 25 µM), 4) an extracellular Ca2+ chelator (EGTA,
1 mM added immediately before the subsequent treatment), or
5) an intracellular Ca2+ chelator
[1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid
tetra(acetoxymethyl) ester (BAPTA-AM); 20 µM, 30 min]. Experiments were performed in Ca2+-containing media (Hanks' balanced
salt solution and HEPES; 1.8 mM), Ca2+-free media, or
saline vehicle. Doses of each agent were previously shown to be
effective under similar in vitro conditions (11, 42, 75, 79, 82,
83). Where indicated, cultures were then incubated with EGF and
oxidant. Using similar protocols, EGF was replaced by OAG or TPA.
Control cells were treated with vehicle.
In a sixth series of experiments, we isolated the monomeric and
polymerized fractions of tubulin (structural protein of the microtubules), which was then analyzed using quantitative immunoblotting.
Immunofluorescent staining and high-resolution laser confocal
microscopy of microtubules.
Cells of monolayers were fixed, processed, and incubated with the
primary antibody (monoclonal mouse anti-
-tubulin) and then the
secondary antibody (FITC-conjugated goat anti-mouse) as described previously (9, 17). After staining, single cells or clumps of two to three cells were observed with an argon laser (
= 488 nm, NA 1.4; Zeiss). The cytoskeletal elements were examined in a
blinded fashion for their overall morphology, orientation, and disruption as previously described (9-14).
Microtubule (tubulin) fractionation and quantitative
immunoblotting of tubulin.
Polymerized (S2) and monomeric (S1) fractions of tubulin were isolated
and subjected to PAGE as previously described (9, 10, 14).
To quantify the relative levels of tubulin, the optical density of the
bands corresponding to immunoradiolabeled tubulin were measured with a
laser densitometer.
Determination of barrier integrity.
Permeability of monolayers was determined using a fluorescent marker,
fluorescein sulfonic acid (FSA; 200 µg/ml, 478 Da), as described
previously (10, 12, 84). After treatments, fluorescent
signals from samples were quantitated by a fluorescence multiplate
reader (excitation 485 nm, emission 530 nm; FL 600, Bio-Tek Instruments).
Measurement of PKC signal activity.
PKC activity of the cytosolic and membrane fractions was assayed as
described previously (11). Briefly, the cytosolic and detergent-solubilized (membrane) PKC fractions were collected and then
processed for their ability to phosphorylate a synthetic peptide as
described previously by others (56) and by us
(11). Sample activity was corrected for protein
concentration by the Bradford method (19), and PKC
activity was expressed as picomoles per minute per milligram of protein.
Western immunoblotting of PKC isoforms.
Membrane and soluble cell fractions were prepared as described in
Measurement of PKC signal activity. After
treatments, the distribution of PKC isoforms to membrane-associated
fractions was assessed by immunoblotting and autoradiography
(88). The PKC isoform-specific antibodies used for
immunoblotting were as follows: mouse monoclonal anti-PKC-
I,
-PKC-
II, -PKC-
, -PKC-
, -PKC-
, and -PKC-
(Santa Cruz
Biotechnology, Santa Cruz, CA) at 0.2 µg/ml and mouse monoclonal
anti-PKC-
(UBI, Lake Placid, NY) at 0.1 µg/ml (58). A
horseradish peroxidase-conjugated goat anti-mouse antibody (1:3,000
dilution; Molecular Probes, Eugene, OR) was used as the secondary antibody.
Measurement of
[Ca2+]i.
Alterations in [Ca2+]i were determined using
the selective fluorescence Ca2+ indicator fluo 3-AM
(Molecular Probes) as described previously (42, 85).
Briefly, monolayers were incubated with fluo 3 for 60 min (final
concentration of 4 µM), and the continuous fluorescent signals from
samples were then quantitated by a fluorescence multiplate reader (FL
600, Bio-Tek Instruments) at 37°C, using excitation and emission
wavelengths of 485 and 530 nm, respectively.
Measurement of
45Ca2+ efflux.
Caco-2 cells were preloaded with 45Ca2+ (10 µCi/ml) for 1 h at 37°C and then incubated with the test
agents. After centrifugation, radioactivity in the supernatant and in
the suspension of lysed cells was determined by scintillation
counting as described previously (11, 42).
Statistical analysis.
Data are presented as means ± SE. Statistical analysis was
performed using analysis of variance followed by Dunnett's
multiple-range test (34). All experiments were carried out
with a sample size of at least four to six observations per group.
P values < 0.05 were deemed statistically significant.
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RESULTS |
Protection by EGF and PKC activators
against oxidant-induced damage to intestinal monolayers.
Preincubation of Caco-2 monolayers with EGF or PKC activators (OAG or
TPA) before H2O2 dose-dependently and
significantly attenuated both barrier hyperpermeability (increases in
FSA clearance; Table 1 and Fig.
