Division of Gastroenterology/Hepatology, University of Colorado Health Sciences Center, Denver, Colorado 80262
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The present studies of cholangiocytes used complementary histological, biochemical, and electrophysiological methods to identify a dense population of subapical vesicles, quantify the rates of vesicular trafficking, and assess the contribution of the actin cytoskeleton to membrane trafficking. FM 1-43 fluorescence measured significant basal rates of total exocytosis (1.33 ± 0.16% plasma membrane/min) in isolated cholangiocytes and apical exocytosis in cholangiocyte monolayers. Cell surface area remained unchanged, indicating that there was a concurrent, equivalent rate of endocytosis. FM 1-43 washout studies showed that 36% of the endocytosed membrane was recycled to the plasma membrane. 8-(4-Chlorophenylthio)adenosine 3',5'-cyclic monophosphate (CPT-cAMP; cAMP analog) increased exocytosis by 71 ± 31%, whereas the Rp diastereomer of adenosine 3',5'-cyclic monophosphothioate (Rp-cAMPS; protein kinase A inhibitor) diminished basal exocytosis by 53 ± 11%. A dense population of 140-nm subapical vesicles arose, in part, from apical membrane endocytosis. Phalloidin staining showed that a subpopulation of the endocytosed vesicles was encapsulated by F-actin. Furthermore, membrane trafficking was inhibited by disrupting the actin cytoskeleton with cytochalasin D (51 ± 13% of control) or jasplakinolide (58 ± 9% of control). These studies indicate that there is a high rate of vesicular trafficking at the apical membrane of cholangiocytes and suggest that both cAMP and the actin cytoskeleton contribute importantly to these events.
exocytosis; endocytosis; vesicular trafficking
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
INTRAHEPATIC BILE DUCTS are lined by an active absorptive-secretory epithelium. Many of the transporter and channel proteins responsible for the movements of ions, solutes, and water across this epithelium have been identified, but the underlying mechanisms responsible for their coordinated regulation are not well defined. In a number of cell types, trafficking of key proteins into and out of the membrane is a central mechanism for controlling transport activity. For example, acid secretion from gastric parietal cells and water permeability of the renal collecting duct are largely controlled through the insertion and retrieval of H+-K+-ATPase proton pumps and aquaporin-2 water channels, respectively (25, 26). In cholangiocytes, the cell type that lines intrahepatic bile ducts, recent studies support a similar paradigm. In intact liver, secretin exposure induces a decrease in the number of >200-nm vesicles and a parallel increase in plasma membrane surface area (4). Furthermore, secretin increases vesicular fluid secretion from acidic vesicle compartments and the water channel aquaporin-1 may be concurrently inserted into the apical membrane (16, 21). Consequently, the purpose of these studies was to evaluate quantitatively the rates of basal and regulated membrane trafficking and to identify a subapical membrane vesicle population that could account for the measured rates. Specific emphasis was placed on evaluating the potential contributions of the actin cytoskeleton to membrane trafficking events. The findings provide direct evidence for robust membrane trafficking of subapical vesicles that are in communication with the apical membrane and are regulated by both cAMP- and actin-dependent pathways.
![]() |
METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Cholangiocyte cell lines and culture.
Two established cholangiocyte models, Mz-ChA-1 (cholangiocarcinoma cell
line) and normal rat cholangiocytes (NRC) cells, were used in these
studies (18, 39). Cultured on culture-treated flasks (no.
430199; Corning, Corning, NY) as previously described (11), Mz-ChA-1 cells were used to investigate the
properties of isolated, nonpolarized cholangiocytes. To evaluate
exocytosis in polarized cells, NRC monolayers were cultured on rat tail
collagen slabs as previously described (29) and passaged
onto collagen-coated semipermeable (24-mm diameter, 0.4-µm pore)
Costar transwell supports (Corning) 7-10 days before all studies.
This protocol permits the development of a high transepithelial
resistance (Rt >1,000 · cm2), net apical HCO
Experimental agents. FM 1-43, jasplakinolide, and Texas red-phalloidin were obtained from Molecular Probes (Eugene, OR). CPT-cAMP and cytochalasin D were obtained from Sigma (St. Louis, MO). All other reagents were purchased from Sigma.
