State of actin in gastric parietal cells
John G.
Forte,
Bernice
Ly,
Qinfen
Rong,
Shoji
Ogihara,
Marlon
Ramilo,
Brian
Agnew, and
Xuebiao
Yao
Department of Molecular and Cell Biology, University of California,
Berkeley, California 94720
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ABSTRACT |
Remodeling of the
apical membrane-cytoskeleton has been suggested to occur when gastric
parietal cells are stimulated to secrete HCl. The present experiments
assayed the relative amounts of F-actin and G-actin in gastric glands
and parietal cells, as well as the changes in the state of actin on
stimulation. Glands and cells were treated with a Nonidet P-40
extraction buffer for separation into detergent-soluble (supernatant)
and detergent-insoluble (pellet) pools. Two actin assays were used to
quantitate actin: the deoxyribonuclease I binding assay to measure
G-actin and F-actin content in the two pools and a simple Western blot
assay to quantitate the relative amounts of actin in the pools.
Functional secretory responsiveness was assayed by aminopyrine
accumulation. About 5% of the total parietal cell protein is actin,
with about 90% of the actin present as F-actin. Stimulation of acid
secretion resulted in no measurable change in the relative amounts of
G-actin and cytoskeletal F-actin. Treatment of gastric glands with
cytochalasin D inhibited acid secretion and resulted in a decrease in
F-actin and an increase in G-actin. No inhibition of parietal cell
secretion was observed when phalloidin was used to stabilize actin
filaments. These data are consistent with the hypothesis that
microfilamentous actin is essential for membrane recruitment underlying
parietal cell secretion. Although the experiments do not eliminate the
importance of rapid exchange between G- and F-actin for the secretory
process, the parietal cell maintains actin in a highly polymerized
state, and no measurable changes in the steady-state ratio of G-actin to F-actin are associated with stimulation to secrete acid.
hydrogen-potassium-adenosinetriphosphatase; cytoskeleton; secretion; phalloidin; cytochalasin D; membrane recruitment
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INTRODUCTION |
THE CYTOSKELETAL SYSTEMS, either actin-based
microfilaments or tubulin-based microtubules, maintain fundamental cell
structure and have been suggested to play a key role in the trafficking of membranes, especially in the regulated recruitment and turnover of
membranes associated with a wide variety of transport processes, e.g.,
pancreatic insulin release (29), antidiuretic hormone (ADH)-regulated
water secretion (30), chromaffin granule secretion (31), and gastric
HCl secretion (6, 21). For several of these regulated vesicular
transport processes, a meshwork of filamentous actin (F-actin) in the
cell cortex just beneath the plasma membrane apparently represents a
barrier for vesicles that are to be recruited to the surface, and
access of vesicles to the target plasma membrane is proposed to occur
through regulated depolymerization of F-actin filaments to the
monomeric, G-actin, form (4, 18, 31, 36). Indeed, the continuous, rapid
interconversion of F-actin and G-actin, carefully controlled by a host
of actin-binding proteins, is a hallmark of cytoskeletal remodeling
associated with mobility of cells. On the other hand, stable actin
filaments are essential to vesicular traffic in a number of systems.
Microfilamentous cores of intestinal microvilli have been proposed to
represent "rails" on which membrane vesicles might be conveyed to
the surface by molecular motor proteins, such as myosin I (13).
Moreover, myosin I and the unconventional myosins have been implicated
as the protein motors to facilitate the cytoplasmic streaming of organelles along actin bundles in algal cells (1, 2). It is not
unreasonable to predict that a volatile F-actin pool (to afford
membrane-membrane interaction) and a stable F-actin pool (to provide
conveyance and microvillar support) may cooperate in regulated
vesicular trafficking.
