Exocytosis has been proposed to participate in the formation of pseudopods. Using video-enhanced microscopy, we directly visualized exocytosis of single vesicles in living Physarum plasmodia migrating on a substrate. Vesicles containing slime, the plasmodial extracellular matrix, of ∼3.5 μm in diameter, shrank at the cell periphery at the average rate of ∼1 μm/second, and became invisible. Immediately after exocytotic events, the neighboring cell surface extended to form a protrusion. The rate of extension was ∼1 μm/second. The protrusion showed lamella-like morphology, and contained actin microfilaments. Electron microscopy suggested that the organization of microfilaments in such protrusions may be a random meshwork rather than straight bundles. These morphologies suggest that protruded regions are pseudopods. Importantly, only the slime-containing vesicle preferentially invaded the hyaline layer that consists of dense actin microfilaments while the other vesicular organelles remained in the granuloplasm. Quantitative analysis demonstrated a linear relationship in terms of their surface area, between individual protrusions and single slime-containing vesicles. It is, therefore, likely that most of the plasma membrane of the protrusion was supplied by fusion of the slime-containing vesicle during exocytosis.

Exocytosis is a fundamental process that is seen in all eukaryotic cells, and plays important roles in cell physiology. It is the mechanism underlying the following two functions of the cell. The first is secretion whereby contents of intracellular membrane vesicles are released to the extracellular space. The other is supply of new membrane materials to the plasma membrane including proteins and lipids. Macromolecules supplied in this way increase the surface area of the plasma membrane. These two functions of exocytosis, i.e. vesicle contents delivery and membrane material addition, give rise to cell shape changes in a cooperative manner in various types of cells. Examples include pseudopod extension (Singer and Kupfer, 1986; Bretscher, 1984), neurite elongation (Bray, 1970; Popov et al., 1993), cytokinesis (Bluemink and Delaat, 1977), and yeast budding (Drubin, 1991). The site of exocytosis has been suggested to be located at the growth cone of growing neurons (Craig et al., 1995; Dai and Sheetz, 1995), tips of extending buds of yeast (Field and Schekman, 1980), and cleavage furrows in dividing cells (Bluemink and Delaat, 1973; Byers and Armstrong, 1986). These localized exocytotic events imply a role for exocytotic vesicles in cell shape changes as a supplier of new membrane materials and the extracellular components to specific regions in cells, where dramatic expansion of the plasma membrane and interactions with the freshly secreted extracellular components are taking place.

In a variety of cell types that locomote on substrate, the leading edge has been suggested to be a site for membrane insertion. Bretscher and co-workers (Bretscher, 1983; Bretscher and Thomson, 1983) showed a graded distribution of surface receptors with remarkable enrichment in the cell periphery of HeLa cells. Hopkins (1985) showed that the majority of the newly appearing transferrin receptors on the cell surface arise near the cell margin of A431 cells using immuno-gold labeling. Since these receptors are transported to the cell surface by exocytotic mechanisms (Goldstein et al., 1985), these results suggest localized exocytosis occurring at the cell periphery. When the surface distribution of the newly synthesized membrane glycoprotein in locomoting CHO cells was examined using vascular stomatis virus G-protein as a probe, Singer and co-workers (Bergmann et al., 1983; Bergmann and Singer, 1983; Kupfer et al., 1987) found that the glycoprotein molecules first appear at the cell surface of the leading edge. In addition, they showed inhibition of cell locomotion by treatment of monensin, an inhibitor of membrane transport from the Golgi apparatus to the plasma membrane, suggesting that exocytosis might be crucial for cell surface expansion at the pseudopod (Kupfer et al., 1987). All these data strongly suggest involvement of exocytosis in pseudopod formation. The exact role of exocytosis in pseudopod formation, however, still remains uncertain.

To directly examine the role of exocytosis in pseudopod formation, we attempted to detect a possible minimal unitary reaction, that is, single exocytotic events coupled with single protrusion processes in the pseudopodial region of the cell. Because of the exceptional optical properties, we employed agar-overlay methods (Kamiya and Kuroda, 1965; Naib-Majani et al., 1983; Fukui et al., 1987) combined with videoenhanced microscopy, which allowed us to visualize a single exocytosis process of vesicles containing slime, the extracellular matrix, in live Physarum plasmodia. We found that cytoplasmic protrusion in the pseudopodial region occurred intermittently and immediately after exocytosis of individual slime-containing vesicles. The protrusion was mostly filled with actin microfilaments and contained few membrane organelles, which is characteristic of pseudopods. Quantitative analysis revealed that newly inserted membrane matched well with pseudopods in terms of their surface area, timing and location. These results strongly suggest the presence of functional coupling of the single exocytotic events to the individual protrusions during pseudopod formation.

Culture and agar-overlay method

The plasmodium of Physarum polycephalum (Carolina Biological Supply Co., Burlington, USA) was fed with oatmeal and housed on damp towel paper set on glass Petri dishes in a plastic case (Camp, 1936).