1) and microtubule cytoskeletal
disruption (decreases in % normal microtubules; Table 1). The highest
dose of EGF (10 ng/ml) provided complete (99%) protection. PKC
activators provided slightly less (90%) protection. OAG protection (of
permeability and of microtubules) was not significantly different from
EGF protection. Thereafter, we used 10 ng/ml of EGF, 50 µM OAG, or 30 nM TPA. A biologically inactive phorbol ester (4
-PDD) did not protect (Fig. 1). Figure 1 also shows that preincubation with PKC inhibitors (chelerythrine or GF-109203X), but not an inactive analog (iGF-109203X), prevented the protective
effects of EGF or PKC activators. Table 2
shows analogous effects for protection measured by percentage of cells
showing a normal microtubule cytoskeleton. PKC activators and EGF had
similar protective effects against monolayer barrier dysfunction caused
by another oxidant, HOCl (Table
3).
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Table 1.
Effect of protective agents on Caco-2 monolayer barrier integrity,
microtubule cytoskeleton, and PKC activity
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Fig. 1.
Protective effect of agents that stimulate protein kinase
C (PKC) signal activity on the epithelial (Caco-2 cell) barrier
integrity in H2O2-exposed monolayers assessed
by fluorescein sulfonic acid (FSA) clearance. Growth factor [epidermal
growth factor (EGF), 10 ng/ml] or PKC activators
[1-oleoyl-2-acetyl-sn-glycerol (OAG), 50 µM or
12-O-tetradecanoyl phorbol 13-acetate (TPA), 30 nM] were
added to the monolayers 10 min before exposure to oxidant
(H2O2, 5 mM). In select experiments, monolayers
were pretreated with either a PKC inhibitor [chelerythrine (1 µM),
GF-109203X (10 nM), or the inactive analog iGF-109203X] or a
biologically inactive phorbol ester [4 -phorbol 12,13-didecanoate
(4 -PDD), 20 nM]. Barrier integrity was calculated as apical to
basolateral flux of FSA divided by the concentration of probe in the
apical chamber, expressed as clearance. *P < 0.05 vs.
vehicle (control); P < 0.05 vs.
H2O2; +P < 0.05 vs. EGF (or OAG or TPA) + H2O2;
&P < 0.05 vs. corresponding
GF-109203X + EGF (or OAG or TPA) + H2O2. n = 4-6/group in all
experiments shown in Figs. 1-10.
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We also assessed microtubule integrity by high-resolution laser
scanning confocal microscopy. Control cells from untreated monolayers
showed a normal and stellar distribution of the microtubule cytoskeleton originating from the microtubule organizer center (MTOC or
perinuclear region) and radiating throughout the cytosol (Fig.
2A, a). Exposure of
monolayers to H2O2 produced extensive fragmentation, disorganization, and collapse of the microtubule cytoskeleton (Fig. 2A, b). Preincubation with EGF
prevented the disruption of microtubules (Fig. 2A,
c). Figure 2, B and C, shows that PKC
activators OAG (Fig. 2B, a) and TPA (Fig.
2C, a) had similar protective effects.
Preincubation with PKC inhibitor, but not inactive iGF-109203X (Fig. 2,
B, c and C, c,
respectively), abolished protection of microtubules by OAG (Fig.
2B, b), TPA (Fig. 2C, b),
or EGF (Table 2).

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Fig. 2.
The intracellular distribution of the microtubule
cytoskeleton as shown by immunofluorescent staining in intestinal cells
from monolayers. A: cell monolayers were treated with
isotonic saline (control, a), 5 mM
H2O2 (b), or EGF (10 ng/ml) and then
H2O2 (c). B: cells were
pretreated, before exposure to H2O2, with OAG
(50 µM, a), PKC inhibitor GF-109203X and then OAG
(b), or inactive analog iGF-109203X and then OAG
(c). C: cells were preincubated, before
H2O2, with TPA (30 nM, a), PKC
inhibitor GF-109203X and TPA (b), or iGF-109203X and TPA
(c). Microtubules in control cells (A, a) appear
as a intact filamentous network that courses radially throughout the
cytosol. In cells exposed to 5 mM H2O2 (without
pretreatment), the microtubules appear disrupted, collapsed, and
fragmented (A, b). In cells pretreated with EGF
(A, c), normal microtubule architecture is highly
preserved. Similarly, microtubule morphology in OAG (B,
a)- or TPA (C, a)-pretreated cells
appears intact and resembles the architecture detected in the
iGF-109203X-preincubated group (B, c and
C, c, respectively). In cells from monolayers
pretreated with GF-109203X + OAG (B, b) or
GF-109203X + TPA (C, b) before exposure to
oxidant, a clear collapse of the microtubule cytoskeleton can be seen.
Bar, 25 µm.
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Effects of EGF on PKC activity and on
intracellular translocation of PKC isoforms to membrane
fractions.