Regulated secretion of HCl by the gastric parietal cell involves an
elaborate surface remodeling as cytoplasmic vesicles containing the
proton pump,
H+-K+-ATPase,
are recruited to the apical surface resulting in the transition from
short to long microvillar extensions (14, 15). The observed inhibition
of secretion by microfilament and microtubule inhibitors has been taken
as suggestive evidence that the cytoskeleton plays a central role in
the apical membrane remodeling associated with stimulation of the
parietal cell (6, 21). Because of the close relationship of actin
microfilaments to the extended secretory microvilli and the implied
importance of the cytoskeleton to parietal cell secretion, we sought a
more direct test of the state of actin associated with secretory
activity. The purpose of the present experiments was to assay the
relative amounts of F- and G-actin in gastric glands and parietal cells
and to determine the changes in the state of actin when parietal cells
were stimulated to secrete HCl. Accordingly, we used the
deoxyribonuclease (DNase) I binding assay to measure G-actin and total
actin content before and after isolated gastric glands or parietal
cells were stimulated by histamine. We also used a simple Western blot
assay to quantitate the relative amounts of actin in the
detergent-soluble and detergent-insoluble pools taken from the glands
and cells in resting and secreting states. For all experiments, the
functional secretory responsiveness was assayed by the ability to
accumulate weak base within an acidic space (aminopyrine accumulation
assay). Changes in the secretory response and state of gastric
glandular actin were also measured after applying agents that
specifically alter the state of the actin cytoskeleton. The experiments
show that parietal cells are rich in actin, especially in the F-actin
form (~90% of total actin), and that there are no significant
changes in the steady-state levels of F-actin and G-actin when the the
cells are stimulated to secrete acid. Cytochalasin D promoted a
depolymerization of F-actin to G-actin and inhibited acid secretion,
whereas stabilization of actin filaments using phalloidin did not
inhibit acid secretion.
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METHODS |
Isolation of gastric glands.
Gastric glands were isolated from New Zealand White rabbits essentially
as described by Berglindh (5) and more recently described by Yao et al.
(39). Isolated gastric glands were suspended in minimal essential
medium (MEM) and maintained in the resting state with
10
5 M cimetidine until
secretagogues were added, as described for individual experiments.
Isolation and separation of glandular epithelial cells.
Cells were prepared by a modified method described by Chew and Brown
(11). Gastric mucosa from perfused stomach was digested for 15 min at
37°C with pronase (0.5 mg/ml) in MEM containing 1 mg/ml bovine
serum albumin (BSA; fraction V) and 2 mg/ml glucose. The partially
digested mucosa was then subjected to three washes of MEM and incubated
for an additional 30-40 min in the same incubation medium with 0.8 mg/ml collagenase (Sigma; ~350 U/mg) After digestion with
collagenase, cells were washed three times in a medium containing (in
mM) 114.4 NaCl, 5.4 KCl, 5.0 Na2HPO4,
1 NaH2PO4,
1.2 MgSO4, 1 CaCl2, 0.5 dithiothreitol (DTT),
10 glucose, 1 pyruvate, and 10 N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic
acid (HEPES; pH 7.4), as well as 2 mg/ml BSA and 10 mg/l phenol red.
Cells were then resuspended in the same medium containing 10 mg/ml BSA, placed on nycodenz gradients (see below) and centrifuged at room temperature in a swinging bucket rotor at 1,000 g for 8 min. The top interface from
the gradient was collected as the parietal cell-rich fraction, which
was characterized at a purity of 85-90% by mitochondrial staining
with nitrotetrazolium blue (5). A mixture of chief cells and mucous
neck cells were in the lowermost band. Nycodenz gradients were prepared
as follows. Nycodenz sterile isotonic solution (density 1.15) was
supplemented with 10 mg/ml BSA and (in mM) 0.5 DTT, 2.4 KCl, 1.2 MgSO4, and 15 HEPES [pH 7.4, adjusted with 0.1 M tris(hydroxymethyl)aminomethane (Tris)]. This
solution was diluted at ratios of 2:1, 1:1, and 1:2 with an isotonic
solution containing (in mM) 132 NaCl, 5.4 KCl, 1.2 MgSO4, 0.5 DTT, and 15 HEPES (pH
7.4) as well as 10 mg/ml BSA and was sequentially layered into
centrifuge tubes.
Isolation of white blood cells.
Blood (~20 ml) was withdrawn from a rabbit into a heparinized
syringe, and the cells were harvested and washed three times with
phosphate-buffered saline (PBS; in mM: 149.6 NaCl, 3 K2HPO4, 0.6 NaH2PO4,
1 MgSO4, and 1 CaCl2) by centrifugation at
2,000 g for 15 min. Cells were
resuspended in PBS and placed on top of the same nycodenz gradients
described for parietal cell isolation, followed by centrifugation at
2,500 g for 10 min in a swinging bucket rotor (see above). The uppermost band at the first gradient interface was collected as white blood cells.