For light microscope observation, we employed the agar overlay method (Kamiya and Kuroda, 1965; Naib-Majani et al., 1983; Fukui et al., 1987). Spread plasmodia were developed from the endoplasmic drops (Achenbach et al., 1979; Achenbach and WohlfarthBottermann, 1986; Ogihara and Sesaki, 1992). In brief, the plasmodia were cut at the surface with a razor blade. Because of the positive inner pressure, the endoplasm poured out to form a droplike lump at the site of the incision. With electron microscopy, we confirmed that the plasma membrane was resealed within 10 seconds. Using sharp forceps, endoplasmic drops were transferred carefully onto coverslips coated with 1.5% plain agar as a substrate. They were then covered with agar films of the same size as the coverslip. Agar films of 3% in concentration and of about 0.5 mm in thickness were used. These sandwiched plasmodia were allowed to spread in a moist chamber kept dark for about 20 hours before observation.

Video microscopy and image analysis

Plasmodia were viewed by video-enhanced differential interference contrast (DIC) microscopy using inverted Olympus IMT-2 with a ×40, 0.55 NA Olympus Plan objective, or Nikon Optiphoto-II with a ×40, 0.7 NA Nikon Plan objective. For time-lapse recordings, images were collected with a charged-coupled device camera (FCD10; Ikegami Inc., Tokyo, Japan) and stored on an s-VHS videotape. Recorded images were averaged using successive three videoframes through an image processor (Σ-III; Nippon Avionics Co., Tokyo, Japan), and digitized with a video image capture board (Videovision Studio; Radius Inc., San Jose, USA) and Adobe Premiere (Adobe System Inc., Mountain view, USA) installed in a Macintosh Quadra 950 (Apple Computer, Cupertino, USA).

For quantitation of the digitized images, NIH Image version 1.54 was used. Diameters of vesicles were determined by measurements of their cross sectional length. Their surface areas of the vesicle were calculated from their diameter, with an assumption that the vesicles are true spheres. The position of the vesicle in a cell was determined by measuring the minimum distance between the vesicle and the cell edge. The accuracy of these measurements was confirmed by measuring latex beads with a known diameter (Polyscience, Inc., Washington, USA).

Length of cytoplasmic protrusions was determined by measuring the distance from their tip to base. The base was assumed to correspond to the relatively flat cell edge of the neighboring region. Since protrusions have the plasma membrane on both the ventral and dorsal sides, and since the thickness of protrusion was negligibly small compared with the length, the surface area of protrusions was calculated by doubling the protruded area that was determined by manually outlining on digitized images.

Dark field microscopy

Plasmodia without an agar overlay were observed using a macro dark field apparatus consisting of a dissection microscope (SMZ-U, Nikon, Tokyo, Japan) and a lamp with fiber optics for light source, and recorded on an s-VHS videotape through a charged-coupled devise camera (FCD-10).

Tetramethylrhodamine-phalloidin labeling

To label surface protrusions with tetramethylrhodamine-conjugated phalloidin, agar-overlaid plasmodia were fixed in ice-cold 4% glutaraldehyde, 4% paraformaldehyde, 100 mM KCl, 50 mM EDTA, and 20 mM K-phosphate buffer at pH 7.0 for 10 minutes (Ishigami et al., 1987) after live recording. Plasmodia were further fixed for an additional 20 minutes in the 1:2 diluted fixative. Permeabilization was carried out in the 1:2 diluted fixative supplemented with 0.5% Triton X-100 for 10 minutes. Autofluorescence of glutaraldehyde was quenched with 1 mg/ml NaBH4 in TPBS (PBS containing 0.1% Tween-20) for 15 minutes. Specimens were washed in TPBS for 30 minutes, and nonspecific binding was blocked for 45 minutes in TPBS with 1% BSA. Samples were stained for 100 minutes with 200 nM tetramethylrhodamine-phalloidin (Sigma Chemical Co., St Louis, USA) dissolved in TPBS, and then washed with TPBS over a period of 60 minutes. Specimens were mounted in PBS containing 50% glycerol and 0.5% β-mercaptoethanol. Fluorescence images were acquired with a Nikon Optiphoto-II using a ×40, 0.7 NA Nikon Plan objective on Kodak Tmax film.

Electron microscopy

Spread plasmodia without an agar overlay on a 1.5% plain agar film were fixed in 2.5% glutaraldehyde, 1% OsO4 and 50 mM Na-cacodylate buffer at pH 7.0 for 1 hour at room temperature. Alternatively, plasmodia were fixed in 2% glutaraldehyde, 0.2% Triton X-100, 2 mM MgCl2, 0.2% tannic acid, and 60 mM Na-cacodylate at pH 7.3 for 1 hour at room temperature followed by post-fixation with 1% OsO4 and 40 mM Na-cacodylate at pH 6.0 for 1 hour on ice in order to preserve the fine structure of actin microfilaments. They were washed three times in water, en bloc stained for 15 minutes in 2% uranyl acetate and carried though a series of dehydration steps in ethanol and propylene oxide. Specimens were embedded in a 50:50 Epon-araldite mixture. After ultrathin sectioning, sections were stained with uranyl acetate followed by lead citrate, and observed with a JEM-1200EX electron microscope at the acceleration voltage of 80 kV.