Quantitative microscopy of FM 1-43 fluorescence.
Isolated Mz-ChA-1 cells were used to measure the rate of exocytosis
with FM 1-43 fluorescence intensities (2, 6). FM 1-43 is a useful quantitative probe of membrane trafficking
because it rapidly (seconds) and reversibly partitions into membranes, is impermeant to lipid bilayers, and increases its quantum efficiency of fluorescence >300-fold in a lipid vs. aqueous environment. Mz-ChA-1 cells were grown on coverslips and perfused with either E-buffer (in mM: 142 NaCl, 4 KCl, 1 KH2PO4, 2 MgCl2, 1.5 CaCl2, 10 D-glucose, and
10 HEPES; pH 7.4; 295-300 mosmol/kgH2O) or E-buffer with 4.7 µM FM 1-43. Unless otherwise specified, all
studies were performed at room temperature. The cells were observed on
a Nikon microscope with a ×60/NA1.2 water-immersion lens. FM 1-43
was excited with band pass filters (peak 480 nm) and collected with an
emission filter (peak 535 nm). The experimental protocols were designed, executed, and captured with the TILLvisION v3.3 software package. During the experimental periods, fluorescence images were
taken from 100-ms exposures at 30-s intervals and captured with a
12-bit cooled charge-coupled device (CCD) IMAGO digital camera.
The pixel size was 0.165 µm. This level and rate of exposure has
negligible bleaching effects (17). NIH Image 6.0 was used to quantify the cellular and background fluorescence intensities from
the same fields in the captured series of images. After the addition of
FM 1-43, there is a rapid rise in fluorescence intensity as the
dye equilibrates with the plasma membrane. Background-corrected cellular fluorescence values were normalized to this initial peak value
(initial plasma membrane surface area; 100%).
Measurement of membrane capacitance.
Whole cell patch-clamp techniques were used to assess the effect of
control, CD (10 µM in EtOH), and Jas (50 nM in EtOH) treatments on
total membrane surface area of Mz-ChA-1 cells. Cells were maintained in
E-buffer and were dialyzed with a standard intracellular pipette solution (in mM: 130 KCl, 10 NaCl, 1 EGTA, 0.5 CaCl2,
and 10 HEPES, pH 7.25; 275 mosmol/kgH2O). After whole cell
configuration was achieved, the cells were voltage clamped at a holding
potential of 40 mV and depolarizing pulses of 4-ms duration were
applied (pulse amplitude 40 mV). Current responses were acquired with a
sampling time of 5 µs on a Macintosh computer with Pulse Control software (15) in conjunction with an ITC16 interface
(Instrutech, Great Neck, NY) and IgorPro3 (WaveMetrics, Lake Oswego,
OR). With custom software, the currents were averaged and inverted and
then fitted to the equation I(t) = Iss + (I0
Iss) exp(
t/
), where I(t) is current response,
Iss is steady-state current,
I0 is peak current, t is time, and
is time constant. From the fitted parameters (I0, Iss, and
), the
membrane capacitance (Cm), access resistance (Ra), and membrane conductance
(Gm) were calculated as previously described
(20). This procedure was repeated every 3 s. The
recordings were stopped if Gm exceeded 1 nS.
Preparation of subapical patches.
To isolate and observe membrane vesicles in the subapical domain of NRC
cells, an apical membrane patch preparation was developed (Fig.
1) with modifications of a previously
described protocol (30). For electron microscopic (EM)
studies, NRC monolayers were washed at 4°C [3× in PBS, 2× in
intracellular (IC) buffer (in mM: 25 HEPES, 25 KCl, and 2.5 Mg
acetate; pH 7.0)] and inverted over a coverslip holding
formvar/poly-L-lysine (1 mg/ml)-coated EM grids.