EGF or the PKC activators OAG or TPA dose-dependently increased PKC
activity as shown in Table 1. Nonprotective doses did not significantly
increase PKC activity. Figure 3 shows
that pretreatment with the PKC inhibitors chelerythrine and GF-109203X
(but not iGF-109203X) significantly inhibited the ability of EGF
or the PKC activators to stimulate PKC activity
associated with the membrane-bound fraction. The combination
of OAG (or TPA) and growth factor elicited no additional effects on PKC
activity compared with those evoked by individual agents alone,
suggesting that both agents work through the same signal pathway.

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Fig. 3.
PKC activity of the membrane-bound fractions from Caco-2
monolayers after the treatments indicated. Monolayers were preincubated
with EGF (10 ng/ml), OAG (50 µM), TPA (30 nM), 4 -PDD (20 nM), or
vehicle before oxidant insult (5 mM H2O2).
Where indicated, monolayers were pretreated with PKC inhibitors
(chelerythrine or GF-109203X or inactive analog iGF-109203X) before
protective agents. Data are means ± SE. *P < 0.05 vs. vehicle; P < 0.05 vs.
H2O2; +P < 0.05 vs. EGF (or OAG) + H2O2;
&P < 0.05 vs. GF-109203X + EGF (or
OAG or TPA) + H2O2.
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Relative levels of PKC activity in both the cytosolic and
membrane-bound fractions of Caco-2 monolayers are shown in Table 4. These data demonstrate that EGF-,
OAG-, or TPA-induced PKC activation is caused by the soluble to
membrane-bound translocation and/or shift of PKC. PKC inhibitors
did not significantly affect the distribution (translocation) of PKC in
the cytosolic and membrane fractions (Table
5), suggesting that the mechanisms of
action of these inhibitors involve potentially more direct actions on PKC itself, such as inhibition of the catalytic domain of PKC enzymes
(Fig. 3).
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Table 5.
The effect of PKC inhibitors and calcium regulating agents on PKC
translocation in Caco-2 cell monolayers
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To determine the PKC isoforms that are involved in protection, we
assessed their intracellular distribution and their shift to the
membrane-associated fractions when activated by treatments. Figure
4 shows that EGF promoted the
distribution of PKC-
, PKC-
I, and PKC-
isoforms into membrane
fractions as shown by increases in the band density of respective
isoforms. OAG and TPA caused distribution of PKC-
and PKC-
I
into membranes. Simultaneously, cytosolic cell fractions showed a
decrease in these isoforms (not shown). These data on EGF-induced PKC
isoform translocation to the membranes are consistent with the data
shown earlier on PKC activity shift from the cytosolic to the membrane
fractions.

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Fig. 4.
The effect of EGF and PKC activators on PKC isoform
translocation to the membrane fractions of Caco-2 cells. Cells were
preincubated with EGF (10 ng/ml) or control (vehicle) for 10 min. EGF
increased the distribution of PKC- (top, apparent mol
mass ~80 kDa), PKC- I (middle, ~78 kDa), and PKC-
(bottom, ~72 kDa) to the membrane fractions indicating
their activation. In select experiments, OAG (50 µM) or TPA (30 nM)
replaced EGF. Membrane extracts (75 µg protein/lane) were analyzed by
SDS-PAGE and Western blots using monoclonal anti-PKC isoform-specific
primary antibody followed by horseradish peroxidase-conjugated
secondary antibody. The region of each gel shown was between the
Mr 67,000 and 93,000 prestained molecular
weights run in adjacent lanes.
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Role of Ca2+
homeostasis in EGF protection of microtubules and
barrier integrity.
We next evaluated the effects on barrier integrity by agents known to
modulate Ca2+ homeostasis (Fig.
5). Switching to a high-Ca2+
(10 mM) medium significantly exaggerated
H2O2-induced barrier dysfunction. In contrast,
incubation of monolayers in Ca2+-free or
low-Ca2+ (100 µM) medium or with an agent that chelates
extracellular Ca2+ (EGTA) markedly prevented oxidant
damage. It is to be noted that for periods up to 1.5 h barrier
function was restored by the removal of Ca2+, but beyond
2 h low-Ca2+ medium disrupted barrier function (not
shown). Moreover, known blockers (verapamil, nifedipine) of VOCC
substantially attenuated H2O2-induced
barrier dysfunction. Furthermore, preincubation of monolayers
with an intracellular Ca2+ chelator (BAPTA-AM) or
with a SOCC antagonist (La3+) significantly prevented loss
of barrier integrity. Finally, a Ca2+ ionophore (A-23187)
not only disrupted monolayer barrier integrity by itself but also
exaggerated the effects of H2O2 on barrier dysfunction. Table 5 shows that these known calcium-modifying agents
had no significant affect on PKC translocation.

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Fig. 5.
Alterations in epithelial monolayer barrier integrity as
determined by FSA clearance after the addition of modulators that
affect Ca2+ homeostasis. Caco-2 monolayers were exposed to
oxidant (5 mM H2O2) or Ca2+
ionophore (A-23187) in 1.8 mM Ca2+-containing media. Where
indicated, monolayers were preincubated, before exposure to oxidant,
with an extracellular Ca2+ chelator (EGTA), an
intracellular Ca2+ chelator (BAPTA-AM), a voltage-operated
Ca2+ channel antagonist (VOCC; verapamil and nifedipine),
or a store-operated Ca2+ channel blocker (SOCC; lanthanum).