For fixation of agar-overlaid plasmodia, another fixative was required to keep their overall morphology intact. Agar sheets covering the plasmodia decreased the efficiency of fixative penetration. Peeling the agar sheet off the plasmodia affected the morphology and hence was not done. We tried five different fixatives: (1) 4% glutaraldehyde, and 60 mM Na-cacodylate at pH 7.0; (2) 4% glutaraldehyde, 2 mM MgCl2, 0.4% tannic acid, and 60 mM Nacacodylate; (3) 4% glutaraldehyde, 0.4% Triton X-100, 2 Mm MgCl2, 0.4% tannic acid, and 60 mM Na-cacodylate; (4) 4% glutaraldehyde, 4% paraformaldehyde, 2 mM MgCl2, 0.4% tannic acid, and 60 mM Na-cacodylate; (5) 4% glutaraldehyde, 3% paraformaldehyde, 1% OsO4, 1.5 mM MgCl2, 0.3% tannic acid, and 60 mM Na-cacodylate. We placed these fixatives for agar-overlaid plasmodia on the agar film and observed their morphology using the DIC optics. Among them, we found that the no. 5 fixative fixed plasmodia most quickly without significant changes in their overall morphology. Although preservation of actin filaments was compromised by this fixation method, membrane structures were well preserved.

For morphometrical analysis, EM negatives were digitized with a high-resolution drum scanner (DT-S1030AI; Dainippon Screen Co., Kyoto, Japan) at the maximum resolution of 5200 dpi. The digital images were stored on a 128 MB magneto-optical disk with a Macintosh 660 AV (Apple Computer, Cupertino, USA). Using NIH Image, mean diameter of the vesicles was estimated from their size distribution in the ultrathin sections according to a standard stereological method of Weibel (1979).

Vesicles containing the slime in Physarum plasmodia

Fig. 1A-E shows representative ultrastructures of vesicles containing the slime, the mucous material covering the surface of the plasmodia, originally described by Stiemerling (1970). The diameter of the vesicles was 3.3±0.6 μm (average ± s.d., n=187). The vesicle often appeared to be closely attached to the plasma membrane (Fig. 1B), suggesting the presence of the docking process as shown in synaptic vesicles (Rothman, 1994). Slime in the unfused vesicle showed a heterogeneous distribution, with dense and sparse regions in places (Fig. 1A,B). In the sparse region, filamentous structures were clearly seen, and the individual filaments were 5.7±0.95 nm (n=100) in average thickness. The filaments appeared to be intertwined into a condensed network, and the extent of such condensation varied from region to region. The slime filaments observed inside the omega-shaped concavities, which have a remarkably similar degree of curvature to that of the vesicle, were distributed with more spatial heterogeneity than the vesicle contents to show numerous aggregations (Fig. C,D). The staining of the slime filaments in the concavities was more dense in electron microscopy than the filaments in the vesicle. In relatively flattened concavities, coagulation of the slime filaments was conspicuous (Fig. 1E).

Fig. 1.

Ultrastructural characterization of the slimecontaining vesicle. (A-E) Slimecontaining vesicles and a possible sequence during exocytosis. (A) Slime-containing vesicle in the cytoplasm. (B) Slime-containing vesicle docked on the plasma membrane. (C and D) Vesicles then fuse with the plasma membrane to form omegashaped concavities. (E) The openings of the concavities gradually become larger and flattened. Slime filaments become conspicuously coagulated in D, and the parallel array of the filaments is seen in E. Asterisks in C-E indicate the agar substrate. (A and C) Coated vesicles were frequently observed on the cytoplasmic surface of the slime-containing vesicle (arrows). (Inset in F) Dark field image of a locomoting plasmodium on the agar substrate. (F) Slime-containing vesicles in the apical region of a lobopod of the plasmodium. Locomoting plasmodia on agar substrate were fixed, processed for electron microscopy, and sectioned perpendicular to the substrate. The upper and the lower sides of the micrograph correspond to the dorsal and the ventral side of the plasmodia, respectively. Asterisks indicate the slime-containing vesicle. Arrowheads show omega-shaped concavities on the plasma membrane. The contour length of the flattered concavity is roughly equal to the vesicle perimeter length. Arrows indicate contractile vacuoles. Bars: 1 μm (A-E); 5 μm (F); 3 mm (inset in F).

Fig. 1.

Ultrastructural characterization of the slimecontaining vesicle. (A-E) Slimecontaining vesicles and a possible sequence during exocytosis. (A) Slime-containing vesicle in the cytoplasm. (B) Slime-containing vesicle docked on the plasma membrane. (C and D) Vesicles then fuse with the plasma membrane to form omegashaped concavities. (E) The openings of the concavities gradually become larger and flattened. Slime filaments become conspicuously coagulated in D, and the parallel array of the filaments is seen in E. Asterisks in C-E indicate the agar substrate. (A and C) Coated vesicles were frequently observed on the cytoplasmic surface of the slime-containing vesicle (arrows). (Inset in F) Dark field image of a locomoting plasmodium on the agar substrate. (F) Slime-containing vesicles in the apical region of a lobopod of the plasmodium. Locomoting plasmodia on agar substrate were fixed, processed for electron microscopy, and sectioned perpendicular to the substrate. The upper and the lower sides of the micrograph correspond to the dorsal and the ventral side of the plasmodia, respectively. Asterisks indicate the slime-containing vesicle. Arrowheads show omega-shaped concavities on the plasma membrane. The contour length of the flattered concavity is roughly equal to the vesicle perimeter length. Arrows indicate contractile vacuoles. Bars: 1 μm (A-E); 5 μm (F); 3 mm (inset in F).

