Cell Physiology

Cholangiocytes exhibit dynamic, actin-dependent apical membrane turnover

R. Brian Doctor, Rolf Dahl, Laura Fouassier, Gordan Kilic, J. Gregory Fitz


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

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.


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 (R t >1,000 Ω · cm2), net apical HCO 3 secretion, and a transepithelial H+ gradient.

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%).

Four compounds were evaluated for their effects on the rate of exocytosis. CPT-cAMP (500 μM in E-buffer), a cell-permeant cAMP analog, was added after an initial rate of exocytosis was obtained. The Rp diastereomer of adenosine 3′,5′-cyclic monophosphothioate (Rp-cAMPS; 5 μM in E- buffer), a potent protein kinase A (PKA) inhibitor, was preincubated for 60–90 min before study. Rp-cAMPS-treated cells showed no overt histological effects. Cytochalasin D (CD; 10 μM in EtOH), an actin filament-depolymerizing agent, and jasplakinolide (Jas; 50 nM in EtOH), an actin filament-stabilizing agent, required 15 min of preincubation to show measurable effects. These concentrations resulted in ∼50% of the cells showing some degree of membrane blebbing. Higher concentrations resulted in >90% of cells with membrane blebs. Cells selected for observation had no observable membrane blebs. Vehicle-treated cells (0.1% EtOH) showed no signs of membrane blebbing, and FM 1–43 exocytosis rates were not different from those of untreated controls. To minimize the potential impact from variations in the basal rates of exocytosis, experimental studies were performed with paired controls.

To measure the rate of apical exocytosis in a polarized cholangiocyte epithelium, FM 1–43 was added to the apical chamber of NRC monolayers (R t > 1,000 Ω · cm2), monolayers were transferred to the microscope stage, and fluorescence intensity was measured at room temperature at 0, 15, and 30 min. The studies used an upright Nikon microscope with a ×10 long working distance objective on a BRC-600 confocal system. FM 1–43 fluorescence was excited with the 488-nm line from a krypton-argon laser. Slit widths were set at the maximal setting. To inhibit the rate of exocytosis, monolayers were incubated and the studies initiated in ice-cold media (33). Parenthetically, the initial fluorescence intensity measurement from NRC cells was consistently higher at 4°C vs. room temperature, suggesting that there may be a temperature dependence of FM 1–43 fluorescence efficiency. All measurements within an experiment were taken at identical settings. The fluorescence intensities were quantified with NIH Image 6.0.

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) =I ss + (I 0I ss) exp(−t/τ), whereI(t) is current response,I ss is steady-state current,I 0 is peak current, t is time, and τ is time constant. From the fitted parameters (I 0, I ss, and τ), the membrane capacitance (C m), access resistance (R a), and membrane conductance (G m) were calculated as previously described (20). This procedure was repeated every 3 s. The recordings were stopped if G m 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.

Fig. 1.

Low-magnification view of osmicated apical membrane patches. To identify membrane vesicles in the subapical domain, the apical membrane of normal rat cholangiocyte (NRC) cells was allowed to adhere to coated electron microscopy (EM) grids and the membrane and physically associated components in the subapical domain were subsequently pulled free from the rest of the cell. Shown are an electron micrograph of NRC cells in the z-axis (A), a cartoon depicting the grid adhesion and membrane isolation process (B), and a low-magnification en face view of osmicated apical membrane patches (C). Islands of material are the adhered apical membrane and intracellular material that remained associated with the membrane.

For fluorescence studies, NRC monolayers were washed at 4°C (1× with PBS, 2× with IC buffer); coated coverslips (10 mg/ml poly-l-lysine) were laid atop the monolayer, inverted, and placed on an ice-chilled pedestal for 15 min; and the coverslips were pulled from the monolayer, washed (3× with IC buffer), and mounted for observation. In studies involving FM 1–43 loading of intracelluar vesicles, the apical chamber of NRC monolayers first received 4.7 μM FM 1–43 for 30 min and the FM 1–43 was removed from the plasma membrane by 5-min exposure to buffer without FM 1–43. In studies involving F-actin staining of vesicles, apical membranes were prepared as above, incubated with Texas red-phalloidin (100 μg/ml in IC buffer; 15 min), and washed (3× 1 min; IC buffer) before mounting.