Pressure was applied to the filter while the residual medium was
aspirated from between the cells and the grids. The grids and adhering
apical membrane constituents were removed, washed with IC buffer, fixed
with 4% glutaraldehyde, and prepared for EM.
|
Microscopic analysis of apical membrane patches. The apical membrane patches were evaluated by EM and fluorescence microscopy. For EM analysis, positively stained and washed specimens were fixed with 1% OsO4 (15 min), washed, treated with 1% tannic acid (10 min), washed, stained with 1% aqueous uranyl acetate (UA), washed, and finally air dried. To provide negative contrast and highlight membranes, negatively stained specimens were prepared either 1) in 2% methylcellulose containing 0.3% UA or 2) in 1% neutralized phosphotungstic acid (PTA) (22). EM grids were examined on a Phillips CM10 electron microscope.
A Nikon microscope with a ×60/1.4 NA ApoPlan oil-emersion objective was used for fluorescence imaging. Images were captured on Kodak EliteII 400 color slide film, digitized on a Nikon LS1000 slide scanner, and analyzed with NIH Image 6.0. ![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Cholangiocytes exhibit high rate of apical exocytosis.
FM 1-43 fluorescence was used to quantify the rate of membrane
exocytosis as illustrated in Fig. 2.
Studies with Mz-ChA-1 cells permit quantification of membrane events at
the single-cell level and allow for greater experimental manipulation,
including exchange of solutions and addition of supplementary
compounds. Studies in NRC monolayers permit the selective apical
addition of FM 1-43 and observation of apical membrane events in a
functional epithelium.
|
Quantitation of exocytosis in cholangiocytes.
The change in fluorescence intensities permits quantitation of the
rates of exocytosis. After initial equilibration with the plasma
membrane (= 100%), the rate of FM 1-43 fluorescence intensity increase was fit by a linear equation corresponding to equilibration of
FM 1-43 with newly exocytosed membrane (Fig. 2B,
region 1). Values corresponded to an exocytic rate of
1.33 ± 0.16%PM/min (n = 8). To determine whether
this rate of exocytosis was representative of exocytic events occurring
at the apical membrane of polarized cholangiocytes, the rate of apical
exocytosis was measured in polarized NRC monolayers, which showed an
average rate of increase of 1.4% apical PM/min (Fig.
3; 142 ± 3% of initial intensity
after 30 min; n = 4). Known to inhibit vesicular
trafficking, increases in FM 1-43 fluorescence were significantly
blunted (106 ± 3% of initial levels after 30 min;
n = 4) in monolayers incubated with cold FM 1-43
medium (31, 33). Together, these studies demonstrate a
robust basal rate of vesicular trafficking to the apical membrane of
cholangiocytes.
|
Endocytic membranes are partially recycled to plasma membrane.
Measurements of Cm were used to evaluate whether
basal exocytosis was paralleled by an increase in plasma membrane
surface area. The initial Cm was 36.5 ± 3.6 pF (n = 13), corresponding to a calculated membrane
surface area of 3,650 µm2 (14). In contrast
to the basal rate of exocytosis, the total plasma membrane surface area
(0.20 + 0.12%PM/min; n = 4) remained relatively
constant (Fig. 2C; Table 1).
This conclusion is supported by comparison of the residual fluorescence
that remains with the cells after FM 1-43 is washed from the
plasma membrane. In all cases, residual fluorescence (i.e., endocytosed
FM 1-43) paralleled the level of increase in fluorescence
intensity after FM 1-43 equilibration with the plasma membrane
(i.e., exocytosis). Thus cholangiocytes maintain a relatively constant
membrane surface area by matching basal rates of exocytosis with
equivalent rates of plasma membrane endocytosis.
|
cAMP-dependent modulation of exocytosis.
cAMP regulates exocytosis in a number of cell types including
cholangiocytes (16). To evaluate the response of Mz-ChA-1 cells to cAMP, cells were exposed to maneuvers designed to increase or
decrease cAMP/PKA activity. Addition of CPT-cAMP (500 µM) increased the FM 1-43 fluorescence intensity in cells over time (i.e., rate of exocytosis) by 71 ± 31% (n = 5) over paired
control cells (Fig. 4A). The
delay in the CPT-cAMP effect varied marginally between preparations and
is likely a result of multiple factors including chamber equilibration,
cell permeation, and intracellular biochemical pathway interactions of
CPT-cAMP.
|
Evidence for subapical membrane vesicles in cholangiocytes.