Plasmodia locomoting on the agar substrate showed fanshaped morphology with well-developed lobopods (large cylindrical pseudopods; Taylor and Condeelis, 1979) and branched strand portions (Fig. 1F, inset). A conspicuous amount of secreted slime was found at the leading edge (Fig. 1F). In addition, the leading edge contains a lot of the slimecontaining vesicles (Fig. 1F, asterisks). Most of the vesicles were localized beneath the plasma membrane of lobopods. On the plasma membrane at the ventral surface of lobopods, many omega-shaped structures were seen (Fig. 1F, arrowheads), suggesting that the exocytosis of the slime-containing vesicles occurs primarily on the ventral side of lobopods.

Direct observation of exocytotic processes of the slime-containing vesicle in live plasmodia

To identify the vesicle that contains the slime in live cells, we observed the plasmodia with video microscopy and then fixed them for electron microscopy (Fig. 2). Referring to the positions of contractile vacuoles (Fig. 2D, asterisks), we identified the slime-containing vesicle in ultrathin sections (Fig. 2D), which had corresponding vesicular morphologies in the videoimages (Fig. 2B). As shown in Fig. 2C, these vesicles moved at about 0.2 μm/second in the cytoplasm, faster than contractile vacuoles that were not significantly translocated during the same observation period. Consistent with electron microscope observation, the diameters of the slime-containing vesicles were 3-4 μm in the videoimages. No other organelles with such a large diameter were found in the cytoplasm except for the nuclei and the contractile vacuole. Although the size of contractile vacuoles was similar to that of the slime-containing vesicle, they could be readily distinguished due to their characteristic ultrastructures such as irregular contour of their membrane surface and the absence of visible contents, and to their unique behavior repeating cycles of contraction and swelling (see details in the next paragraph).

Fig. 2.

Identification of the slime-containing vesicle in live plasmodia. Agar-overlaid plasmodia were observed both with video microscopy (A and B) and electron microscopy (D). The slimecontaining vesicles (circles) translocated in live plasmodia for 50 seconds as shown in A and B, and the tracks of displacement between A and B are shown in C (filled circles) with 10 secondintervals. Four contractile vacuoles (asterisks in A and B) were virtually immobile (C; open circles). (D) Ultrastructure of the live recorded region of the plasmodium. Immediately after light microscope observation in B, the plasmodium was fixed, and sectioned parallel to the substrate. Circles in D indicate slimecontaining vesicles observed in videoimages. Asterisks in D indicate contractile vacuoles which match those in C. Bars, 10 μm.

Fig. 2.

Identification of the slime-containing vesicle in live plasmodia. Agar-overlaid plasmodia were observed both with video microscopy (A and B) and electron microscopy (D). The slimecontaining vesicles (circles) translocated in live plasmodia for 50 seconds as shown in A and B, and the tracks of displacement between A and B are shown in C (filled circles) with 10 secondintervals. Four contractile vacuoles (asterisks in A and B) were virtually immobile (C; open circles). (D) Ultrastructure of the live recorded region of the plasmodium. Immediately after light microscope observation in B, the plasmodium was fixed, and sectioned parallel to the substrate. Circles in D indicate slimecontaining vesicles observed in videoimages. Asterisks in D indicate contractile vacuoles which match those in C. Bars, 10 μm.

To further confirm whether or not these vesicles that were visible with video microscopy are the vesicles secreting the slime, we attempted to observe exocytotic processes of these vesicles in live plasmodia. Taking advantage of agar-overlay methods in the optical properties, we were able to observe the disappearance of these vesicles in the cytoplasm (Fig. 3A).

Fig. 3.

Shrinkage of a slime-containing vesicle. (A) Timelapse images of the slime-containing vesicle during its disappearance. The vesicles in agar-overlaid plasmodia were recorded with video-enhanced microscopy. Recording starts at the upper left image and proceeds at 1/6-second intervals to the lower right image. (B) Representative time course of the changes in diameter of the slime-containing vesicle. Inset shows the diameter changes of a contractile vacuole during representative contraction. (C) Average rate of the changes in diameter of the slime-containing vesicle and the contractile vacuole. Error bars indicate s.d. (n=7). Bar in A, 10 μm.

Fig. 3.

Shrinkage of a slime-containing vesicle. (A) Timelapse images of the slime-containing vesicle during its disappearance. The vesicles in agar-overlaid plasmodia were recorded with video-enhanced microscopy. Recording starts at the upper left image and proceeds at 1/6-second intervals to the lower right image. (B) Representative time course of the changes in diameter of the slime-containing vesicle. Inset shows the diameter changes of a contractile vacuole during representative contraction. (C) Average rate of the changes in diameter of the slime-containing vesicle and the contractile vacuole. Error bars indicate s.d. (n=7). Bar in A, 10 μm.