Permeabilization with
-toxin.
In cases in which phalloidin was added to stabilize F-actin filaments,
cells were permeabilized with
-toxin, which renders the cells
permeable to molecules in the range of 1 kDa while maintaining 60-80% of the functional secretory responsiveness (35, 38). Before treatment with
-toxin, intact glands or cells were washed once in a K+-rich permeabilization
medium (K medium) including (in mM) 20 HEPES, 10 Tris (pH 7.4), 100 KCl, 20 NaCl, 1.2 MgSO4, 1 NaH2PO4, and 40 mannitol. Glands or parietal cells were resuspended in K medium
at a 10% cytocrit and incubated with
-toxin at 37°C for 45 min
in the case of glands and 20 min in the case of isolated cells.
Cimetidine (100 µM) was also included in this and all subsequent steps to antagonize endogenous histamine that might be released. After
permeabilization, the suspensions were diluted to a 5% cytocrit in K
medium to which 10 mM succinate and 1 mM pyruvate were included as
oxidative substrates. The extent of parietal cell permeabilization was
routinely estimated by trypan blue uptake as described by Thibodeau et
al. (35).
Stimulation of parietal cells and measurement of aminopyrine
accumulation.
[14C]aminopyrine was
added to the cellular suspension in either MEM (intact glands and
cells) or K medium (permeabilized cells) to a final concentration of 4 × 10
4 mM. Aliquots
(0.5 ml) of the suspension were distributed into preweighed 1.5-ml
tubes already containing secretagogues or activators as specified in
each individual experiment. The performance of and calculations for the
aminopyrine uptake assay were essentially as described by Yao et al.
(39). Intact preparations were stimulated with histamine (100 µM) and
3-isobutyl-1-methylxanthine (IBMX; 50 µM).
-Toxin-permeabilized
cells were stimulated by adding 0.1 mM adenosine
3',5'-cyclic monophosphate (cAMP) and 1 mM ATP to the K
medium (35).
Detergent extraction of glands and cells.
Detergent-soluble and detergent-insoluble fractions were prepared as
follows. All steps were performed at room temperature. Typically, a
gland or cell pellet was treated with a 20-fold volume of extraction
buffer (usually ~500 µl) containing 0.1% Nonidet P-40 (NP-40), 5 mM
KH2PO4,
27 mM
Na2HPO4
(pH 7.2), 2 mM MgSO4, 2 mM
ethylene glycol-bis(
-aminoethyl
ether)-N, N, N', N'-tetraacetic acid, 0.2 mM ATP, 0.5 mM DTT, 2 M glycerol, and 1 mM
phenylmethylsulfonyl fluoride (a protease inhibitor) for 15 min with
agitation. The preparations were then centrifuged (400 g) for 1 min and separated into a
soluble fraction (NP-40 supernatant) and an insoluble cytoskeletal fraction (NP-40 pellet). Preliminary experiments were carried out to
optimize the time of extraction. To minimize interconversion between
actin forms, the NP-40 supernatant extracts were assayed for G-actin
within 1-5 min of cell extraction. The NP-40 cytoskeletal pellets
were resuspended in ~500 µl of extraction buffer and subsequently assayed for G-actin and total actin. Because our preliminary tests showed virtually no G-actin remaining in the NP-40 pellets, assays on
this fraction were delayed until the NP-40 supernatants were completed.
DNase I assay of G-actin and total actin.
The assay of G-actin and total actin was essentially according to the
DNase I inhibition assay (8) with modifications described by Heacock
and Bamburg (19). The DNA solution consisted of 80 µg/ml DNA (type
XIV from herring testes, Sigma Chemical, D6898) in 0.1 M
Tris · HCl, 4 mM
MgSO4, and 1.8 mM
CaCl2 (pH 7.5). The DNase I stock
solution was made up as 10 mg/ml DNase I (bovine pancreas, Sigma
Chemical, DN-25) in 0.125 M Tris · HCl, 5 mM
MgCl2, 2 mM
CaCl2, and 1 mM
NaN3 (pH 7.5) and stored at
20°C. For the DNase working solution, the stock DNase was
diluted 1:100 in 20 mM imidazole, 30 mM NaCl, and 15% glycerol (pH
7.0) and maintained on ice until use.