These high rates of membrane turnover imply the presence of an abundant
vesicle population. Standard histological analysis in whole mount
preparations of the apical membrane/subapical domain from NRC cells
(Fig. 1) were osmicated to highlight filamentous and proteinaceous
elements in the apical patches (Fig. 5,
A and B). The osmicated patches varied in
electron density, reflecting differences in the depth of cellular
material that remained physically associated with the apical membrane.
In areas of lower electron density, a broad filamentous network was
observed (Fig. 5A). The presence of plasma
membrane-associated clathrin triskelion structures (Fig. 5B)
indicates that these areas are at the subapical plasma membrane
surface. To highlight the presence of membranous structures by negative
contrast, UA (Fig. 5, C and D) and PTA (Fig. 5,
E and F) were applied to apical membrane
preparations. Both staining methods revealed a dense population of
~140-nm oval-shaped bodies (Fig. 5, B and C).
Consistent with membrane bilayers, higher magnification of the
UA-stained patches (Fig. 5D) showed 5.2-nm excluded spaces encircling the denser 140-nm bodies. These observations provide direct
evidence for an abundant population of membrane vesicles in the
subapical domain of NRC cells.
|
Subapical vesicles are derived, in part, from plasma membrane
endocytosis.
To determine whether the subapical vesicle population is in
communication with the plasma membrane, the apical membrane was incubated with FM 1-43 for 30 min to permit endocytosis and the subsequent presence of FM 1-43 in subapical vesicles was examined under fluorescence microscopy. Thicker patches showed pleiomorphic vesicles of varying fluorescence densities (data not shown). Less dense
patches with low background levels revealed a broadly distributed population of FM 1-43-stained vesicles (Fig.
6). The resolution power for green light
with a 1.4 NA lens is ~200 nm. Although light microscopy is unable to
resolve 140-nm vesicles, the endocytosed vesicles (Fig. 6B)
were of an appropriate size compared with intact NRC cells (Fig.
6A), were similar in size to 200-nm fluorescent beads (Fig.
6C), and were relatively homogeneous in fluorescence density.
|
F-actin distributes with subpopulation of endocytic vesicles.
The actin cytoskeleton modulates vesicular trafficking in a number of
cell types. With Texas red-phalloidin staining of NRC apical membrane
patches, F-actin was clearly localized to a population of subapical
vesicles. To evaluate the relationship between the F-actin-labeled
vesicle population and endocytic vesicles, the subapical endocytic
vesicle population in NRC cells was initially labeled with FM 1-43
and apical membrane patches were prepared and stained for F-actin (Fig.
7). Interestingly, after 30 min of FM
1-43 loading, essentially all (~98%) of the F-actin labeled vesicles colocalized with FM 1-43-labeled endocytic vesicles. However, only ~13% of the FM 1-43-labeled vesicles costained
for F-actin.
|
Disruption of F-actin cytoskeleton blunts membrane exocytosis.
To determine whether F-actin contributes to membrane exocytosis, the
comparative rates of exocytosis were quantified with FM 1-43 after
CD or Jas pretreatment. Interestingly, both manipulations resulted in
decreased rates of exocytosis (Fig. 8,
A and B). Quantitative analysis showed that the
rates of exocytosis in CD- and Jas-treated cells were only 51 ± 13% (n = 9) and 58 ± 9% (n = 6)
of control rates, respectively (Fig. 8C).
Cm measurements in Mz-ChA-1 cells treated with
CD or Jas showed that the total membrane surface area remained largely
unchanged despite the decreased rates of exocytosis [Table 1; CD:
0.8 + 1.2%PM/min (n = 6); JAS:
0.7 + 0.7%PM/min (n = 4)]. Thus, despite the substantial
decrease in the rate of exocytosis after perturbation of the actin
cytoskeleton, the cells had diminished rates of endocytosis that again
paralleled the decrease in exocytosis. The observation that either
actin filament stabilization or depolymerization results in diminished rates of membrane exocytosis suggests that the actin cytoskeleton may
contribute at multiple points along the vesicle trafficking pathway.