They gradually shrank in diameter while keeping the roundness in shape, and finally became undetectable. As shown in Fig. 3B and C, the diameter shortened linearly and the shortening rate was 1.1±0.3 μm/second (s.d., n=7). After disappearance, the vesicles were not found in the other focal planes at the same spot. This indicates that vertical translocation of the vesicles did not occur. On the other hand, the average rate of contraction of contractile vacuoles was 12±5.0 μm/second (n=7), ten times as fast as the shrinkage rate of the slime-containing vesicle (Fig. 3C). The slime-containing vesicles never swelled up again after they disappeared once, whereas contractile vacuoles repeated contraction and swelling (Fig. 3B, inset). It is, therefore, probable that the disappearance of the slime-containing vesicle is accounted for by exocytosis. Consistent with this notation, disappearance of secretory vesicles during exocytosis has been reported in chromaffin cells (Edwards et al., 1984; Terakawa et al., 1991), and salivary grand acinar cells (Segawa et al., 1991). Interestingly, the slime-containing vesicles occasionally fused with each other in the cytoplasm before exocytosis. The fused vesicles became larger in diameter than unfused vesicles (data not shown). Fusion between secretory vesicles has been reported previously in mast cells (Lawson et al., 1977).

Cell surface protrusion after exocytosis of the slimecontaining vesicle

When the slime-containing vesicles disappeared at the cell edge, a part of the neighboring cell surface protruded (Fig. 4). We observed a total of 246 cases of exocytosis, and found that the onset of such protrusion was within 5 seconds of vesicle disappearance in 134 cases. The protruded region showed lamella-like morphology, and did not appear, in most cases, to contain intracellular granular organelles visible by videoenhanced DIC optics. The rate of protrusion was ∼1 μm/second, which is quite similar to the rate of pseudopod formation in plasmodia previously reported by Ueda and Kobatake (1978).

Fig. 4.

Surface protrusion after exocytosis of the slime-containing vesicle in agar-overlaid plasmodia. The sequence starts at the upper left image and proceeds at 0.6-second intervals to the lower right image. The arrowhead indicates the slime-containing vesicle, and the arrow indicates the onset of protrusion. Bar, 10 μm.

Fig. 4.

Surface protrusion after exocytosis of the slime-containing vesicle in agar-overlaid plasmodia. The sequence starts at the upper left image and proceeds at 0.6-second intervals to the lower right image. The arrowhead indicates the slime-containing vesicle, and the arrow indicates the onset of protrusion. Bar, 10 μm.

Since one of the most important characteristics of the pseudopod is the presence of actin microfilaments (Condeelis, 1993), we examined microfilament distribution in newly formed protrusions. Plasmodia were fixed immediately after protrusion following exocytosis, as shown in Fig. 5A-D. Flu-orescent phalloidin staining demonstrates that the cytoplasmic protrusion contains actin microfilaments (Fig. 5E). The intensity was relatively weak in this protrusion compared to neighboring regions, probably reflecting the difference in thickness between the protrusion and the remaining cell body. To characterize the ultrastructural basis of the protrusion, we fixed cells for electron microscopy immediately after protrusion had occurred (Fig. 6). In the protruded region (Fig. 6G), most of the membrane organelles were excluded, suggesting that their main constituent is actin microfilaments, taken together with phalloidin staining as shown above. The protrusions displayed ultrastructures that showed continuous texture from the cortex to the protrusion. The cortical region contains the actin microfilament meshwork (Naib-Majani et al., 1983), although OsO4 contained in the fixative we used destroyed fine details of microfilament structures, as reported previously (Small, 1981). The presence of actin filaments in protrusions at the cell edge was confirmed by the use of another fixative with different constituents as follows: 2% glutaraldehyde, 0.2% Triton X-100, 2 mM MgCl2, 0.2% tannic acid, and 60 mM Na-cacodylate at pH 7.3 followed by post-fixation with 1% OsO4 and 40 mM Na-cacodylate at pH 6.0. Plasmodia were not agar-overlaid. Actin microfilaments were excellently preserved and their organization was clearly demonstrated to be meshwork filling the entire space of the protrusions (Fig. 7). By contrast, in the region between the cortex and the granular cytosol, a distinct pattern of ultrastructure running parallel to the cell edge was seen (Fig. 6G, arrows) and it probably represents microfilament bundles (Ishigami et al., 1981). These observations indicate that the protrusion contains an actin microfilament meshwork without conspicuous membrane organelles.

Fig. 5.

Protrusion contains actin microfilaments. Agar-overlaid plasmodia were observed with video-enhanced microscopy: (A) 0, (B) 4, and (C) 11 seconds. The same plasmodia were fixed at 18 seconds (D) and stained with tetramethylrhodamine-conjugated phalloidin (E). The arrowhead in A indicates the slime-containing vesicle before exocytosis. Bar, 10 μm.

