Department of Surgery, Beth Israel Deaconess Medical Center, Boston, Massachusetts 02215
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ABSTRACT |
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G526, 2002. First published December
5, 2001; 10.1152/ajpgi.00393. 2001.Many studies have implicated
F-actin in the regulation of gastric acid secretion using cytochalasin
D (CD) to disrupt apical actin filaments in oxyntic cells. However, it
is known that CD also affects mucosal permeability by disrupting tight junction structure. Here we investigated the contribution of F-actin to
mucosal permeability and acid secretion in the stomach using CD.
Stomachs were mounted in Ussing chambers and acid secretion (stimulated
or inhibited), transepithelial resistance (TER), mannitol flux,
bicarbonate transport, and dual mannitol/sodium fluxes were determined
with or without CD. H+ back diffusion was predicted from
its diffusion coefficient. Incubation with CD resulted in a significant
reduction in stimulated acid secretion. TER was unchanged in stimulated
tissues but significantly reduced in inhibited tissues. Mannitol flux,
bicarbonate transport, and H+-back diffusion increased
significantly with CD. However, the rates of bicarbonate and
H+ flux were not large enough to account for the inhibition
of acid secretion. These findings demonstrate that actin filaments
regulate paracellular permeability and play an essential role in the
regulation of acid secretion in the stomach.
cytochalasin D; gastric; parietal cell
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INTRODUCTION |
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ACTIN HAS RECENTLY
TAKEN center stage in the stomach due to its potential importance
in the regulation of gastric acid secretion. Actin accounts for nearly
5% of the total protein in parietal cells, of which 90% is
filamentous (F-actin) and 10% is globular (G-actin) (8).
In addition, there is a differential localization of actin isoforms in
parietal cells, where apical actin consists of -actin and
basolateral actin consists of
-actin (8, 30, 31).
Apical actin filaments localize predominantly to microvilli in
stimulated parietal cells (24, 30) or to surface folds in
stimulated oxynticopeptic cells (12). Although not studied in detail, apical actin filaments probably associate with the tight
junction in gastric epithelial cells, as they do in epithelial cells of
other gastrointestinal tissues and in cultured epithelial cells
(2, 11, 14, 16, 20, 21, 26, 27).
To determine that actin filaments are involved in gastric acid secretion, F-actin polymerization was inhibited in both intact tissues (3, 10) and in isolated gastric glands (8, 23) with cytochalasin B, D, or E. In the rat stomach, in vivo, cytochalasin E caused a dose-dependent inhibition of H+-secretion within 30 min (10). In the piglet stomach, in vitro >30 µM cytochalasin B abolished acid secretion within 50 min (3). Cytochalasin D (10 µM) promoted the depolymerization of F- to G-actin and inhibited acid secretion in rabbit gastric glands irrespective of the stimulant used, be it carbachol, dibutyryl cAMP, or histamine (8, 23). In each study, the disruption of actin filaments and resulting inhibition of acid secretion was attributed to a direct effect of the disrupted actin filaments on some aspect of stimulated acid secretion. The role of tight junctions in regulation of paracellular permeability, essential for trapping H+ in the lumen so it can be measured, was not considered.
In epithelial cells, tight junctions are structurally associated with a perijunctional actomyosin ring that contracts and acts to regulate paracellular permeability of the mucosa (28). In the intact ileum (20, 21), jejunum (18), and gallbladder (2), as well as in cultured Caco-2 (17), T84 (11) and Madin-Darby canine kidney (MDCK; 14, 16, 26, 27) cells, a significant reduction in transepithelial resistance (TER) and an increase in paracellular permeability accompanies the disruption of actin filaments with cytochalasin B or D. In addition, permeability in the presence of cytochalasin D is size-selective, where mannitol (3.6 Å) but not inulin (11-15 Å) can cross the disrupted junction (20). Chen, et al. (4) found that cytochalasin D also decreased TER and increased mannitol but not inulin flux, in monolayer cultures of canine gastric epithelial cells. These results suggest that TER and paracellular permeability are also regulated by F-actin (at the tight junction) in gastric epithelial cells.