In other experiments, Ca2+-free, low-Ca2+ (100 µM) or high-Ca2+ (10 mM) media were used before oxidant.
Clearance was determined as described in Fig. 1. *P < 0.05 vs. vehicle; P < 0.05 vs.
H2O2.
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We then evaluated the role of Ca2+ homeostasis in growth
factor-induced protection against oxidant damage. Figures
6 and 7
show what happened when we preincubated monolayers with EGF, OAG, or TPA, with or without a Ca2+ ionophore or inhibitors of
membrane Ca2+ pumps. The Ca2+ ionophore and
each inhibitor of the membrane-bound Ca2+-ATPase pump
(quercetine, vanadate) prevented protection by growth factor (Fig.
6A) and by OAG or TPA (Fig. 6B) against
hyperpermeability and against microtubule instability (Fig. 7) in
monolayers exposed to oxidant. Unlike Ca2+ ionophore
(A-23187), Ca2+-ATPase inhibitors by themselves had no
injurious effects on the microtubules or barrier integrity.

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Fig. 6.
Alterations of the protective effects of EGF
(A) or PKC activators (B) by either
Ca2+ ionophore (A-23187) or inhibitors of membrane-bound
Ca2+-ATPase (quercetine and vanadate). Monolayers were
preincubated with A-23187 or Ca2+-ATPase inhibitor in
combination with EGF (10 ng/ml) or PKC activators (OAG, 50 µM; TPA,
30 nM), and then exposed to oxidant (5 mM
H2O2). Where shown, monolayers were incubated
with other calcium-modifying agents as indicated in Fig. 5. FSA
clearance was determined as described in Fig. 1. *P < 0.05 vs. vehicle; P < 0.05 vs.
H2O2; +P < 0.05 vs. corresponding EGF (or OAG or TPA) + H2O2.
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Fig. 7.
Percentage of Caco-2 cells of monolayers exhibiting
normal microtubule architecture as assessed by high-resolution laser
confocal microscopy. Conditions as in Figs. 5 and 6. *P < 0.05 vs. vehicle; P < 0.05 vs.
H2O2; +P < 0.05 vs. EGF (or OAG or TPA) + H2O2;
&P < 0.05 vs. H2O2
(high or low calcium) or H2O2 + EGTA (or
BAPTA-AM, verapamil, or lanthanum).
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Manipulation of Ca2+ homeostasis by the same agents that
prevented loss of barrier integrity (Fig. 5) also prevented disruption of microtubule cytoskeleton as quantitated by the percentage of cells
with normal microtubules assessed by laser confocal microscopy (Fig.
7). For instance, pretreatment of monolayers with a VOCC antagonist
(nifedipine) significantly prevented microtubule instability: 74 ± 3% of cells showed normal microtubules vs. 25 ± 4% in
H2O2-exposed monolayers.
To investigate the underlying cause of microtubule stability and/or
instability, we performed quantitative Western immunoblotting of the
polymerized tubulin pool (S2 fraction, index of stability) and
monomeric tubulin pool (S1 fraction, index of disassembly) in response
to various treatment regimes. Figure 8
shows that similar to H2O2, A-23187 caused a
significant reduction in the S2 stable polymerized tubulin and an
increase in the S1 monomeric tubulin, indicating depolymerization of
the microtubules. High-Ca2+ (10 mM) medium, which had
exacerbated loss of microtubule stability and barrier dysfunction
(Figs. 6 and 7), also significantly exaggerated H2O2-induced microtubule disassembly by
increasing tubulin depolymerization. In contrast, removal of
Ca2+ (low-Ca2+ or Ca2+-free media,
chelation of extracellular Ca2+ by EGTA or of intracellular
Ca2+ by BAPTA-AM) maintained tubulin assembly, indicating
microtubule stability. In additional experiments, VOCC blockers
(verapamil, nifedipine) or a SOCC antagonist (La3+) also
significantly prevented H2O2-induced
microtubule depolymerization, as shown by enhancement of tubulin
assembly. For instance, percent tubulin polymerization for monolayers
pretreated with nifedipine was 60 ± 0.5% vs. 35 ± 0.4%
for H2O2 alone and 66 ± 0.25% for vehicle.

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Fig. 8.
Quantitative immunoblotting analysis of the distribution
of polymerized tubulin (S2, index of stability) and monomeric tubulin
(S1, index of disassembly) in Caco-2 monolayers (A and
B). Conditions were identical to those in Figs. 5-7.