Assays for G-actin were carried out at room temperature by adding 20 µl of DNase I working solution and the desired amount of unknown
sample, or standard G-actin, to 3 ml of DNA solution in a quartz
cuvette and immediately measuring the rate of DNA hydrolysis at 260 nm
in a Varian spectrophotometer. The resulting curve for d(optical
density)/dt has three
portions: a slow initial rate and a major linear portion, followed by a
saturating portion. The slope of the linear portion of the curve was
inversely proportional to the concentration of G-actin as determined by
standard curves constructed with purified G-actin (the higher the
concentration of G-actin, the greater the inhibition of DNA hydrolysis
rate). The volume of unknown samples added to the cuvette was adjusted so that the slopes fell within the linear range, and assays were always
carried out in duplicate or triplicate.
To determine total actin, standards or unknown samples were treated on
ice for 15 min with an equal volume of guanidine
hydrochloride solution to depolymerize F-actin. The
depolymerizing solution contained 1.5 M guanidine hydrochloride, 1 M
sodium acetate, 1 mM CaCl2, 1 mM
ATP, and 20 mM Tris · HCl (pH 7.5). After a brief centrifugation to remove remnant nuclear material, aliquots were combined in a cuvette with DNase I and DNA solutions for actin assay as
described above. F-actin was calculated as the difference between total
actin and initially measured G-actin.
Actin measurements by Western blotting.
In some cases total actin content in the NP-40 supernatant and pellet
was measured by running aliquots on sodium dodecyl
sulfate-polyacrylamide gel electrophoresis (12% running gel), blotting
the protein to nitrocellulose, and probing for actin using a commercial
actin antibody (Amersham Life Science). The blots were developed with a
second antibody coupled to horseradish peroxidase, and the density of
the signal was quantitated by digital imaging using National Institutes
of Health Image software. At least three graded samples of both
NP-40 supernatant and pellet material were applied to the lanes
of a single gel, along with a series of four to six actin standards
ranging from 5 to 50 ng actin. In this way we established that
the unknown samples would be bracketed by standards and in a linear
range for every experiment and blot.
Protein assay.
The Bradford assay was used to measure protein for all cell extracts
and pellets and for actin standards.
 |
RESULTS |
Time course of protein extraction.
To establish an appropriate time of extraction, we evaluated the time
course of treatment with NP-40 extraction buffer at room temperature,
followed by separation into supernatant and pellet fractions. Figure
1 shows the time course of
protein and actin extraction from isolated parietal cells. Protein was
rapidly released into the supernatant (~45% of total protein was
released within 1 min) and reached a steady state by 10-15 min, at
which time ~80% of total cell protein was in the supernatant and
20% in the residual pellet (Fig.
1A). The relative amounts of G-
and F-actin in the supernatant and pellet fractions, as measured by the
DNase 1 assay, are shown in Fig. 1B.
The majority of parietal cell actin remained with the pellet throughout
the extraction, and most of the pellet actin was in the F-actin form.
The supernatant contained both G- and F-actin. G-actin increased
initially and leveled off by 10 min. A small amount of G-actin was
initially present in the pellet, and this decreased so that by 15 min
there was no measurable G-actin in the pellet. The combined G-actin, supernatant plus pellet, remained constant at ~10% of total parietal cell actin throughout the time of extraction. The F-actin in the supernatant showed a slight decline over the first 5-10 min of extraction. Because the total G-actin-to-F-actin ratio remained constant over the time course of extraction, it appeared that the
extraction conditions were stable. Also, on the basis of these experiments we decided to treat cells and glands with extraction buffer
for 15 min as a standard protocol of extraction.

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Fig. 1.
Time course for treatment of isolated parietal cells with NP-40
extraction buffer. After treatment, samples were separated into
supernatant and pellet fractions as described in
METHODS and protein was assayed by
Bradford assay. A: values are % of
total protein extracted into supernatant fraction (means ± SE); for
5, 10, and 15 min, n = 3; for all
other time points, n = 2. B: values for actin in 1 set of
samples. G-actin (G) and total actin were measured by deoxyribonuclease
I assay; F-actin (F) is reported as total actin G-actin. Sup,
supernatant; Pel, pellet.