Thus the purpose of this study was to determine whether apical F-actin regulates gastric acid secretion. If apical actin filaments regulate mucosal permeability in the stomach, it is possible that significant hydrogen ion back diffusion (mucosal to serosal) and bicarbonate ion flux (serosal to mucosal) will titrate secreted H+ when actin filaments are disrupted with cytochalasin D. We found, however, that despite a significant increase in paracellular permeability in the presence of cytochalasin D, the back diffusion of hydrogen ion and flux of bicarbonate are minimal and cannot account for the significant reduction in acid secretion.
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MATERIALS AND METHODS |
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Preparation of the bullfrog gastric mucosa for the Ussing chamber
studies.
Animals used for this study were maintained in accordance with the
guidelines of the Committee on Animals at the Beth Israel Deaconess
Medical Center and those prepared by the committee on Care and Use of
Laboratory animals by the National Research Council. Bullfrogs
(Rana catesbeiana) caught in the wild were purchased from
West Jersey (Wenonah, NJ) and kept at room temperature in large water
tanks until use. Stomachs were removed from pithed frogs, and the
fundic mucosa was stripped from underlying external muscle layers and
submucosa to bare the muscularis mucosa as described previously
(12). Stripped mucosae were divided into two halves (one
experimental and one control); each was mounted between two Lucite
halves of an Ussing-type chamber with an exposed mucosal area of 0.636 cm2. Mucosal surfaces (from here onward called
"luminal") were bathed with a solution containing (in mM): 102.4 Na+, 4.0 K+, 0.8 Mg2+, 1.8 Ca2+, 91.4 Cl, 10.1 SO
, 0.8 SO
Mannitol flux studies with cytochalasin D in stimulated and
inhibited tissues.
To stimulate acid secretion, 0.1 mM histamine and 1 mM carbachol
(histamine-carbachol) were added to the nutrient solution. To inhibit
acid secretion, both luminal and nutrient compartments were washed with
buffer and then the tissues were incubated with 1 mM aqueous cimetidine
until acid secretion reached 0 µeq · h1 · cm
2. Once the
tissues were stimulated or inhibited, 0.1% DMSO or 20 µM
cytochalasin D in 0.1% DMSO was added to the nutrient solution and
3H-mannitol (50 µCi, 15-30 Ci/mmol, NEN Life Science
Products, Boston, MA) was added to the luminal solution. For mannitol
flux studies, duplicate aliquots (0.25 ml each) were taken every 30 min
from the nutrient solution, replacing this aliquot with an equal volume
of unlabeled nutrient buffer containing histamine-carbachol or
cimetidine and DMSO or cytochalasin D and DMSO (CD/DMSO), respectively. Samples from both stimulated and inhibited tissues were diluted with 3 ml of scintillation fluid (Atomlight, NEN Life Science Products,
Boston, MA) and the amount of 3H in each sample was
determined by liquid scintillation (Packard Instrument, Meriden, CT).
Mucosal-to-serosal flux was calculated by standard techniques.
Electron microscopy. Frog tissues from the Ussing-chamber were fixed overnight at 4°C with 2% glutaraldehyde in 0.1 M cacodylate buffer (pH 7.4), postfixed for 1 h at 4°C with 1% osmium tetroxide in 0.1 M cacodylate buffer (pH 7.4), and stained overnight at 4°C with 2% aqueous uranyl acetate. Tissues were dehydrated in graded alcohols and propylene oxide and embedded in LX112 resin. Thin sections, cut parallel to the long axis of gastric glands, were placed on Formvar- and carbon-coated grids and examined with a JEOL 100CX electron microscope.
Measurement of bicarbonate transport (serosal to mucosal) in
inhibited tissues.
Tissues were stripped and mounted in Ussing chambers, as described
above, except that nutrient and luminal solutions were maintained at pH
7.4. To inhibit tissues, 0.3 mM omeprazole (Astra Hassle, Sweden) was
added to the nutrient solution until the rate of acid secretion reached
0 µeq · h1 · cm
2 and TER
was at least 400 Ohm/cm2. Bicarbonate transport was
measured using a pH-stat device and was calculated from the volume of 5 mM HCl needed to titrate the luminal solution to a constant pH of 7.4. Anoxia was applied to differentiate between passive and active
bicarbonate transport. For this, 100% N2 was applied to
the luminal solution and 5% CO2- 95% N2 to
the nutrient solution for 30 min.
[3H]mannitol-22Na flux studies.