Spectral analysis showed a fairly broad range for FM 1–43 fluorescence, with 22% of its fluorescence signal in the Texas red window. Conversely, <5% of the Texas red-phalloidin fluorescence was observed in the FITC window. In studies with both FM 1–43 and Texas red-phalloidin, FM 1–43 and Texas red-phalloidin concentrations were optimized (see above) to minimize the FM 1–43 signal in the Texas red window. There remained a weak but detectable fluorescence crossover from both fluors. Reported results with FM 1–43 and Texas red-phalloidin were similar in parallel studies comparing 1) FM 1–43 and Texas red-phalloidin in separate preparations or 2) FM 4–64 (Molecular Probes) and FITC-phalloidin fluors. FM 4–64 has functional characteristics similar to those of FM 1–43 but a fluorescence spectrum (absorption/emission = 558 nm/734 nm) more specific to the Texas red window.

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 either1) in 2% methylcellulose containing 0.3% UA or2) 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.


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.

Fig. 2.

Cholangiocytes display basal exocytosis. FM 1–43 was used to evaluate the rates of exocytosis and membrane recycling in Mz-ChA-1 cells. A: FM 1–43 fluorescence images of Mz-ChA-1 cells are shown before FM 1–43 addition (0 min, top; light image), after initial equilibration with the plasma membrane (3 min), after longer incubation in FM 1–43 (15 min), and after 18 min of FM 1–43 incubation followed by 12 min of perfusion in the absence of FM 1–43 to remove FM 1–43 from the plasma membrane (18/12 min, top; increased sensitivity to demonstrate internalized fluorescent membranes). To permit direct comparison, all bottom panels were observed at identical exposure times and settings.B: under basal conditions, the constitutive rate of exocytosis in Mz-ChA-1 cells (slope of the points in region 1) was 1.33 ± 0.16% plasma membrane surface area (PM)/min. After washing of FM 1–43 from the plasma membrane, the rate of membrane recycling of endocytosed FM 1–43 back to the plasma membrane (slope of the points in region 2) was 0.48 ± 0.06%PM/min. C: membrane capacitance measurements were normalized to initial values and were relatively stable over the time course of these studies.

Figure 2, A and B, illustrates the qualitative and quantitative fluorescence response of Mz-ChA-1 cells under control conditions after the addition and removal of FM 1–43. Qualitatively, before the addition of FM 1–43, cells have a relatively low background fluorescence (Fig. 2 A). All images in Fig. 2 A, bottom, were taken with identical exposures, settings, and processing steps. After the addition of FM 1–43 to the extracellular buffer there was a rapid increase in FM 1–43 fluorescence (Fig. 2 A) from the equilibration of FM 1–43 with the plasma membrane. The delay in fluorescence is due primarily to the transit time of the dye through the perfusion apparatus. This initial peak fluorescence value represents the plasma membrane surface area (PM). Over time (Fig.2 A), fluorescence intensity increased because of exocytic insertion and staining of new membrane (6). Endocytic retrieval of FM 1–43-labeled membrane occurred concurrently during this period. After the removal of FM 1–43 from the medium, there was a rapid decrease in fluorescence representing the efflux of FM 1–43 from the plasma membrane. After washing, only FM 1–43 that was endocytosed remained (Fig. 2 A; 18-min FM exposure/12-min washing; top 18/12 min panel represents same cell with increased sensitivity to highlight cytoplasmic/perinuclear distribution of FM 1–43).

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. 2 B,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.

Fig. 3.