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Steady state of actin in gastric glands, parietal cells, and other
cell types.
Experiments were designed to measure the steady-state levels of G-actin
and F-actin in freshly isolated gastric glands and in partially
purified cell populations isolated from the glands. As indicated in
Table 1 for gastric glands, the NP-40
supernatant contained ~30-35% of the total actin, which
included both G-actin and F-actin; the NP-40-insoluble cytoskeletal
fraction contained the largest pool of actin (~64% of total),
recovered almost exclusively as F-actin. About 15% of the total
gastric glandular actin was measurable as G-actin; the remaining 85%
was measured only after guanidine treatment and therefore taken to be
F-actin. The data for isolated cells, also depicted in Table 1,
indicate that the more highly purified parietal cell populations
contained a higher proportion of F-actin than either the parietal
cell-poor fractions or the intact gastric glands. In isolated cell
fractions, judged to be 85-91% parietal cells, F-actin
represented ~90% of the total actin, and most of this was found in
the NP-40-insoluble cytoskeletal fraction. On the other hand, cell
fractions relatively poor in parietal cells contained less total actin,
which in turn was somewhat more skewed toward the G-actin form (i.e.,
~80% of total actin occurred as F-actin).
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Table 1.
Relative distribution of G-actin and F-actin in extracts and pellets
taken from several cellular sources
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Concerns have been raised regarding methodology that may lead to
inaccurate measurements of G-actin and F-actin in cells (19). The high
percentage of F-actin in gastric glands, parietal cells, and even
gastric nonparietal cells could be the result of polymerization of
G-actin induced by the NP-40 extraction buffer. To test this hypothesis, white blood cells, which contain relatively high
proportions of G-actin (24, 25, 33), were isolated and extracted with the same NP-40 extraction buffer and conditions used for the gastric samples. The last column in Table 1 illustrates the distribution of
actin in white blood cells. About one-half of the total actin in the
white blood cells was measured as G-actin in the NP-40 supernatant;
relatively small amounts of F-actin (~7% of total) were extracted
into the NP-40 supernatant (Table 1). These results support the notion
that the high F-actin percentage is not an artificial state induced by
the NP-40 extraction buffer, and they demonstrate that parietal cells
belong to the class of cells in which the distribution of actin is very
heavily skewed toward the filamentous cytoskeletal form.
Measurements of actin in resting and stimulated gastric glands.
We next sought to evaluate the steady-state levels of actin forms in
gastric glands and parietal cells during various states of secretory
activity. Table 2 shows actin
and protein measured in freshly isolated gastric glands maintained in a
resting state compared with glands stimulated with secretagogues to
secrete acid. Stimulated glands demonstrated good secretory
responsiveness, showing five- to sixfold increases in
[14C]aminopyrine
uptake ratios over the resting counterparts. However, there were no
significant differences between the resting and stimulated glands for
the steady-state distribution of actin in the NP-40 supernatant and
pellet or in the distribution of G- and F-actin in those fractions. For
both resting and stimulated preparations, actin represents ~5% of
the total glandular protein, and this was predominantly in the F-actin
form. As with previous results, G-actin comprised only ~10% of the
total actin.
An experimental time course of stimulation and inhibition is shown in
Fig. 2. Gastric glands were treated with
histamine for 20 min and then exposed to a powerful
histamine-H2 blocker, SKF-9347 (7), to restore the glands to a resting state. Aliquots of glands were
taken at various times over the treatment regimen and assayed for G-
and F-actin in the NP-40 supernatant and total actin in the pellet.
Because there was typically negligible G-actin in the pellet, total
pellet actin was assumed to be equivalent to pellet F-actin. Again
there were no significant differences in the steady-state distribution
of actin forms between resting glands and those stimulated to secrete
acid. Over the 60 min after acid secretion was inhibited by SKF-9347,
there was a slight decrease in the amount of F-actin that sedimented
with the NP-40 pellet, but this reached the 5% level of significance
only for the preparations at 30 min postinhibition.

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Fig. 2.
