Regulation of acid secretion and paracellular permeability by F-actin in the bullfrog, Rana catesbeiana

Tarik A. Abdul-Ghaffar Al-Shaibani and Susan J. Hagen

Department of Surgery, Beth Israel Deaconess Medical Center, Boston, Massachusetts 02215


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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


    INTRODUCTION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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 beta -actin and basolateral actin consists of gamma -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.


    MATERIALS AND METHODS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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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<UP><SUB>4</SUB><SUP>2−</SUP></UP>, and 19.3 mannitol and continuously gassed with 100% O2. Luminal solutions were unbuffered and kept at pH 4.7 with a pH-stat device (Radiometer America, Cleveland, OH). Serosal surfaces (from here onward called "nutrient") were bathed with (in mM) 102.4 Na+, 4.0 K+, 0.8 Mg2+, 1.8 Ca2+, 91.4 Cl-, 0.8 SO<UP><SUB>4</SUB><SUP>2−</SUP></UP>, 0.8 H2PO<UP><SUB>4</SUB><SUP>2−</SUP></UP>, 10 glucose, 17.8 HCO<UP><SUB>3</SUB><SUP>−</SUP></UP> (pH 7.2) and continuously gassed with 95% O2-5% CO2. Acid secretion was measured using a Radiometer pH-stat device and calculated from the volume of 10 mM NaOH needed to titrate the nutrient solution to a constant pH of 4.7. Transepithelial resistance (TER) was calculated from Ohm's law using measurements of potential difference that were monitored continuously by KCl-saturated agar bridges connected via two calomel electrodes to a voltmeter.

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 · h-1 · 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 · h-1 · 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 · h-1 · 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.


    RESULTS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Cytochalasin D inhibits stimulated acid secretion. Acid secretion in the frog fundic mucosa attained a maximal rate of 6.68 ± 0.17 µeq · h-1 · 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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Fig. 1.   Effect of 0.1% DMSO or 20 µM cytochalasin D in 0.1% DMSO (CD/DMSO) on acid secretion and transepithelial resistance (TER) in the stimulated and inhibited bullfrog gastric mucosa. A and B: in tissues stimulated with histamine plus carbachol for 60 min, addition of CD/DMSO reduced acid secretion within 3 h (A) without causing a change in TER (B). C: in contrast to stimulated tissues, TER dropped significantly in inhibited tissues treated with CD/DMSO. Data were obtained from 12 experiments and are expressed as means ± SE.

Cytochalasin D decreases TER in inhibited tissues. Tissues inhibited with cimetidine had an acid secretion rate of 0.00 µeq · h-1 · 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 (×10-2) µM · h-1 · cm-2. Stimulated tissues treated with DMSO had a slower initial rate of mannitol flux [4.17 ± 0.32 (×10-2) µM · h-1 · cm-2], which increased to 7.15 ± 3.24 (×10-2) µM · h-1 · cm-2 by 90-240 min (Fig. 2). Stimulated tissues incubated with CD/DMSO for 60 min had a similar rate of mannitol flux [7.15 ± 0.91 (×10-2) µM · h-1 · cm-2]. In contrast, there was a significant increase in mannitol flux in both inhibited and stimulated tissues by 90-240 min after incubation with CD/DMSO (Fig. 2). In inhibited tissues, the rate of mannitol flux increased 282% to 32.32 ± 2.34 (×10-2) µM · h-1 · cm-2. In stimulated tissues, the rate of mannitol flux increased 239% to 20.92 ± 3.49 (×10-2) µM · h-1 · cm-2. In all cases, the rate of mannitol flux was greater in inhibited, compared with stimulated, tissues.


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Fig. 2.   Effect of 0.1% DMSO or 20 µM cytochalasin D in 0.1% DMSO (CD/DMSO) on the mucosal to serosal flux (JMS) of mannitol. Mannitol flux was comparable in inhibited tissues treated with DMSO and in inhibited tissues treated with CD/DMSO for 60 min. Results were similar for stimulated tissues. In contrast, mannitol flux increased significantly in tissues treated with CD/DMSO for >90 min. Note that the flux of mannitol was greater in inhibited tissues. Data were obtained from 8 experiments (inhibited) or 10 experiments (stimulated) and are expressed as means ± SE. *Significant difference (P < 0.01 in inhibited tissues and P < 0.005 in stimulated tissues) compared with control tissues (90-240 min) treated with DMSO.

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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Fig. 3.   Electron micrograph of a gastric gland from tissues inhibited with cimetidine and then incubated for 4 h with 20 µM CD in 0.1% DMSO. Note that incubation with CD/DMSO had only minimal effect on gland structure consisting of dilation of the gland lumen (L) and reduction in the number of surface folds along the apical surface (arrows) of oxynticopeptic cells (OP). Tight junctions (TJ) are present at the luminal surface and are similar to those in control tissues treated with DMSO. The area outlined with a box is featured in Fig. 4. LP, lamina propria. Original magnification ×3,750; bar, 5 µm.