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The present study of cholangiocytes used complementary biochemical, histological, and electrophysiological techniques to quantitatively assess the trafficking of membrane vesicles at the plasma membrane and to reveal a dense subapical vesicle population capable of supporting the measured rates of the trafficking events.
Cholangiocytes have substantial rate of apical membrane turnover. Rates of exocytosis under basal conditions are cell type specific. A seminal work with fluid phase markers in isolated macrophages and a fibroblast cell line documented basal rates of endocytosis equal to 3.1% and 0.8%PM/min, respectively (32). The rates of exocytosis were inferred to be equivalent to these values. Recently, HTC cells, a hepatocyte cell line, were shown to have a constitutive rate of membrane turnover equal to 2.0%PM/min (17). In contrast, spontaneous exocytosis at neuronal synapses is nominal (3). In earlier cholangiocyte studies, no release of acridine orange from acidic vesicles was observed under basal conditions (16). Using FM 1-43 fluorescence to directly quantify membrane insertion into the plasma membrane of cholangiocytes, the present studies demonstrated a significant rate (1.33 ± 0.16%PM/min) of exocytosis under basal conditions. The apparent disparity in these two cholangiocyte studies likely reflects differences in the assays used and the specific vesicle populations being measured. Specifically, acridine orange fluorescence targets only acidic vesicles or vacuoles whereas FM 1-43 fluorescence measures all exocytosed membrane. Thus the acidic vesicle population may traffic to the plasma membrane only in response to specific stimuli.
Interestingly, both methods detected increases in cholangiocyte exocytosis within minutes of increasing intracellular cAMP (cAMPi) and persisting over a 10-min period (Fig. 4; Ref. 16). This contrasts with insulin-induced exocytic response (17). In HTC cells, the addition of insulin results in an abrupt, steplike increase in FM 1-43 fluorescence over 1-2 min followed by a return of exocytosis to the basal rate. Although insulin addition to hepatocytes likely induces the rapid insertion of a distinct population of insulin-responsive vesicles, increased cAMPi in cholangiocytes may induce either an increased rate of delivery of a more general vesicle population or the comparatively slow mobilization of a distinct population of cAMP-responsive vesicles. Nevertheless, the differences in response patterns suggest the presence of multiple mechanisms of regulated vesicle trafficking.Comparative quantitation of vesicle density, exocytosis, and
endocytosis.
Reminiscent of the dense apical endosome population observed in LR
Gold-embedded thin sections of rat ileum (41), the present studies detected a dense population of vesicles in the subapical domain
of NRC cells (Figs. 5, 6, and 7). In NRC cells, the apical membrane
comprises 57% of the total plasma membrane (8). By using
the capacitance measurements of plasma membrane surface area in
Mz-ChA-1 cells (3,650 µm2) to approximate the plasma
membrane surface area of an NRC cell, the apical membrane surface is
estimated to be ~2,100 µm2 [(3,650
µm2) × (0.57) 2,100 µm2].
Given the observed rate of apical exocytosis in NRC cells of ~1.4%
of the apical membrane surface area per minute (Fig. 2C), this equates to ~30 µm2 of vesicular membrane being
added to the apical membrane per minute [(2,100
µm2) × (0.014)
30 µm2]. The
membrane surface area of an 140-nm vesicle is ~0.06 µm2
[4
r2 = (4) × (3.14) × (0.07 µm)2
0.06 µm2,
where r is vesicle radius]. If the 140-nm vesicles
observed in the subapical domain are solely responsible for the
exocytosed membrane, ~500 vesicle fusion events would be required per
minute per cell [(0.06 µm2) × (500) = 30 µm2]. Furthermore, with
Cm remaining essentially unchanged under basal
conditions (Table 1), the exocytic events must be paralleled by
endocytic events and the presence of an endocytic vesicle population with equivalent surface area. This predicted rate of endocytic vesicle
formation is similar in magnitude to the 125 pinocytotic vesicles per
minute that form in resting macrophages (32). Despite these projections, conventional histological analysis of cholangiocytes has not demonstrated a vesicle population capable of accounting for
this robust rate of vesicular trafficking. However, the whole mount,
negative-contrast apical subdomain preparations (Fig. 5) revealed a
dense population of vesicles that are demonstrably in communication
with the apical membrane (Figs. 6 and 7) and present in sufficient
numbers to account for these significant rates of membrane trafficking.