Percent polymerization of tubulin pool = [S2 divided by total
tubulin pool (S2 + S1)]. *P < 0.05 vs. vehicle;
P < 0.05 vs. H2O2;
+P < 0.05 vs. EGF (or OAG or TPA) + H2O2; &P < 0.05 vs. H2O2 (high or low calcium) or
H2O2 + EGTA (or BAPTA-AM, verapamil, or
lanthanum).
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Furthermore, a Ca2+ ionophore and inhibitors of the
membrane-bound Ca2+-ATPase pump (quercetine and vanadate)
each prevented EGF (Fig. 8A)- and OAG or TPA (Fig.
8B)-induced tubulin assembly in monolayers exposed to
oxidant, paralleling similar findings on both microtubules (Fig. 7) and
monolayer barrier integrity (Fig. 6). In contrast, pretreatment of
monolayers with the inactive 4
-PDD did not prevent tubulin
disassembly in H2O2-exposed monolayers (37 ± 1% vs. 35 ± 0.4% for H2O2).
Moreover, in monolayers incubated with H2O2, preincubation with a PKC inhibitor, GF-109203X, abolished the increase
in tubulin assembly by EGF (31 ± 0.60%), OAG (34 ± 0.30%), or TPA (36 ± 0.45%). As expected, the inactive analog
iGF-109203X was ineffective when preadministered before EGF (65 ± 0.40%), OAG (62 ± 0.34%), or TPA (61 ± 0.53%) and
H2O2 insult.
Intracellular calcium concentrations under disruptive and
protective conditions.
To further evaluate the role of Ca2+ in our Caco-2 cell
model, we used the Ca2+-sensitive dye fluo 3 (Fig.
9A) to monitor
[Ca2+]i. Monolayers exposed to
H2O2 (5 mM shown) exhibited a significant and
rapid increase in [Ca2+]i within 2 min
(peak = 455 ± 11 nM vs. 124 ± 10 nM for controls) followed by a gradual decrease. [Ca2+]i
remained significantly elevated at 30 min (304 ± 15 nM).
H2O2 at 0.1 (peak
[Ca2+]i = 171 ± 10 nM) and 0.5 (225 ± 8 nM) mM resulted in a significant but less marked rise in
[Ca2+]i than 5 mM
H2O2. Monolayers treated with a protective dose
of EGF (10 ng/ml) alone also had a rapid initial rise in
[Ca2+]i, which returned to normal by 10 min.
Monolayers pretreated with EGF (10 ng/ml) and subsequently exposed to
H2O2 had a similar early transient rise in
[Ca2+]i extending to 2 min, but unlike
H2O2 alone, this was followed by a rapid and
significant decline in [Ca2+]i (136 ± 12 nM) that was similar to control levels (127 ± 9 nM). A
nonprotective dose of EGF (1 ng/ml; EGF alone = 140 ± 15 nM) in combination with H2O2 (EGF + H2O2 = 312 ± 18 nM at 30 min) did
not normalize [Ca2+]i. Effects on
[Ca2+]i statistically similar to those of EGF
were observed when monolayers were pretreated with the PKC activators
OAG and TPA (Fig. 9B). As expected, inactive phorbol ester
4
-PDD did not normalize [Ca2+]i against
H2O2 insult (442 ± 16 nM at 10 min and
296 ± 19 nM at 30 min).

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|
Fig. 9.
Intracellular Ca2+ alterations as determined by the
Ca2+-sensitive fluorescent probe fluo 3-AM. Cells preloaded
with fluo 3-AM were preincubated with a Ca2+-ATPase
inhibitor (vanadate or quercetine) or a PKC inhibitor (chelerythrine
and/or GF-109203X or inactive analog iGF-109203X) and then exposed to
H2O2 (5 mM) in the presence or absence of
pretreatment with EGF (A) or OAG or TPA (B).
*P < 0.05 vs. vehicle (control); P < 0.05 vs. H2O2;
+P < 0.05 vs. EGF (or OAG or TPA) + H2O2.
|
|
Preincubation of monolayers (Fig. 9, A and B)
with membrane Ca2+-ATPase pump inhibitors (quercetine or
vanadate) or PKC inhibitors (chelerythrine and/or GF-109203X) abrogated
EGF (Fig. 9A)- and OAG or TPA (Fig. 9B)-induced
normalization of [Ca2+]i.
[Ca2+]i levels remained elevated during the
entire 30-min observation period. Interestingly, neither PKC inhibitors
nor Ca2+-ATPase inhibitors by themselves had any
significant effects on baseline calcium levels.
Calcium efflux under disruptive and protective conditions.
Because Ca2+-ATPase blockers prevented EGF-induced
protection and maintained [Ca2+]i at
high levels, we surmised that Ca2+ efflux is a key
mechanism for growth factor-induced protective effects on the
normalization of [Ca2+]i and the maintenance
of both microtubules and barrier function. Indeed, direct measurement
of Ca2+ efflux from monolayers prelabeled with
45Ca2+ (Fig.
10; data shown for 10-min observation
period) showed that protective doses of EGF (Fig.