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Fig. 4.   High magnification electron micrograph of the area boxed in Fig. 3. Note that the tight junctions appear normal by electron microscopy when tissues are treated for 4 h with 20 µM cytochalasin D in 0.1% DMSO. Membrane organization and dilatations (arrows) and perijunctional actin look similar to control tissues treated with DMSO. Original magnification, ×26,250; bar, 0.5 µm.

HCO<UP><SUB>3</SUB><SUP>−</SUP></UP> flux contributes little to luminal acid neutralization during acid secretion. To test the hypothesis that an increase in paracellular permeability would increase the serosal to mucosal HCO<UP><SUB>3</SUB><SUP>−</SUP></UP> flux and serve to neutralize luminal acid (Fig. 5), we measured the rate of HCO<UP><SUB>3</SUB><SUP>−</SUP></UP> flux in inhibited tissues treated with DMSO or with CD/DMSO. In tissues treated with DMSO, the serosal to mucosal HCO<UP><SUB>3</SUB><SUP>−</SUP></UP> flux was 0.14 ± 0.03 µeq · h-1 · cm -2 after 5 h. The serosal to mucosal HCO<UP><SUB>3</SUB><SUP>−</SUP></UP> flux in tissues treated with CD/DMSO was 0.34 ± 0.03 µeq · h-1 · cm -2 after 5 h, which was a significant (threefold) increase compared with control tissues (Fig. 5).


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Fig. 5.   Effect of 0.1% DMSO or 20 µM cytochalasin D in 0.1% DMSO (CD/DMSO) on the serosal to mucosal flux (JSM) of bicarbonate (HCO<UP><SUB>3</SUB><SUP>−</SUP></UP>) in the bullfrog gastric mucosa. HCO<UP><SUB>3</SUB><SUP>−</SUP></UP> flux increased significantly within 5 h in tissues treated with CD/DMSO. Anoxia (N2) for 30 min followed by reoxygenation for 60 min did not cause a significant change in HCO<UP><SUB>3</SUB><SUP>−</SUP></UP> flux in either group. These results demonstrate that the increase in HCO<UP><SUB>3</SUB><SUP>−</SUP></UP> flux in tissues treated with CD/DMSO is by passive paracellular diffusion. Data are from 8 experiments and represent means ± SE. *Significantly different (P < 0.01) compared with control tissues treated with DMSO.

To determine whether the mean increase in HCO<UP><SUB>3</SUB><SUP>−</SUP></UP> flux in CD/DMSO-treated tissues was due to active HCO<UP><SUB>3</SUB><SUP>−</SUP></UP> transport or by passive paracellular diffusion, anoxia (N2) was applied to the tissues for 30 min (Fig. 5). The rate of HCO<UP><SUB>3</SUB><SUP>−</SUP></UP> flux in tissues treated with CD/DMSO during anoxia was not significantly different from that of tissues in normal oxygen conditions (Fig. 5). In addition, reoxygenation of tissues treated with CD/DMSO resulted in no significant difference in HCO<UP><SUB>3</SUB><SUP>−</SUP></UP> transport (Fig. 5). Anoxia followed by reoxygenation also had no significant effect on control tissues treated with DMSO. These results show that bicarbonate transport occurred by passive paracellular diffusion in inhibited tissues treated with DMSO or with CD/DMSO.

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 × 10-5 cm2/s) × 102.4 mM Na+/Dman (0.677 × 10-5 cm2/s) × 19.3 mM mannitol, is 10.46. Because the calculated ratio of 10.63 is nearly identical to the ratio due to diffusion alone (10.46), these results verify that the increase in ion permeability, in the presence of CD/DMSO, is by passive diffusion through the paracellular space.


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Fig. 6.   Na+ back diffusion predicted from the rate of mucosal-serosal Na+ and mannitol flux across the paracellular space. A: frog fundic mucosa was inhibited with omeprazole and then switched to closed-circuit conditions in the presence of 0.1% DMSO (DMSO) or 20 µM CD in 0.1% DMSO (CD/DMSO). Na+ and mannitol fluxes were determined simultaneously with dual tracer techniques. Data fit a line with a slope of 10.63 (r2 = 0.91), which indicates that the increase in ion permeability during treatment with CD/DMSO is by passive diffusion across the paracellular space. B: the movement of Na+ by diffusion in free solution across the paracellular space was calculated using mannitol flux data in Fig. 6A. The slope of 10.46 corresponds to the ratio of the free-solution diffusion coefficients of each species corrected for the concentration in the luminal solution. The data for mannitol was plotted with a slope of 10.46 to determine the predicted Na+ concentration. C: the measured rates of Na+ flux in 6 experiments with DMSO and 7 experiments with CD/DMSO (from Fig. 6A) were averaged. In the presence of CD, there is a significant increase in Na+ flux across the paracellular space. D: the predicted rates of Na+ flux in 6 experiments with DMSO and 7 experiments with CD/DMSO (from Fig. 6B) were averaged. The results show that a significant increase in Na+ flux across the paracellular space would be predicted to occur in the presence of CD/DMSO. The data represent means ± SE. *Significant difference (P < 0.005).