Actin cytoskeleton modulates vesicular trafficking at multiple
points.
In neurons, filamentous actin is concentrated near presynaptic
membranes (9, 12) and can moderate multiple steps in the spatial and temporal organization of reserve and readily releasable pools of synaptic vesicles as well as their migration to the
presynaptic membrane (7, 19, 23, 40). Studies in
nonneuronal cells suggest that the actin cytoskeleton may form a
cortical barrier to vesicle trafficking to the plasma membrane
(5, 24, 38) and moderate endocytosis (10,
28). In an opossum kidney proximal tubule cell line, latrunculin
B, an actin filament-disrupting agent, blocked the endothelin-induced
exocytic insertion of Na+/H+ exchanger 3 into
the apical membrane and its subsequent increase in activity
(27). Interestingly, disruption of the actin cytoskeleton with cytochalasin D, the actin filament-disrupting agent used in the
present study, failed to have an similar effect. In gastric parietal
cells, specific actin cytoskeletal interactions contribute to the
H+-K+-ATPase insertion and profound
reorganization of the apical membrane. The -actin isoform is
specifically concentrated at the canalicular region (42),
and there is a phosphorylation-dependent association of ezrin, an
actin-membrane linking protein, with the apical membrane of stimulated
parietal cells (34, 35). In pancreatic acinar cells,
perturbation of the actin cytoskeleton impacts both zymogen granule
secretion and subsequent membrane retrieval (1, 36, 37).
Furthermore, filamentous actin associates transiently with zymogen
granules during granule translocation to the apical membrane (37).
![]() |
ACKNOWLEDGEMENTS |
---|
The authors thank Dr. Bill Betz and Steve Fadul for technical and theoretical discussions regarding FM dyes and Dr. Kathryn Howell for discussions on membrane vesicles and trafficking.
![]() |
FOOTNOTES |
---|
This work was supported by American Liver Foundation Grant ALF PN 9801-014 and Cystic Fibrosis Foundation Grant DOCTOR01GO (to R. B. Doctor), an American Liver Scholar Award (to G. Kilic), and National Institute of Diabetes and Digestive and Kidney Diseases Grants R01-DK-46082 and DK-43278 (to J. G. Fitz).
Address for reprint requests and other correspondence: R. B. Doctor, Box B158, 4200 E. 9th Ave., Denver, CO 80262 (E-mail: brian.doctor{at}uchsc.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.
First published December 5, 2001;10.1152/ajpcell.00367.2001
Received 21 March 2001; accepted in final form 28 November 2001.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1.
Baudin, H,
Stock C,
Vincent D,
and
Grenier J.
Microfilamentous system and secretion of enzyme in the exocrine pancreas.
J Cell Biol
66:
165-181,
1975[Abstract].
2.
Betz, W,
and
Bewick G.
Optical analysis of synaptic vesicle recycling at the frog neuromuscular junction.
Science
255:
200-203,
1992[ISI][Medline].
3.
Betz, W,
Mao F,
and
Bewick G.
Activity-dependent fluorescent staining and destaining of living vertebrate motor nerve terminals.
J Neurosci
12:
363-375,
1992[Abstract].
4.
Buanes, T,
Grotmol T,
Landsverk T,
and
Raeder MG.
Secretin empties bile duct cell cytoplasm of vesicles when it initiates ductular HCO
5.
Cheek, T,
and
Burgoyne R.
cAMP inhibits both nicotine-induced actin disassembly and catecholamine secretion from bovine adrenal chromaffin cells.
J Biol Chem
262:
11663-11666,
1987
6.
Cochilla A, Angleson J, and Betz W. Monitoring secretory membrane
with FM1-43 fluorescence. Ann Rev Neuroscience:
1-10, 1999.