Fig. 5.

Protrusion contains actin microfilaments. Agar-overlaid plasmodia were observed with video-enhanced microscopy: (A) 0, (B) 4, and (C) 11 seconds. The same plasmodia were fixed at 18 seconds (D) and stained with tetramethylrhodamine-conjugated phalloidin (E). The arrowhead in A indicates the slime-containing vesicle before exocytosis. Bar, 10 μm.

Fig. 6.

Ultrastructural characterization of the protrusion. Agar-overlaid plasmodia were observed with video microscopy (A-E) and then fixed for electron microscopy (F-H). (A-D) The slime-containing vesicle (arrow in A) was exocytosed and surface protrusion (arrows in B-E) followed. Note the hyaline appearance of the protrusion. (F-H) Ultrathin sections of the same plasmodia sectioned parallel to the substrate. Fixation was done about 5 seconds after E. Three sections are chosen so that the one in F is furthest to the substrate, the one in H is the closest, and the one in G is between. The cell edge marked with a bar in F roughly corresponds to such a region in G and the protrusion in H. Black asterisks in E and F indicate unidentified vesicular structures observed both in the videoimage and electron micrograph. The white asterisk indicates the slime-containing vesicle. In the region between the cortex and the granular cytosol, distinct pattern of ultrastructure which probably represents microfilament bundles (Ishigami et al., 1981) runs parallel to the cell edge (arrows in G). Bars: 5 μm (E,F,G); 2 μm (H).

Fig. 6.

Ultrastructural characterization of the protrusion. Agar-overlaid plasmodia were observed with video microscopy (A-E) and then fixed for electron microscopy (F-H). (A-D) The slime-containing vesicle (arrow in A) was exocytosed and surface protrusion (arrows in B-E) followed. Note the hyaline appearance of the protrusion. (F-H) Ultrathin sections of the same plasmodia sectioned parallel to the substrate. Fixation was done about 5 seconds after E. Three sections are chosen so that the one in F is furthest to the substrate, the one in H is the closest, and the one in G is between. The cell edge marked with a bar in F roughly corresponds to such a region in G and the protrusion in H. Black asterisks in E and F indicate unidentified vesicular structures observed both in the videoimage and electron micrograph. The white asterisk indicates the slime-containing vesicle. In the region between the cortex and the granular cytosol, distinct pattern of ultrastructure which probably represents microfilament bundles (Ishigami et al., 1981) runs parallel to the cell edge (arrows in G). Bars: 5 μm (E,F,G); 2 μm (H).

Fig. 7.

Organization of actin microfilaments in the protrusion. Spread plasmodia without an agar overlay were fixed for electron microscopy. See Materials and Methods for details in the fixative. A random meshwork pattern of actin microfilaments is well preserved. Bar, 0.5 μm.

Fig. 7.

Organization of actin microfilaments in the protrusion. Spread plasmodia without an agar overlay were fixed for electron microscopy. See Materials and Methods for details in the fixative. A random meshwork pattern of actin microfilaments is well preserved. Bar, 0.5 μm.

Among the ultrathin sections of the newly protruded portion of the cell, the section close to the substrate contained the proximal part to the distal part of the protrusion (Fig. 6H). Judging from the overall morphology, the distal portion of the protrusion with the round shape in Fig. 6H appears to correspond to the protrusion in Fig. 6G. This means that the proximal part of this protrusion is hidden by the overlaying cytoplasm in Fig. 6G. Since the vesicle in live plasmodia disappeared 4.6 μm from the cell edge, and since this distance roughly corresponded to the length of the proximal part of the protrusion, it is suggested that the protrusion started where the vesicle disappeared. Interestingly, the protrusion appears to push the secreted slime outward, judging from the stretched appearance of the slime around the tip of the protrusion.

The slime-containing vesicles invaded the hyaline layer (Fig. 8A-E) which contains a dense microfilament meshwork. In the hyaline region, slime-containing vesicles disappeared, and cytoplasmic protrusion occurred. Moreover, electron microscopy confirmed that the slime-containing vesicles (Fig. 8F, asterisks) were often found in the cortical layer just beneath the plasma membrane, which corresponds to the hyaline layer, whereas most other organelles were not found in this region.

Fig. 8.

Invasion of the slimecontaining vesicle into the hyaline layer before exocytosis. (A-E) Videoimages showing that the slime-containing vesicle (arrows) invades the hyaline layer on the cell edge of agaroverlaid plasmodia. The vesicle disappeared in the hyaline layer, and protrusion (arrowheads) was observed at the site of exocytosis. Images were taken during a period of 81 seconds. (F).Electron micrograph showing the presence of the slimecontaining vesicle (asterisks) near the cortical region just beneath the plasma membrane. The one on the right has invaded the cortical regions, the one on the left is found in the granuloplasm, and the one inbetween lies in the intermediate zone. Agar-overlaid plasmodia locomoting on the agar substrate were fixed, processed for electron microscopy, and sectioned parallel to the substrate. Note that vesicular organelles other than the slime-containing vesicles are densely confined in the granuloplasm. Bars: 10 μm (E); 2 μm (F).