Measurement of state of actin in gastric glands over time course of
stimulation to secrete acid and subsequent inhibition. Freshly isolated
gastric glands were sequentially treated with
10 4 M cimetidine for 20 min, 10 4 M histamine for 20 min, and 5 × 10 5 M
SKF-9347 (H2 antagonist) for 60 min. At indicated time points, separate samples were taken for
aminopyrine uptake assay (A) and for
assay of actin (B). For actin
analyses, glands were extracted with NP-40 buffer, separated into
supernatant and pellet, and assayed for G- and F-actin as described in
METHODS. Values are means ± SE;
no. of samples at each time point for aminopyrine
(A) or actin
(B) analysis is given in
parentheses.
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A summary of experiments for resting and stimulated parietal cells is
shown in Table 3. As was the case for the
gastric glands, there were no significant differences between resting
and stimulated cells in the steady-state levels and distribution of
actin forms.
Cytochalasin D produces secretory changes and changes in the state
of gastric glandular actin.
Figure 3 shows that treatment of gastric
glands with cytochalasin D disrupted actin filaments, as judged by a
significant decrease in the NP-40-insoluble cytoskeletal F-actin pool
and an increase in G-actin content. In addition, there was a slight increase in the F-actin in the NP-40 supernatant. These changes in the
relative distribution of G- and F-actin occurred at both 1 and 10 µM
cytochalasin. Aminopyrine accumulation in response to histamine plus
IBMX was also significantly depressed in these same
cytochalasin-treated glands, consistent with earlier studies showing
inhibition of acid secretion in functionally intact gastric mucosa (6).

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Fig. 3.
Cytochalasin D (CD) inhibits glandular acid secretion and disrupts
F-actin filaments. Isolated gastric glands were treated for 20 min at
37°C to sustain resting conditions (rest.;
10 4 M cimetidine) or
stimulated conditions (stim.;
10 4 M histamine + 5 × 10 5 M
3-isobutyl-1-methylxanthine). Cytochalasin D was also added to
stimulated gland preparations at concentrations indicated. After
incubation, separate samples were taken for aminopyrine uptake assay
(A) and for assay of G- and F-actin
in supernatant and pellet fractions
(B) as described in
METHODS. Values are means ± SE;
n = 3.
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Stabilization of actin filaments by phalloidin does not alter the
secretory response.
To ensure ready access of gastric cells to phalloidin, glands or
parietal cells were permeabilized with
-toxin, a model that we
previously showed to be functionally responsive to cAMP. We used
rhodamine-labeled phalloidin to visualize the ready permeation of
phalloidin into parietal cells (not shown). Treatment of
-toxin-permeabilized gastric glands with 1 or 10 µM
phalloidin did not interfere with their ability to respond to cAMP as a
stimulant (Fig. 4). Initial experiments
employing the DNase I assay to measure G- and F-actin in
phalloidin-treated preparations met with difficulty in that high
concentrations of guanidine were necessary to disrupt the phalloidin-F-actin association and depolymerize actin for assay. To
avoid this difficulty, we used the alternative Western blot assay of
actin in the NP-40 supernatant and pellet fractions from gastric glands
and parietal cells. A major disadvantage is that the blotting method
does not distinguish between G- and F-actin but simply assays total
actin in the NP-40-soluble and -insoluble fractions.

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Fig. 4.
Phalloidin does not inhibit stimulation of acid secretion by
-toxin-permeabilized gastric glands. Glands were permeabilized with
-toxin as described in METHODS and
incubated for 20 min with
10 4 M cimetidine (resting)
or with 10 4 M cAMP + 10 3 M ATP (stimulated)
containing 0, 1, or 10 µM phalloidin as indicated. After 20 min,
glands were taken for
[14C]aminopyrine
uptake assay as an index of acid secretion. Values are means ± SE
for 3 separate preparations.
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Table 4 shows the results of aminopyrine
accumulation and actin measurements for
-toxin-permeabilized
parietal cells treated with 5 µM phalloidin in the resting and
stimulated states. For control cells, with no phalloidin added, the
results were similar to those obtained with the DNase I assay (cf.
Tables 2 and 3), with the exception that the blotting method indicated
proportionally less actin in the NP-40 supernatant (~15% of total)
than the DNase I assay (~33% of total). This difference between the
two methods was consistent over many experimental measurements, and we
have no explanation for the discrepancy. In any event, there were no changes in the distribution of actin in the NP-40-soluble and -insoluble fractions associated with stimulation of the parietal cells.