Tissues were stripped, mounted in Ussing chambers (nutrient solution pH
7.2 and luminal solution pH 4.7), and inhibited with 0.3 mM omeprazole
until the rate of acid secretion reached 0 µeq · h1 · cm
2 and TER
was at least 400 Ohm/cm2. Buffered solutions were replaced
(on both sides) with nutrient buffer (pH 7.4) and DMSO or CD/DMSO were
added for 90 min. This is the time required to maximally increase
paracellular permeability. Tissues were then switched from open- to
closed-circuit conditions and allowed to equilibrate for 30 min in the
presence of luminal 3H-mannitol (50 µCi) and
22Na (10 µCi, 287.3 mCi/mg; NEN Life Science, Boston,
MA). Four duplicate aliquots (0.25 ml) were then taken every 30 min
from the nutrient solution, replacing this aliquot with an equal volume of unlabeled nutrient buffer containing omeprazole and DMSO or CD/DMSO,
respectively. Samples were diluted with 3 ml of scintillation fluid
(Atomlight) and the disentegrations per minute of
3H-22Na in each sample was determined by liquid
scintillation. Quench curves for 3H and 22Na
were done before counting. Mucosal-to-serosal flux was calculated by
standard techniques.
Statistical analysis. Combined data were expressed as mean ± SE. Statistical analyses of data were done with SigmaStat software (Jandel Scientific Software, San Rafael, CA) using the unpaired t-test for analysis of two groups or one-way analysis of variance for many groups. Best-fit regression lines were calculated in SigmaPlot for dual 3H-22Na experiments. Differences were regarded as statistically significant at P < 0.05.
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RESULTS |
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Cytochalasin D inhibits stimulated acid secretion.
Acid secretion in the frog fundic mucosa attained a maximal rate of
6.68 ± 0.17 µeq · h1 · cm
2 by 60 min after stimulation and remained constant for
3 h in the presence of 0.1% DMSO (Fig.
1A). In contrast, tissues that were maximally stimulated and then incubated from the serosal surface
with 20 µM cytochalasin D in 0.1% DMSO (CD/DMSO) showed a
significant reduction in acid secretion from 6.46 ± 0.06 to 1.69 ± 0.54 µeq · h
1 · cm
2 within 3 h (Fig. 1A). In stimulated
tissues, TER was 81.6 ± 5.79 Ohm/cm2 in tissues
treated for 3 h with DMSO and 78.4 ± 16.39 Ohm/cm2 after 3 h in tissues treated with CD/DMSO
(Fig. 1B).
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Cytochalasin D decreases TER in inhibited tissues.
Tissues inhibited with cimetidine had an acid secretion rate of 0.00 µeq · h1 · cm
2 (not
shown) and a TER of ~500 Ohm/cm2 (Fig. 1C).
Incubation of inhibited tissues with DMSO resulted in a modest decline
in TER to 305.3 ± 17.8 Ohm/cm2 within 4 h (Fig.
1C). In contrast, incubation of inhibited tissues with
CD/DMSO caused a significant decrease in TER to 106.8 ± 12.1 Ohm/cm2 within 4 h (Fig. 1C).
Cytochalasin D causes an increase in paracellular permeability. The significant decrease in TER in inhibited tissues treated with CD/DMSO suggested that treatment with cytochalasin D results in an increase in paracellular permeability. To test this hypothesis, unidirectional (mucosal to serosal) mannitol fluxes were studied in both inhibited and stimulated tissues treated with DMSO or with CD/DMSO.
In inhibited tissues treated with DMSO, or CD/DMSO for 60 min, the rate of mannitol flux was comparable at 12.3 ± 1.3 (×10
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Cytochalasin D does not disrupt the structure of oxynticopeptic
cells as determined by electron microscopy.
To determine whether CD/DMSO damages the mucosa to result in an
increase in paracellular permeability, mucosal structure was evaluated
in inhibited tissues by electron microscopy (Figs.
3 and
4). Low magnification micrographs
showed that changes in mucosal structure in inhibited tissues treated
for 4 h with CD/DMSO were limited to dilation of the gland lumen
and a reduction in the number of surface folds at the apical surface of
oxynticopeptic cells (Fig. 3). Tight junctions in oxynticopeptic cells
treated with CD/DMSO were identical to those from control tissues and showed no increase in membrane dilatations or condensed perijunctional actin (Figs. 3 and 4).
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HCO1 · cm
2 after
5 h. The serosal to mucosal HCO
1 · cm
2 after
5 h, which was a significant (threefold) increase compared with
control tissues (Fig. 5).