Fig. 8.

Invasion of the slimecontaining vesicle into the hyaline layer before exocytosis. (A-E) Videoimages showing that the slime-containing vesicle (arrows) invades the hyaline layer on the cell edge of agaroverlaid plasmodia. The vesicle disappeared in the hyaline layer, and protrusion (arrowheads) was observed at the site of exocytosis. Images were taken during a period of 81 seconds. (F).Electron micrograph showing the presence of the slimecontaining vesicle (asterisks) near the cortical region just beneath the plasma membrane. The one on the right has invaded the cortical regions, the one on the left is found in the granuloplasm, and the one inbetween lies in the intermediate zone. Agar-overlaid plasmodia locomoting on the agar substrate were fixed, processed for electron microscopy, and sectioned parallel to the substrate. Note that vesicular organelles other than the slime-containing vesicles are densely confined in the granuloplasm. Bars: 10 μm (E); 2 μm (F).

Quantitative relationships between exocytosis and protrusion

To determine spatial and temporal relationships between the disappearance of the slime-containing vesicle and the protrusion of the cell surface, quantitative analyses were performed. Fig. 9A shows a representative time course of the vesicle disappearance and surface protrusion. Protrusion started immediately after the disappearance of the slime-containing vesicle; no such protrusion was observed preceding the disappearance. In general, protrusions came out from the cell edge with time lags ranging from 0 to 5 seconds after disappearance of the slime-containing vesicle (Fig. 9B).

Fig. 9.

Quantitative analysis of the relationship between exocytosis and protrusion. (A) Representative timecourse of the slimecontaining vesicle shrinkage and protrusion. Filled circles indicate diameter of the vesicle. Open circles indicate length of the protrusion. (B) Time lag between exocytosis and protrusion is virtually proportional to the position of the slime vesicle from the cell edge. The time lags were determined using videoimages with 0.25 second intervals. (C) Surface area of protrusions is proportional to that of the slime-containing vesicle. (D) Ratio of surface areas between protrusion and vesicle depends on the position of vesicles from the cell edge.

Fig. 9.

Quantitative analysis of the relationship between exocytosis and protrusion. (A) Representative timecourse of the slimecontaining vesicle shrinkage and protrusion. Filled circles indicate diameter of the vesicle. Open circles indicate length of the protrusion. (B) Time lag between exocytosis and protrusion is virtually proportional to the position of the slime vesicle from the cell edge. The time lags were determined using videoimages with 0.25 second intervals. (C) Surface area of protrusions is proportional to that of the slime-containing vesicle. (D) Ratio of surface areas between protrusion and vesicle depends on the position of vesicles from the cell edge.

If the membrane of a newly formed protrusion was supplied by fusion of the slime-containing vesicle with the plasma membrane, the surface area of the protrusion would be correlated to the surface area of the vesicle. To test for this possibility, the surface area of individual vesicles was calculated from their diameter assuming that their shapes are true spheres in the cytoplasm. Surface area of protrusions was determined by doubling the protruded area. Fig. 9C shows a virtually linear relationship between the surface area of the protrusion and that of the slime-containing vesicle. Furthermore, the ratio of protruded area to vesicle surface area showed values ranging from 0.3 to 1.0, depending on the location of vesicles (Fig. 9D). The further a vesicle was located from the cell edge, the less a protruded surface area became. Collectively all these data support the idea that membrane materials of the slime-containing vesicle were used as the plasma membrane of protrusions during exocytosis.

We visualized single exocytotic events, and correlated them to the process of cell surface protrusion. Exocytosis was observed with video-enhanced microscopy combined with agar-overlay methods applied to live plasmodia (Figs 3, 4, 5, 6 and 8). The exocytotic vesicle contains and secretes slime, as revealed by electron microscopy (Fig. 1). Slime is the extracellular matrix that consists of D-galactan with an average degree of polymerization of 560 (Henney, 1982). Preceding exocytosis, the slime-containing vesicle selectively invaded the hyaline layer which consists of the actin microfilament meshwork (Fig. 8). No other vesicular structures were found in the hyaline layer. This is quite an important feature of the slime-containing vesicles, which enables the vesicles to access to the plasma membrane before fusion. Cell surface in the neighboring region protruded at the site where exocytosis occurred (Figs 4, 5, 6 and 8). The protrusion rate was quite similar to the rate of pseudopod extension previously reported (Ueda and Kobatake, 1978). Protruded regions then formed a new hyaline layer. They contained actin microfilaments (Fig. 5). The organization of microfilaments was suggested to be meshwork of randomly oriented filaments rather than straight bundles (Figs 6 and 7). These results, taken together, suggest that Physarum plasmodia extend individual pseudopods after single exocytotic events of the slime-containing vesicle. Such protrusion, coupled with exocytosis of the slime-containing vesicle, may be the minimal process which underlies the pseudopod formation, and an assembly of many protrusions could lead to formation of a large pseudopod called a lobopod (Taylor and Condeelis, 1979) observed in Physarum plasmodia.