Treatment of parietal cells with 5 µM phalloidin produced a small
decrement in the NP-40 supernatant actin and a slight increase in
cytoskeletal actin, consistent with a stabilization of F-actin (Table
4). Similar to what was observed for gastric glands, phalloidin did not
inhibit the transition from rest to stimulation in
-toxin-permeabilized cells treated with cAMP.
 |
DISCUSSION |
From results presented here we draw the following conclusions. Parietal
cells are rich in actin, representing ~5% of the total protein, with
~90% of the actin present in the filamentous form (F-actin).
Treatment of gastric glands with cytochalasin D resulted in the
expected inhibition of acid secretion as well as a decrease in F-actin
and an increase in G-actin. On the other hand, hyperstabilization of
actin filaments with phalloidin produced no inhibition of parietal cell
secretion. Stimulation of acid secretion via the cAMP pathway resulted
in no measurable change in the relative amounts of G-actin, short
filamentous actin, and cytoskeletal F-actin. These data are consistent
with the hypothesis that microfilamentous actin is essential for
membrane recruitment underlying parietal cell secretion. However,
activation of secretion involves little or no change in steady-state
F-actin-to-G-actin transformation. This last conclusion was most
surprising in view of the rather profound apical membrane
transformations associated with stimulation and the evidence that comes
from several other secretory systems.
In stimulated adrenal chromaffin cells, disassembly of cortical actin
was found to precede catecholamine secretion and was correlated with
increased levels of cytosolic Ca2+
(10). Disassembly of actin was also observed in digitonin-permeabilized chromaffin cell actin treated with micromolar levels of
Ca2+ (9). Using immunogold
electron microscopy to double-label chromaffin granules and F-actin,
Nakata and Hirokawa (27) found that the cytoskeletal reorganization in
response to stimulation of cultured chromaffin cells was not massive
but appeared to involve discrete depolymerization of the cortical
cytoskeleton proximal to each exocytic site. In the case of
ADH-regulated water transport, rapid depolymerization of F-actin was
observed in toad bladder epithelial cells stimulated with ADH or
8-bromoadenosine 3'5'-cyclic monophosphate (12).
Concomitant with exocytosis and increased water transport, monomeric
G-actin increased from 37 to 54% of the total actin (18). In most
cells the concentration of G-actin greatly exceeds its critical
concentration for polymerization, indicating that G-actin-binding
proteins are required to sustain the steady state and suggesting that
regulation of the G-actin sequestration system may be fundamental to
the exocytic event. Confocal microscopy (20) and quantitative
immunogold techniques (15) were used to demonstrate that actin
depolymerization in ADH-treated toad bladder cells occurred only at the
apical pole, and that this cortical layer of actin selectively
depolymerized between microvilli but not within the microvilli per se.
On the other hand, net depolymerization of F-actin by low levels of
cytochalasin B produced no increase in aggrephore fusion or water flow
(37). To test whether the cortical actin cytoskeleton provides a
barrier for cholecystokinin (CCK)-regulated exocytosis in
pancreatic zymogen secretion, Muallem et al. (26) measured the release
of amylase from acinar cells permeabilized with streptolysin O so that
they could introduce
-thymosins as specific monomeric actin-binding proteins. Low concentrations of the actin monomer-sequestering peptides
triggered a rapid and robust exocytosis with a profile similar to the
initial phase of CCK stimulation, but high concentrations of the
polypeptides inhibited all phases of exocytosis, suggesting that some
basic cytoskeletal structure was essential for the secretory event.
O'Konski and Pandol (28) offered alternative evidence for a
functionally stable pool of F-actin, showing that
diminished secretory activity caused by hyperstimulating acinar cells
was associated with marked degradation of the apical cytoskeleton. Collectively, these studies are consistent with the hypothesis that
cortical F-actin impedes access for membrane recruitment and that some
transient and selective depolymerization must occur for secretion;
however, they also indicate that a minimal actin cytoarchitecture is
necessary for exocytosis, possibly acting as a structural support for
apical surface activity and/or a pathway for vesicular
transport.