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Back-diffusion of H+ contributes little to the reduction in acid secretion that occurs with cytochalasin D. With increased paracellular permeability, H+ may be lost from the lumen by movement across the tight junction into the serosal solution. Because it is difficult to study the flux of H+, per se, H+ movement was predicted from measurements of Na+ and mannitol flux under short circuit conditions across the tight junction. Rationale for this study was based on the validated assumption that the movement of mannitol, Na+, and H+ through the paracellular space would be related to the movement of these species in free solution (7). We can determine the rate of Na+ and mannitol flux across the paracellular space under our experimental conditions. Then, the rate of H+ flux across the paracellular space can be predicted, under the same experimental conditions, from its diffusion coefficient in free solution.
When dual Na+ and mannitol fluxes were measured in the inhibited stomach, in the presence of DMSO or CD/DMSO, the ratio of Na+ to mannitol flux was 10.63 (Fig. 6A). Na+ and mannitol fluxes had a strong correlation (r2 = 0.91) and the intercept was different from 0 (Fig. 6A). The ratio of the free-solution diffusion coefficients of each species corrected for the concentration in the luminal solution, DNa (1.334 × 10
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DISCUSSION |
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The present study shows that disruption of actin filaments with
cytochalasin D significantly inhibits acid secretion in the frog
gastric mucosa. After incubation with cytochalasin D, acid secretion
decreased from 6.46 ± 0.06 to 1.69 ± 0.54 µeq · h1 · cm
2 and was
accompanied by a significant increase in passive ion flux across the
tight junction. We show here, however, that the increase in ion flux
with cytochalasin D would not result in significant acid depletion,
either by titration with HCO
1 · cm
2) from 6.46 to 6.12 µeq · h
1 · cm
2.
Similarly, the cytochalasin D-induced increase in H+ back
diffusion would reduce acid secretion (by 0.00396 ± 0.00029 µeq · h
1 · cm
2) from 6.12 to 6.116 µeq · h
1 · cm
2.
Thus our results demonstrate that disruption of actin filaments with
cytochalasin D has a specific inhibitory effect on gastric acid
secretion that cannot be explained by an increase in passive ion flux
across the tight junction.
Our results, using cytochalasin D, agree with previously reported
findings in whole tissue preparations in which cytochalasin E or B
inhibited acid secretion in the rat and piglet gastric mucosa,
respectively (3, 10). In addition, our results are consistent with those where cytochalasin B, E, or D inhibited acid
secretion in isolated rabbit gastric glands (6, 8). However, our results are in contrast to studies in the frog gastric mucosa, where cytochalasin B had no effect on acid or pepsinogen secretion (13). It is not known why our results differ
from other studies in the same species. Perhaps our method of stripping the submucosa from the muscularis mucosa (12) facilitates
the penetration of cytochalasin D in the in vitro preparation.
Alternatively, cytochalasin D may have a different effect on the frog
gastric mucosa than does cytochalasin B. Experiments done by us to
address these issues provided the following results. First, stripping the submucosa significantly influenced the rate of stimulated acid
secretion. Acid secretion was 2.14 ± 0.28 µeq · h1 · cm
2 in
unstripped tissues, which is reduced considerably from the rate of acid
secretion in stripped tissues as described in Fig. 1. Without the
mucosa stimulated maximally, it is difficult to assess the effects of
cytochalasin D or B on stimulated acid secretion. Second, cytochalasin
B reduced the rate of stimulated acid secretion more in stripped (86%)
than in unstripped (55%) tissues (data not shown). These results
demonstrate that both cytochalasin D and B inhibit stimulated acid
secretion in the frog gastric mucosa, but that drug effectiveness is
dependent on tissue preparation.