10A) and PKC activators (Fig. 10B) markedly
and significantly increased Ca2+ efflux. Inhibitors of
the membrane Ca2+-ATPase pump (quercetine, vanadate)
or inhibitors of PKC, at doses that prevent EGF protection,
abolished this increase in efflux. Also, H2O2
modestly but significantly reduced Ca2+ efflux, which
may partly explain the increase in [Ca2+]i
induced by this agent.

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Fig. 10.
Effect on 45Ca2+ efflux from
Caco-2 monolayers by EGF (A), PKC activators (OAG or TPA or
inactive 4 -PDD) (B), PKC inhibitors (chelerythrine
and/or GF-109203 X or the iGF-109203 X), or Ca2+-ATPase
inhibitors (quercetine or vanadate). After preincubation with these
agents monolayers were exposed to oxidant (5 mM
H2O2). *P < 0.05 vs.
vehicle. P < 0.05 vs.
H2O2. +P < 0.05 vs. EGF
(or OAG or TPA) + H2O2.
|
|
Finally, we did not observe any significant Ca2+ efflux in
monolayers pretreated with the nonprotective doses of 1 ng/ml EGF (49 ± 4%), 10 µM OAG (51 ± 3%), or 1 nM TPA (48 ± 2%) compared with vehicle (48 ± 3%), paralleling the lack of
effects of these doses on FSA clearance (barrier function), microtubule
integrity, PKC activity, and [Ca2+]i.
 |
DISCUSSION |
Investigating the mechanisms by which growth factors protect
intestinal cells and the intestinal permeability barrier against oxidant-induced damage, as the current study has done, is clinically important because free radical damage results in a "leaky gut" that
is thought to be one of the underlying mechanisms in IBD (14, 36,
41). We focused on mechanisms involving the microtubule cytoskeleton for two reasons: 1) because it is well
established that maintaining an intact microtubule cytoskeleton is
essential for the integrity of barrier permeability as well as normal
cellular function and structure of enterocytes (8-10, 13,
14, 31), and 2) because our recent studies (10,
14) found that maintaining an intact microtubule cytoskeleton is
necessary for protection by growth factors. On the basis of the results
from the current study, we conclude that the ability of EGF to prevent
oxidant-induced microtubule disruption and barrier leakiness in our
model is mediated by enhanced PKC activity and normalization of
Ca2+ homeostasis. To our knowledge, this is the first time
this mechanism has been ascribed to the defense and repair of GI
epithelial cells.
We felt particularly confident in drawing this conclusion because it
was supported by several independent lines of evidence. First,
pretreatment of intestinal monolayers with EGF, which prevents oxidant-induced hyperpermeability, simultaneously increases PKC activity and evokes a cascade of changes that are consistent with the
proposed mechanism. EGF activates specific PKC isoforms, increases the
proportion of cells showing a normal microtubule architecture, increases the size of the polymerized tubulin pool while decreasing the
size of the monomeric tubulin pool, normalizes cytosolic
Ca2+, and increases Ca2+ efflux.
Second, these effects of EGF are selectively mimicked by PKC activators
(OAG and TPA) or agents known to affect Ca2+ homeostasis.
Third, the protective effects of EGF or PKC activators are attenuated
by specific PKC inhibitors. Fourth, agents that are known to
dysregulate Ca2+ homeostasis worsen oxidant-induced
cytoskeletal or barrier disruption or induce disruption themselves.
Fifth, our previous work (10, 14) shows a significant
correlation between barrier disruption and microtubule disruption, and
the current study shows additional significant correlations: protection
against barrier disruption and PKC activity (r = 0.89, P < 0.05), protection against barrier disruption and
[Ca2+]i (r = 0.95, P < 0.05), protection against barrier disruption and
Ca2+ efflux (r = 0.98, P < 0.05), protection against microtubule disruption and PKC activity
(r = 0.90, P < 0.05), protection
against microtubule disruption and [Ca2+]i
(r = 0.97, P < 0.05), protection
against microtubule disruption and Ca2+ efflux
(r = 0.96, P < 0.05), and increase in
PKC and in Ca2+ efflux (r = 0.88, P < 0.05).
Looking further into the mechanisms of EGF protection, our study
suggests that not all PKC isoforms and not all Ca2+
pathways are equally important. Our studies on redistribution of PKC to
membrane fractions suggest that
-,
I-, and
-isoforms of PKC
are activated by EGF. Consistent with these findings, it is known that
PKC profoundly affects cellular functions in nonintestinal cell types
(3-5, 22, 26, 28, 32, 35, 47, 48, 50, 71, 74, 78, 88,
94), and several studies show that growth factors (e.g., EGF)
activate PKC in several cell types including the epithelium (6,
18, 88, 92). For example, EGF caused translocation of both
PKC-
and PKC-
I into membranes in canine gastric cells
(88) and caused PKC-
membrane association in mammary
epithelial cultures (18). Moreover, PKC has been suggested to be a mediator of EGF-induced alterations in the actin component of
the cytoskeleton in HeLa and corneal endothelial cells (22, 39) and of EGF-mediated inhibition of canine parietal cell
function (88).