Using data from the known rate of mannitol flux in Fig. 6A, it is possible to calculate the rate of Na+ flux that would be predicted to occur by diffusion alone. This would be done by plotting the data for mannitol flux (from Fig. 6A) with a slope of 10.46 (ratio of the free-diffusion coefficients as calculated above) and extrapolating the predicted Na+ concentration on the y-axis (Fig. 6B). Because there were six experiments done with DMSO-treated tissues and seven experiments done with cytochalasin D-treated tissues, the data were extrapolated and then averaged for each group. Using extrapolated data, the mean rate of Na+ flux with DMSO is 0.519 ± 0.097 µeq · h-1 · cm-2 (Fig. 6D). This rate is not significantly different from the measured rate of Na+ flux with DMSO, which is 0.669 ± 0.075 µeq · h-1 · cm-2 (Fig. 6C). Likewise, the predicted rate of Na+ flux in the presence of CD/DMSO (using extrapolated data from Fig. 6B) is 2.918 ± 0.214 µeq · h-1 · cm-2 (Fig. 6D). This value is not significantly different from the measured rate of Na+ flux with CD/DMSO, which is 3.08 ± 0.327 µeq · h-1 · cm-2 (Fig. 6C). Thus equivalent results can be obtained concerning the movement of Na+ across the tight junction either experimentally (by measuring ion flux directly) or by predicting the movement of Na+ from its diffusion coefficient in free solution.

We used the above strategy to predict the rate of H+ diffusion through the paracellular space. The ratio of H+ to mannitol was calculated, using the H+ concentration in the luminal solution that, when titrated to pH 4.7 during acid secretion, is 10-4.7/M (or 1.99 × 10-5/M). The ratio, DH (9.311 × 10-5 · cm2 · s) × 0.0199 mM H+/Dman (0.677 × 10-5 · cm2 · s) × 19.3 mM mannitol, is 0.0142. We plotted the data for mannitol flux in Fig. 6A with a slope of 0.0142 and then extrapolated values for H+ flux from the y-axis (Fig. 7A). Because there were six experiments done with DMSO- and seven experiments with cytochalasin D-treated tissues, the data were extrapolated and then averaged for each group. The predicted mean rate of H+ flux was 12.5 ± 4.9 (×10-4) µeq · h-1 · cm-2 in the presence of DMSO and 39.6 ± 2.91 (×10-4) µeq · h-1 · cm-2 in the presence of CD/DMSO (Fig. 7B). It should be noted that mannitol flux in inhibited tissues is significantly greater than in stimulated tissues (Fig. 2). Thus our predicted rates of H+ flux through the paracellular space represent an overestimate of the true rate of H+ flux in stimulated tissues.


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Fig. 7.   H+ back diffusion predicted from the rate of mucosal-serosal H+ and mannitol flux across the paracellular space. A: the slope of 0.0142 corresponds to the ratio of the free-solution diffusion coefficients for H+ and mannitol corrected for the concentration in the luminal solution. Data for mannitol flux (Fig. 6A) were plotted with a slope of 0.0142 to determine the predicted H+ concentration. B: predicted rates of H+ flux in 6 experiments with DMSO and 7 experiments with CD/DMSO (from Fig. 7A) were averaged. Results show that a significant increase in H+ flux across the paracellular space would be predicted to occur in the presence of CD/DMSO. Rate of H+ flux is 12.5 ± 4.9 (×10-4) µeq · h-1 · cm-2 in tissues treated with DMSO and 39.6 ± 2.91 (×10-4) µeq · h-1 · cm-2 in tissues treated with CD/DMSO. The data represent means ± SE. *Significant difference (P < 0.005).


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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 · h-1 · 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<UP><SUB>3</SUB><SUP>−</SUP></UP> (serosal to mucosal flux) or by back diffusion of H+. In fact, the cytochalasin D-induced increase in HCO<UP><SUB>3</SUB><SUP>−</SUP></UP> flux would reduce acid secretion (by 0.34 ± 0.03 µeq · h-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 · h-1 · 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.


    ACKNOWLEDGEMENTS

The authors thank Dr. Jeffrey Matthews for helpful discussions, especially concerning the 3H-22Na flux studies.


    FOOTNOTES

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.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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Am J Physiol Gastrointest Liver Physiol 282(3):G519-G526
0193-1857/02 $5.00 Copyright © 2002 the American Physiological Society




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