We show here that TER in the gastric mucosa is influenced by the rate
of passive paracellular movement of ions, consistent with studies using
cytochalasin D in the intestine (20) or with chemical
hypoxia in T84 cells (22). In the stimulated gastric mucosa (frog), previous studies have shown that TER is low (80-100 Ohm/cm2) due to a high rate of net Cl and
Na+ flux across the mucosa (9). In contrast,
previous studies have shown that TER is high (400-600
Ohm/cm2) in the inhibited gastric mucosa (frog) due to a
significant decrease, compared with stimulated tissues, in net
Cl
and Na+ flux across the mucosa
(9). In both cases, an assumption was made that the tight
junction has similar permeability properties so that it is not a factor
in determining TER. When the mucosa is treated with cytochalasin D,
however, we demonstrate a significant increase in paracellular
permeability in both stimulated and inhibited tissues. In inhibited
tissues, this increase in passive ion permeability fully accounts for
the decline in TER, demonstrating that TER is dependent, in part, on
mucosal permeability in the gastric mucosa. In stimulated tissues,
treatment with cytochalasin D both inhibits acid secretion (that should
increase TER) and increases paracellular permeability (that should
decrease TER). It is not clear, therefore, why TER remains low during
treatment with cytochalasin D. It is possible that the rise in TER
(when acid secretion is inhibited with cytochalasin D) offsets the
decline in TER (when paracellular permeability increases in the
presence of cytochalasin D). Alternatively, H+ transport
may be inhibited in the presence of cytochalasin D without a
concomitant decrease in acid (secretion)-mediated Cl
and
Na+ flux. Further experiments will be required to
understand, in detail, the effects of cytochalasin D on TER in the
stimulated gastric mucosa. In addition, studies to determine the
properties of tight junctions in inhibited and stimulated tissues will
be required to verify that permeability is similar in both conditions.
Although cytochalasin D influences mucosal permeability and TER in this study, we show that there are no gross alterations in tight junction structure as determined by electron microscopy. These results are consistent with studies by Unno, et al, (29) who showed that ATP depletion induced by hypoxia increased permeability and changed the localization of actin at the tight junction, but had no effect on tight junction structure as determined by electron microscopy. Madara and Daharmsathaphorn (20a) showed that strand counts at the tight junction, as determined by freeze fracture microscopy, are predictive of the TER and rate of passive ion flux. Cytochalasin D reduces the number of strands associated with the tight junction, in concert with a reduction in TER and an increase in paracellular permeability (20). Electron microscopic examination of the tight junction, however, has been beneficial to determine abnormalities in tight junction structure including contraction of the perijunctional actomyosin ring, an increase in perijunctional actin, and development of membrane dilatations (15, 21).
The results presented here show that the rate of mannitol flux is greater in inhibited than in stimulated tissues. Although the reason for this difference is not known, we propose that the "secretory flush," or the serosal to mucosal movement of water, makes it more difficult for ions to diffuse in the opposite direction (from mucosal to serosal) during acid secretion. Alternatively, there may be distinct differences in the structure of tight junctions in inhibited and stimulated tissues. Such differences could facilitate the regulation of ion permeability during acid secretion.
The mechanism by which actin filaments regulate gastric acid secretion is not known. Forte and colleagues (8, 25) suggest that actin filaments participate in vesicular trafficking and the translocation of tubulovesicular membranes to the apical surface after stimulation. Chew et al. (5) showed that signaling proteins required for acid secretion are associated with F-actin. Actin filaments are thought to participate in Ca2+ mobilization after stimulation (23) and/or may serve to organize ion exchangers after stimulation (1). Because little direct experimental evidence has been obtained to identify the way in which actin filaments regulate gastric acid secretion, this should be a priority for further investigation.
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ACKNOWLEDGEMENTS |
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The authors thank Dr. Jeffrey Matthews for helpful discussions, especially concerning the 3H-22Na flux studies.
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FOOTNOTES |
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This work was supported by the National Institute of Diabetes and Digestive and Kidney Diseases Grant R01-DK-15681 (to S. J. Hagen) and DK-34854 (to the Harvard Digestive Diseases Center). Dr. Al-Shaibani was supported by the Fulbright Scholar Program, United States Department of State, the Arabian Gulf University, Manama, Bahrain and the United Medical group, Riyadh, Saudi Arabia, represented by Omar Ali Babtain.
T. A. Abdul-Ghaffar Al-Shaibani's current address is: Arabian Gulf University, College of Medicine and Medical Sciences, PO Box 22979, Manama, Bahrain.
Address for reprint requests and other correspondence: S. J. Hagen, Dept. of Surgery, Beth Israel Deaconess Medical Center, Dana 805, 330 Brookline Ave., Boston, MA 02215 (E-mail: shagen{at}caregroup.harvard.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.
10.1152/ajpgi.00393.2001
Received 7 September 2001; accepted in final form 25 November 2001.
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