Remodeling of the apical membrane-cytoskeleton has been implicated in
acid secretion by parietal cells. Although there is some disagreement
on the specific source of the expanded secretory surface (15, 32),
morphological evidence clearly demonstrates the conversion of short to
long microvilli when cells are stimulated. A major question is, How do
the microvilli grow: do they extend from the tips, or do they
"grow" into the cell? Related to this are the nature and dynamics
of the microfilament support system for the microvilli. Because of the
morphological arrangement within the restricted space of parietal cell
canaliculi, secretion-dependent microvillar growth would have to occur
into the cytoplasmic space and could not occur by tip elongation as
usually happens in the case of filapodia and microspikes or developing
intestinal microvilli. This view is consistent with the observation
that the bases of the microvilli extend deep into the cytoplasm of
maximally stimulated cells. Thus it seems reasonable to conclude that
microvillar elongation occurs by adding membrane at the microvillar
base, with consequent extension or "growth" into the cell. In
resting parietal cells, organized bundles of microfilaments have been
shown to extend into the cytoplasm 1 µm or more beneath the short
microvilli (6), potentially representing the structural framework for
elongation. However, it is not clear whether filament length is fixed
or whether they undergo regulated growth and extension to accommodate
elongated microvilli. Moreover, the polarity of the microfilamentous
bundles within parietal cell microvilli has not been determined. The
presumed action of membrane-cytoskeletal attachment proteins such as
ezrin or nonconventional myosins, both of which are strongly expressed in parietal cells (3, 17), would provide the bonding sites for shaping
the expanded membrane surface into elongated microvilli.
The present data indicate that parietal cell microvillar elongation
occurs without a major shift in the steady-state G-actin-to-F-actin ratio. This apparent intransigence of the steady-state actin forms does
not preclude the importance of actin turnover associated with secretory
activation. It is possible that the turnover between G- and F-actin may
be rapid and continuous, so that depolymerization from one pool is
compensated by polymerization into another pool within the time
resolution of our measurements. Alternatively, there
could a large pool of stable microfilamentous actin, for example,
associated with microvilli, and a smaller pool of cortical dynamic
actin between microvilli, whose depolymerization would be difficult to
detect in the large background of stable F-actin filaments. Both of
these general possibilities would allow for activation-dependent
cytoskeletal reorganization within an apparently constant
G-actin-to-F-actin ratio. On the other hand, the addition of phalloidin
to
-toxin-permeabilized parietal cells did not inhibit or prevent
the cAMP-mediated accumulation of aminopyrine by glands or parietal
cells, suggesting that stabilization of parietal cell F-actin did not
alter the response to stimulation. In this regard, phalloidin
stabilization of actin filaments in the T84 intestinal cell line did
not alter the activation and insertion of
Cl
channels at the apical
membrane, although cAMP-induced activation of the
Na+-K+-2Cl
cotransporter at the basolateral membrane appeared to be microfilament dependent and responsible for inhibitory effects in the
phalloidin-loaded state (22, 23). These observations suggest that
disorganization of the apical actin network may not be required for
regulated exocytic recruitment of transport proteins in all cases. In
fact, secretion of insulin by permeabilized pancreatic islet cells was reportedly enhanced by stabilizing F-actin with phalloidin (34).
In summary, the results of this study support the conclusion that
filamentous actin is important for initiating or maintaining the
regulated secretion of HCl by the parietal cell. The parietal cell
maintains actin in a highly polymerized state, with 90% of total actin
as F-actin, and there were no measurable changes in the steady-state
ratio of G-actin to F-actin associated with stimulation of parietal
cells to secrete acid. In addition, treatment of
-toxin-permeabilized parietal cells with the F-actin stabilizing
agent phalloidin did not inhibit secretory activity. Although these
latter data are consistent with a functional role for a cytoskeletal
framework in parietal cell activation, they do not eliminate the
importance of the rapid recycling of actin between the F- and G-actin
forms as secretion and recovery from secretory activity occur. More dynamic measurements of actin turnover and cytolocalization of the
dynamic actin pools will be necessary to address the recycling issue.
 |
ACKNOWLEDGEMENTS |
This work was supported in part by National Institute of Diabetes
and Digestive and Kidney Diseases Grant DK-10141.
 |
FOOTNOTES |
Address for reprint requests: J. G. Forte, Dept. of Molecular and Cell
Biology, 241 LSA, University of California, Berkeley, CA 94720-3200.
Received 4 August 1997; accepted in final form 9 September 1997.
 |
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