While surface protrusion that is coupled with exocytosis of the slime-containing vesicle showed the characteristics of pseudopods as discussed above, it is not clear whether or not these protrusions are directly involved in locomotion of the plasmodium. For example, it has been shown that not all pseudopods that extend from the cell surface lead to cell migration (Condeelis, 1993). Also a rearward transport process of the surface structure (Abercrombie et al., 1970; Heath and Holifield, 1991), if present in the Physarum plasmodium, would considerably reduce the contribution of the surface protrusion to the net migration of the plasmodium. There is also a possibility that the protrusion functions merely in the cell spreading process, not tightly associated with the net migration which obviously requires the coupling with retraction of the posterior part of the plasmodium.

The shortening rate of the diameter of the slime-containing vesicle during exocytosis was ten times as slow as the contraction rate of the contractile vacuole (Fig. 3D), although their diameter was quite similar. Such a rate difference between the exocytotic vesicle and the contractile vacuole revealed in this article strongly supports a proposal for different mechanisms underlying these two types of membrane deformation. Contraction of the contractile vacuoles could be driven by an ATPdependent active process involving actin microfilaments and myosin I, as reported recently (Doberstein et al., 1993). On the other hand, in exocytosis, resumption of the planar nature of the plasma membrane after fusion of vesicles and formation of the omega shape as an intermediate state of the exocytosis could be a passive process probably due to the intrinsic nature of the membrane including surface tension and/or to positive inner pressure of the cytoplasm (Kamiya, 1959). Alternatively, the slow rate of the slime-containing vesicle shrinkage could be explained by the viscosity of the contents to be extruded. Slime is a branched filamentous structure as shown in the electron micrograph in Fig. 1 (Wolf et al., 1981). Biochemical characterization (Henney, 1982) confirms its branched structure and hence its viscous nature. Consistent with this notion, the slow rate of exocytosis in general has been explained by a difference in the diffusibility of the vesicle contents in acinar cells and goblet cells which secrete the viscous mucus (Segawa et al., 1991).

A series of experiments has raised the hypothesis that exocytosis is required for pseudopod formation in locomoting cells (Bretscher, 1983, 1984; Bergmann et al., 1983; Bretscher and Thomson, 1983; Singer and Kupfer, 1986; Kupfer et al., 1987). This hypothesis would predict some quantitative relationships between exocytosis and cell surface extension, if we assume that insertion of the new membrane materials via exocytosis directly allows the cell surface to expand. One of the critical questions to be addressed has been whether or not the amount of membrane materials supplied by exocytosis can explain the expansion of the plasma membrane at the site of the pseudopod. But this question has not been so far directly examined, because the size of the exocytotic vesicle is generally too small to be correlated to pseudopod formation. Here we directly correlated single exocytosis to pseudopod formation in a quantitative manner, taking advantage of the fact that the slime-containing vesicles are extremely large and hence visible with light microscopy. Also the proximity of the exocytosis site and the protrusion site is an advantage of the plasmodium. Protruded areas were substantially proportional to the surface area of the vesicle (Fig. 9C) and did not exceed the membrane area that was supplied by exocytosis (Fig. 9D). Calculation of the surface areas of the slime vesicle and the protrusion based on their morphologies could be exact enough to obtain accurate values, firstly because the slime-containing vesicle is almost a complete sphere with a smooth membrane contour, and secondly because the protrusion has negligible thickness and a smooth membrane contour. These results strongly indicate that the slime-containing vesicle supplied the pre-existing plasma membrane with new membrane materials to expand its surface during pseudopod formation.

It is of interest to understand the geometrical relationship between exocytosis and protrusion. Do protrusions really start at the site of exocytosis? To answer this question, we tried to correlate the site of the exocytosis to that of the protrusion. Some protrusions started obviously at the region where the slime-containing vesicle disappeared, i.e. in which exocytosis occurred at the very edge of the plasmodia. In the other cases in which the slime-containing vesicles disappeared at sites relatively distant from the cell edge, not all the sites of protrusion were distinct. Since protrusion at the cell edge occurred immediately after the exocytosis (Fig. 9A), the observed time lags in exocytosis distant from the cell edge allowed us to estimate how far the initiation site of the protrusion is from the cell edge. The time lag was in a range from 0 to 5 seconds (Fig. 3B), and can be converted to distance in a range from 0 to 5 μm, taking the protrusion rate to be 1 μm/second (Fig. 9A). These values matched well with the distance between where the slime containing vesicle disappeared and where protrusions came out from the cell edge. In addition to these quantitative analyses, electron microscopy suggests that these time lags could be explained by three-dimensional topography of the cell edge which renders protrusions invisible in a top view using the light microscope until the protrusion comes out from the edge after a certain delay. In fact, the proximal part of the protrusion is hidden by overlying cytoplasm, when exocytosis occurred away from the cell edge (Fig. 6). Collectively, it is highly plausible that protrusion started at the site where exocytosis occurred.

The authors thank Hiroshi Ike, Ken-ichi Jono, Yoshikatsu Sato and Asaka Ito of the International Christian University for morphometry, and Jeff Silverstein of the University of Washington for critical reading of the manuscript.

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