In our studies, OAG and TPA only induced activation of classic PKC-
and PKC-
I. This finding is not surprising because studies in non-GI
models (i.e., fibroblasts) showed that these PKC activators can only
induce activation of classic PKC isoforms (33). Our current studies using an acute model of PKC activation are also consistent with previous reports in which acute administration of a low
dose of a PKC activator (TPA) caused rapid activation of PKC in
parietal cells and subsequently inhibited cell secretions via
PKC-mediated processes (18). Also, within 5-15 min
after exposure to TPA (0.1 µM), there were rapid PKC-mediated effects on the enhancement of cell migration concomitant with rapid
reorganization of the actin component of the cytoskeleton in corneal
endothelial cells (39). In other non-GI cells (e.g.,
fibroblasts), after 15 min in TPA, PKC redistributed to cell membranes
and was associated with rapid cytoskeletal remodeling (33, 86,
93). In further support of our findings are recent studies
proposing that diacylglycerol apparently modulates intestinal
epithelial (Caco-2) permeability (5, 48). The effects of
PKC activation in cellular models are complex and, we acknowledge, vary
with different experimental settings and cell types (5, 26,
48). In our studies, PKC inhibitors did not affect PKC
translocation (Table 5). Our PKC activity measurements (Fig. 3),
however, show that these inhibitors prevent PKC activation and that the
inhibitory effects of PKC inhibitors in our studies were probably
caused by their direct inhibition of the catalytic domain of PKC as
suggested by others (6, 68, 78, 92).
[Ca2+]i in eukaryotic cells is exquisitely
balanced, and derangement of [Ca2+]i
homeostasis can lead to the disruption of many cell functions including
the cell's cytoskeletal network (37, 40, 76). Our calcium
experiments, which used a wide variety of known calcium-modifying agents, were all consistent with the idea that decreases in
[Ca2+]i and enhanced efflux of
Ca2+ are particularly important in EGF protection. This
conclusion is supported by several findings. EGF enhanced
Ca2+ efflux and normalized
[Ca2+]i. Protection by EGF was significantly
blunted by Ca2+ ionophore and membrane Ca2+
pump inhibitors. Additionally, manipulations that lowered
extracellular or intracellular Ca2+ (e.g., EGTA, BAPTA-AM,
low Ca2+, or VOCC and SOCC Ca2+ channel
antagonists) protected against oxidant-induced disruption. Maneuvers
that raised extracellular or intracellular Ca2+ levels
(e.g., high-Ca2+ media) had the opposite effect,
potentiating oxidant-induced cytoskeletal injury and barrier
dysfunction. It is to be noted that the effects of these known
Ca2+-modifying agents did not involve alterations in PKC
translocation in our model because these agents did not significantly
change PKC distribution (Table 5). Furthermore, EGF prevented the
sustained rise in [Ca2+]i that is induced by
oxidants concomitantly with maintaining intact microtubules, tubulin
assembly, and barrier integrity.
Although the precise mechanism through which changes in
Ca2+ homeostasis mediate protection by EGF is not
established, our findings suggest that EGF elicits its protective
effects by PKC-induced changes in [Ca2+]i.
EGF or PKC activators abolished the prolonged elevation of [Ca2+]i that was induced by oxidants and
normalized [Ca2+]i; these effects were
abrogated by PKC inhibitors. Our study also suggests that maintaining
Ca2+ homeostasis through Ca2+ efflux is an
important mechanism through which EGF protects. The fact that membrane
Ca2+-ATPase inhibitors, Ca2+ ionophores, and
PKC inhibitors prevented EGF protection supports the proposed
mechanism. Our proposed mechanism is consistent with previous studies
in non-GI cells in which phorbol 12-myristate 13-acetate (a PKC
activator) enhanced Ca2+ efflux via a proposed
PKC-dependent extrusion mechanism in human T lymphocytes, osteoblasts,
and neuronal cells (3, 4, 90, 94). Another study showed
that TPA reduced the amplitude of Ca2+ influx in human lung
epithelial cells, suggesting stimulation of Ca2+ efflux
(46).