7.
Cole, J,
Villa B,
and
Wilkinson R.
Disruption of actin impedes transmitter release in snake motor terminals.
J Physiol (Lond)
525:
579-586,
2000
8.
Doctor, R,
Dahl R,
Salter K,
and
Fitz J.
Reorganization of cholangiocyte membrane domains represents an early event in rat liver ischemia.
Hepatology
29:
1364-1374,
1999[ISI][Medline].
9.
Drenckhahn, D,
and
Kaiser H.
Evidence for the concentration of F-actin and myosin in synapses and in the plasmalemmal zone of axons.
Eur J Cell Biol
31:
235-240,
1983[ISI][Medline].
10.
Durrbach, A,
Louvard D,
and
Coudrier E.
Actin filaments facilitate two steps of endocytosis.
J Cell Sci
109:
457-465,
1996
11.
Feranchak, AP,
Roman RM,
Doctor RB,
Salter KD,
Toker A,
and
Fitz JG.
The lipid products of PI3 kinase contribute to regulation of cholangiocyte ATP and chloride transport.
J Biol Chem
274:
30979-30986,
1999
12.
Fifkova, E,
and
Delay R.
Cytoplasmic actin in neuronal processes as a possible mediator of synaptic plasticity.
J Cell Biol
95:
345-350,
1982[Abstract].
13.
Goldenring, J.
Pools of actin in polarized cells: some filaments are more stable than others.
Am J Physiol Cell Physiol
281:
C386-C387,
2001
14.
Hille, B.
Ionic Channels of Excitable Membranes. Sunderland, MA: Sinauer, 1992.
15.
Horrigan, F,
and
Bookman R.
Releasable pools and the kinetics of exocytosis in adrenal chromaffin cells.
Neuron
13:
1119-1129,
1994[ISI][Medline].
16.
Kato, A,
Gores G,
and
LaRusso N.
Secretin stimulates exocytosis in isolated bile duct epithelial cells by a cyclic AMP-mediated mechanism.
J Biol Chem
267:
15523-15529,
1992
17.
Kilic, G,
Doctor R,
and
Fitz J.
Insulin stimulates membrane conductance in a liver cell line: evidence for insertion of ion channels through a PI3 kinase-dependent mechanism.
J Biol Chem
276:
26762-26768,
2001
18.
Knuth, A,
Gabbert H,
Dippold W,
Klein O,
Sachsse W,
Bitter-Suermann D,
Prellwitz M,
and
Meyer zum Buschenfelde KH.
Biliary adenocarcinoma. Characterisation of three new human tumor cell lines.
J Hepatol
1:
579-596,
1985[ISI][Medline].
19.
Landis, D,
Hall A,
Weinstein L,
and
Reese T.
The organization of cytoplasm at the presynaptic active zone of a central nervous system synapse.
Neuron
1:
201-209,
1988[ISI][Medline].
20.
Lindau, M,
and
Neher E.
Patch-clamp techniques for time-resolved capacitance measurements in single cells.
Pflügers Arch
411:
137-146,
1988[ISI][Medline].
21.
Marinelli, RA,
Pham L,
Agre P,
and
LaRusso NF.
Secretin promotes osmotic water transport in rat cholangiocytes by increasing AQP-1 water channels in plasma membrane.
J Biol Chem
272:
12984-12988,
1997
22.
Martin, S,
Tellam J,
Livingstone C,
Slot J,
Gould G,
and
James D.
The glucose transporter (GLUT-4) and vesicle-associated membrane protein-2 (VAMP-2) are segregated from recycling endosomes in insulin-sensitive cells.
J Cell Biol
134:
625-635,
1996[Abstract].
23.
Morales, M,
Colicos M,
and
Goda Y.
Actin-dependent regulation of neurotransmitter release at central synapses.
Neuron
27:
539-550,
2000[ISI][Medline].
24.
Muallem, S,
Kwiatkowska K,
Xu X,
and
Yin H.
Actin filament disassembly is a sufficient trigger for exocytosis in non-excitable cells.
J Cell Biol
128:
589-598,
1995[Abstract].
25.