Apical exocytosis in NRC monolayers. FM 1–43 fluorescence intensity was measured from the apical membrane of NRC monolayers. Interestingly, they showed a linear increase in fluorescence to 142 ± 3% of the initial fluorescence intensity measurement after 30 min. In contrast, paired incubation of NRC cells at 4°C, which inhibits membrane trafficking, blunted the fluorescence intensity increase (106 ± 3% of the initial fluorescence intensity measurement after 30 min at 4°C).

Endocytic membranes are partially recycled to plasma membrane.

Measurements of C m were used to evaluate whether basal exocytosis was paralleled by an increase in plasma membrane surface area. The initial C m 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. 2 C; 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.

View this table:
Table 1.

Alterations in cholangiocyte membrane capacitance

These findings indicate parallel rates of exocytosis and endocytosis of ∼1.33%PM/min. After FM 1–43 is removed from the extracellular buffer and washed from the plasma membrane, the rate at which membrane vesicles recycle back to the plasma membrane can be quantified by measuring the rate at which FM 1–43 fluorescence intensity decreases because of FM 1–43-laden endocytic membranes returning to the plasma membrane and unloading FM 1–43 into the extracellular buffer (Fig. 2 B, region 2). This rate of FM 1–43 fluorescence decrease was 0.48 ± 0.06%PM/min (n = 8). This observation indicates that ∼36% (0.48%PM/min recycling ÷ 1.33%PM/min endocytosis = 0.36) of the endocytosed membrane recycled to the plasma membrane under basal conditions. Conversely, 64% of endocytosed FM 1–43 remained within the cell. Fluorescence localization of the FM 1–43 that remained in the cells showed that these endocytosed membranes migrated into a cytoplasmic/perinuclear locale (Fig.2 A, 18/12 min).

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. 4 A). 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.

Fig. 4.

cAMP modulates the rate of exocytosis in cholangiocytes. As measured by the comparative increase in FM 1–43 fluorescence intensity over time, the addition of 8-(4-chlorophenylthio)adenosine 3′,5′-cyclic monophosphothioate (CPT-cAMP; arrow), a cell-permeant cAMP analog, resulted in a significant 71 ± 31% increase in the rate of exocytosis. In paired experiments, preincubation of Mz-ChA-1 cells with the Rp diastereomer of adenosine 3′,5′-cyclic monophosphothioate (Rp-cAMPS), a potent protein kinase A (PKA) inhibitor, resulted in a decrease in the basal rate of exocytosis (47 ± 11% of control). These findings indicate the level of cAMP-PKA activity contributes to the regulation of exocytosis.

To determine whether background levels of cAMP and PKA activity contribute to the basal rate of exocytosis, FM 1–43 fluorescence intensity values were measured in control cells vs. paired cells that were preincubated with Rp-cAMPS (5 μM), a potent inhibitor of PKA. Cells treated with Rp-cAMPS had a significant decrease (47 ± 11% of controls; n = 6) in their basal rates of exocytosis (Fig. 4 B). Thus increases or decreases in cAMP/PKA activity are paralleled by changes in the rates of exocytosis.

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. 5 A). The presence of plasma membrane-associated clathrin triskelion structures (Fig. 5 B) 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. 5 D) 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.

Fig. 5.

Alternate EM processing reveals an abundant vesicle population in the subapical domain of NRC cells. Osmicated apical membrane patches revealed structural elements associated with the apical domain (A), and, at higher magnification, clathrin lattices (arrow) associated with the cytoplasmic face of the apical membrane were observed (B). Negative staining with uranyl acetate in methylcellulose (C, D) or phosphotungstic acid (E, F) revealed a dense population of 140-nm bodies in the subapical domain. D: at higher magnification, a distinctive, 5.2-nm membrane track is observed encircling the vesicles (arrowheads). Bar, 500 (A, C,E) or 100 (B, D, F) nm.