Several findings suggest that the proposed mechanism for EGF protection
is generalizable. First, in the present study, we observed similar
effects when HOCl was the oxidant rather than H2O2. This is consistent with reports
(41) that both oxidants are known to be elaborated by
activated neutrophils at sites of inflammation and that both appear to
be important in IBD. Second, in previous studies (10, 12,
14) we reported similar protective effects for TGF-
, a
structurally similar growth factor elaborated by intestinal mucosa that
appears to act through the same EGF receptor. Indeed, other studies
have shown that it is not possible to convincingly disassociate the
biological activities of EGF and TGF-
in any cell population,
including GI epithelium (32, 59, 77). It is therefore
reasonable to assume that most biological effects of TGF-
will also
be produced by EGF. Regarding protection of intestinal tissue, the only
major difference between EGF and TGF-
appears to be their site of
origin. Whereas TGF-
is elaborated directly by the intestinal
epithelium (32), EGF found in the intestine is normally
synthesized elsewhere. For instance, several previous studies showed
that salivary EGF as well as EGF contained within secretions of
Brunner's glands and exocrine pancreas are the major source of
intestinal EGF and that they play a key role in protection of small and
large bowel (15, 59, 61, 65). For example, EGF prevented
damage to the intestinal epithelium in an animal model of colitis
(IBD), similar to our oxidant model of IBD in vitro (15,
61). EGF also prevented Clostridium difficile toxin-induced epithelial damage to human colon in vitro
(70). Although the relative contributions of these
two growth factors to in vivo defense and repair of the intestinal
mucosal barrier remain to be fully established, our study already
suggests that targeting this protective mechanism in new drug
development studies may be worthwhile, either by enhancing the
signaling pathways of endogenous growth factors or by introducing
exogenous agents that are EGF and/or TGF-
mimetics. Another key step
will be to establish that this putative mechanism for intestinal
barrier protection by growth factors is operative in vivo in animals
and humans, an idea that has already received some support (2, 15, 30, 44, 51-53, 55, 61, 64, 66, 67, 70, 80, 91). For
instance, Wright et al. (91) demonstrated the existence of
EGF-like immunoreactivity in a novel cell lineage derived from intestinal stem cells in the injured intestinal mucosa in the instances
of IBD and peptic ulcer disease. Overall, the preponderance of evidence
suggests that our EGF model is a relevant model for studying the
intracellular mechanism of intestinal protection.
An apparent limitation of our conclusion that PKC signal activation is
a major mediator of EGF-induced protection is that protection of
barrier integrity against H2O2 insult by PKC
agonists (OAG, TPA) was slightly less (92% and 85%, respectively)
compared with EGF (98%). However, not all PKC isoforms were activated
by OAG and TPA as they were by EGF. Thus it appears that the greater protection by EGF is due to its ability to activate additional PKC
isoforms. An alternative interpretation is that as much as 10-15%
of protection may be due to non-PKC pathways. A similar pattern was
found for protection against HOCl. Also, although it is reasonable to
state that increases in PKC and decreases in
[Ca2+]i are necessary components of the
protective mechanism for EGF, they may not be entirely sufficient;
other molecular mechanisms may also be involved. Some studies
(20, 24, 27, 29) have suggested that protection by growth
factors may involve stimulation of ornithine decarboxylase (ODC) and
subsequent synthesis of essential polyamine compounds. For instance,
EGF enhances intestinal repair against ethanol injury, and this effect
is mediated, in part, by ODC because an irreversible inhibitor of this
enzyme,
-difluoromethyl ornithine, significantly prevented
protection (20). Moreover, EGF can modulate the expression
of the rate-limiting enzyme ODC, which is a key step in the
biosynthesis of polyamines (24, 29, 51). Specifically, EGF
not only enhances the intestinal mucosal expression of ODC but also
increases the levels of another polyamine regulatory enzyme, diamine
oxidase, in Caco-2 cells. A recent series of studies, including our own
(7, 8), showed that polyamines contribute, at least in
part, to the repair of GI injury (51, 87). We also showed
(7, 8) that polyamines are important for cytoskeletal
reorganization and healing under in vivo conditions. Thus the
ODC-polyamine system could provide an additional component to the
mechanism through which EGF protects the intestinal mucosa. Future
studies will be needed to determine whether there are interactions
between PKC signal enhancement and normalization of Ca2+
homeostasis by EGF and EGF-induced changes in the ODC-polyamine system.
In summary, our studies demonstrate that EGF protects the intestinal
monolayer barrier against oxidant damage by preventing damage and
disruption of the microtubule cytoskeleton, and the current study shows
that activation of PKC signaling and normalization of intracellular
Ca2+ appear to be key mechanisms. Although the total
protective mechanism for growth factors remains to be established, on
the basis of the current report and our previous work
(9-14) there is already a rationale for considering
development of certain new drug targets for IBD. In particular,
inhibitors (e.g., selective antioxidants) could be developed that
target those reactions that have been shown by our studies to be
required for oxidative-induced damage. Alternatively, agents could be
developed that enhance or mimic the protective effects of growth
factors against oxidant-induced insult such as occurs under
inflammatory conditions.
 |
ACKNOWLEDGEMENTS |
This work was supported in part by a grant from Rush University
Medical Center.
 |
FOOTNOTES |
Portions of this work were presented at Research Forum of the annual
meeting of the American Gastroenterological Association in San Diego,
CA, 2000 (10a).
Address for reprint requests and other correspondence: A. Banan, Rush Univ. Medical Center, Division of Digestive Diseases, 1725 W. Harrison, Suite 206, Chicago, IL 60612 (E-mail:ali_banan{at}rush.edu).
The costs of publication of this
article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement"
in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Received 6 June 2000; accepted in final form 21 November 2000.
 |
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