Okamoto, C,
and
Forte J.
Vesicular trafficking machinery, the actin cytoskeleton and H+-K+-ATPase recycling in the gastric parietal cell.
J Physiol (Lond)
532:
287-296,
2001
26.
Park, C,
Leem C,
Jang Y,
and
Shim Y.
Vesicular transport as a new paradigm in short-term regulation of transepithelial transport.
J Korean Med Sci
15:
123-132,
2000[ISI][Medline].
27.
Peng, Y,
Amemiya M,
Yang X,
Fan L,
Moe O,
Yin H,
Preisig P,
Yanagisawa M,
and
Alpern R.
ETB receptor activation causes exocytic insertion of NHE3 in OKP cells.
Am J Physiol Renal Physiol
280:
F34-F42,
2001
28.
Qualmann, B,
Kessels M,
and
Kelly R.
Molecular links between endocytosis and the actin cytoskeleton.
J Cell Biol
150:
F111-F116,
2000
29.
Salter, K,
Roman R,
LaRusso N,
Fitz J,
and
Doctor R.
Modified culture conditions of normal rat cholangiocytes induces the expression of bile duct epithelial properties.
Lab Invest
80:
1775-1778,
2000[ISI][Medline].
30.
Sanan, D,
and
Anderson R.
Simultaneous visualization of LDL receptor distribution on clathrin lattices on membranes torn from the upper surface of cultured cells.
J Histochem Cytochem
39:
1017-1024,
1991[Abstract].
31.
Silverstein, S,
Steinman R,
and
Cohn Z.
Endocytosis.
Annu Rev Biochem
46:
669-722,
1977[ISI][Medline].
32.
Steinman, R,
Brodie S,
and
Cohn Z.
Membrane flow during pinocytosis.
J Cell Biol
68:
665-687,
1976[Abstract].
33.
Tomada, H,
Kishimoto Y,
and
Lee Y.
Temperature effect on endocytosis and exocytosis by rabbit alveolar macrophages.
J Biol Chem
264:
15445-15450,
1989
34.
Urshidani, T,
Hanzel D,
and
Forte J.
Protein phosphorylation associated with stimulation of rabbit gastric glands.
Biochim Biophys Acta
930:
209-219,
1987[ISI][Medline].
35.
Urshidani, T,
Hanzel D,
and
Forte J.
Characterization of an 80-kDa phosphoprotein involved in parietal cell stimulation.
Am J Physiol Gastrointest Liver Physiol
256:
G1070-G1081,
1989
36.
Valentijn, K,
Gumkowski F,
and
Jamieson J.
The subapical cytoskeleton regulates secretion and membrane retrieval in pancreatic acinar cells.
J Cell Sci
112:
81-96,
1999
37.
Valentijn, J,
Valentijn K,
Pastore L,
and
Jamieson J.
Actin coating of secretory granules during regulated exocytosis correlates with the release of rab3D.
Proc Natl Acad Sci USA
97:
1091-1095,
2000
38.
Vitale, M,
Seward E,
and
Trifaro J.
Chromaffin cell cortical actin network dynamics control the size of the release-ready vesicle pool and the initial rate of exocytosis.
Neuron
14:
353-363,
1995[ISI][Medline].
39.
Vroman, B,
and
LaRusso NF.
Development and characterization of polarized primary cultures of rat intrahepatic bile duct epithelial cells.
Lab Invest
74:
303-313,
1996[ISI][Medline].
40.
Wang, X,
Zheng J,
and
Poo M.
Effects of cytochalasin treatment on short-term synaptic plasticity at developing neuromuscular junctions in frogs.
J Physiol (Lond)
491:
187-195,
1996[Abstract].
41.
Wilson, J,
Whitney J,
and
Neutra M.
Identification of an endosomal antigen specific to absorptive cells of suckling rat ileum.
J Cell Biol
105:
691-703,
1987[Abstract].
42.
Yao, X,
Chaponnier C,
Gabbiani G,
and
Forte J.
Polarized distribution of actin isoforms in gastric parietal cells.
Mol Biol Cell
6:
541-557,
1995[Abstract].