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. 6 B) were of an appropriate size compared with intact NRC cells (Fig.6 A), were similar in size to 200-nm fluorescent beads (Fig.6 C), and were relatively homogeneous in fluorescence density.

Fig. 6.

Subapical vesicle population arises, in part, from apical membrane endocytosis. All images shown were taken at equivalent magnifications. A: MeOH-fixed NRC cells are shown as a reference of cell size (bar = 20 μm). B: apical incubation of NRC cells with FM 1–43 results in fluorescence labeling of subapical vesicles. The vesicle density was less than observed by negative-contrast electron microscopy. C: fluorescence imaging of 200-nm fluorescent beads (Molecular Probes) allows for comparative sizing of particles and suggests that the FM 1–43-laden vesicles are similar in size to the 140-nm vesicles observed by negative-contrast electron microscopy.

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.

Fig. 7.

F-actin codistributes with a subpopulation of endocytosed subapical vesicles. FM 1–43 staining of endocytic vesicles (left) reveals a broad endocytic vesicle population across the subapical domain. Texas red-phalloidin staining (right) shows specific interaction with a smaller population of vesicles from the same field. Arrowheads denote FM 1–43-loaded vesicles that colabeled for F-actin (13% of the FM 1–43-positive endocytotic vesicle population). After 30 min of FM 1–43 incubation, only ∼1% of the Texas red-phalloidin-positive vesicles were not coincidentally positive for FM 1–43 (asterisks).

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. 8 C).C m 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.

Fig. 8.

Alterations in the actin cytoskeleton modify membrane exocytosis. To assess the involvement of the actin cytoskeleton in membrane exocytosis, the actin cytoskeleton was either disrupted with cytochalasin D (A) or stabilized with jasplakinolide (B). Interestingly, both agents resulted in significantly diminished rates of exocytosis (C; cytochalasin D: 51 ± 13% of control; jasplakinolide: 58 ± 13% of control).


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. 2 C), 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[4πr 2 = (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, withC m 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 β-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).

Using cytochalasin D and jasplakinolide, two agents with broad effects on the actin cytoskeleton, the present studies demonstrated that the actin cytoskeleton plays a significant role in membrane trafficking in cholangiocytes (Fig. 8). If the actin cytoskeleton acted at a single step in membrane trafficking, actin filament depolymerization (i.e., cytochalasin D) and actin stabilization (i.e., jasplakinolide) would be predicted to have opposing effects. The similarity in their responses (Fig. 8) suggests that the actin cytoskeleton is likely involved in multiple steps in the membrane trafficking pathway. Identification of discrete actin-associated proteins and individual steps in the membrane trafficking pathways will be required to dissect the specific roles of the actin cytoskeleton in this process. As an initial step, filamentous actin was observed to associate with a subpopulation of endocytic vesicles (Fig. 7). This interaction could result from F-actin interacting transiently with all endocytosed membrane vesicles or targeting a discrete subset of endocytic vesicles. For example, the actin cytoskeleton was previously postulated to regulate the recycling of endocytosed cargoes back to the apical membrane (13). Future studies will focus on identifying specific actin-associated proteins on this vesicle population and evaluating their functional significance in the formation and trafficking of this pool of endocytic vesicles.

In summary, the principal findings from these cholangiocyte models are1) there are robust rates of basal exocytosis and endocytosis corresponding to 1.3%PM/min; 2) approximately one-third of the endocytosed membrane recycles back to the plasma membrane; 3) the rate of exocytosis is modulated by cAMP;4) the actin cytoskeleton contributes to these trafficking events; 5) there is a dense submembranous vesicle population that can account quantitatively for this level of vesicle trafficking; and 6) filamentous actin encapsulates a subpopulation of endocytic vesicles. Together, these findings support an important potential role for vesicle trafficking in modifying the composition of the apical plasma membrane in response to changing physiological demands.


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.


  • 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


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