809Journal of Cell Science 110, 809-818 (1997)Printed in Great Britain © The Company of Biologists Limited 1997JCS9507

Protrusion of cell surface coupled with single exocytotic events of secretion

of the slime in Physarum plasmodia

Hiromi Sesaki* and Satoshi Ogihara†

Department of Biology, Graduate School of Science, Osaka University, Toyonaka, Osaka 560, Japan

*Present address: Charles H. Best Institute, University of Toronto, 112 College Street, Toronto, Ontario, Canada M5G 1L6†Author for correspondence (e-mail: ogihara@bio.sci.osaka-u.ac.jp)

Exocytosis has been proposed to participate in theformation of pseudopods. Using video-enhancedmicroscopy, we directly visualized exocytosis of singlevesicles in living Physarum plasmodia migrating on asubstrate. Vesicles containing slime, the plasmodial extra-cellular matrix, of ~3.5 µµm in diameter, shrank at the cellperiphery at the average rate of ~1 µµm/second, and becameinvisible. Immediately after exocytotic events, the neigh-boring cell surface extended to form a protrusion. The rateof extension was ~1 µµm/second. The protrusion showedlamella-like morphology, and contained actin microfila-ments. Electron microscopy suggested that the organiz-ation of microfilaments in such protrusions may be arandom meshwork rather than straight bundles. These

morphologies suggest that protruded regions arepseudopods. Importantly, only the slime-containing vesiclepreferentially invaded the hyaline layer that consists ofdense actin microfilaments while the other vesicularorganelles remained in the granuloplasm. Quantitativeanalysis demonstrated a linear relationship in terms oftheir surface area, between individual protrusions andsingle slime-containing vesicles. It is, therefore, likely thatmost of the plasma membrane of the protrusion wassupplied by fusion of the slime-containing vesicle duringexocytosis.

Key words: Pseudopod, Exocytosis, Physarum, Slime, Extracellularmatrix

SUMMARY

INTRODUCTION

Exocytosis is a fundamental process that is seen in all eukary-otic cells, and plays important roles in cell physiology. It is themechanism underlying the following two functions of the cell.The first is secretion whereby contents of intracellularmembrane vesicles are released to the extracellular space. Theother is supply of new membrane materials to the plasmamembrane including proteins and lipids. Macromoleculessupplied in this way increase the surface area of the plasmamembrane. These two functions of exocytosis, i.e. vesiclecontents delivery and membrane material addition, give rise tocell shape changes in a cooperative manner in various types ofcells. Examples include pseudopod extension (Singer andKupfer, 1986; Bretscher, 1984), neurite elongation (Bray,1970; Popov et al., 1993), cytokinesis (Bluemink and Delaat,1977), and yeast budding (Drubin, 1991). The site of exocyto-sis has been suggested to be located at the growth cone ofgrowing 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 exocytoticevents imply a role for exocytotic vesicles in cell shape changesas a supplier of new membrane materials and the extracellularcomponents to specific regions in cells, where dramaticexpansion of the plasma membrane and interactions with thefreshly secreted extracellular components are taking place.

In a variety of cell types that locomote on substrate, theleading edge has been suggested to be a site for membraneinsertion. Bretscher and co-workers (Bretscher, 1983;Bretscher and Thomson, 1983) showed a graded distributionof surface receptors with remarkable enrichment in the cellperiphery of HeLa cells. Hopkins (1985) showed that themajority of the newly appearing transferrin receptors on thecell surface arise near the cell margin of A431 cells usingimmuno-gold labeling. Since these receptors are transportedto the cell surface by exocytotic mechanisms (Goldstein etal., 1985), these results suggest localized exocytosisoccurring at the cell periphery. When the surface distributionof the newly synthesized membrane glycoprotein in loco-moting CHO cells was examined using vascular stomatisvirus G-protein as a probe, Singer and co-workers (Bergmannet al., 1983; Bergmann and Singer, 1983; Kupfer et al., 1987)found that the glycoprotein molecules first appear at the cellsurface of the leading edge. In addition, they showed inhibi-tion of cell locomotion by treatment of monensin, an inhibitorof membrane transport from the Golgi apparatus to theplasma membrane, suggesting that exocytosis might becrucial for cell surface expansion at the pseudopod (Kupferet al., 1987). All these data strongly suggest involvement ofexocytosis in pseudopod formation. The exact role of exocy-tosis in pseudopod formation, however, still remainsuncertain.

To directly examine the role of exocytosis in pseudopod

810 H. Sesaki and S. Ogihara

formation, we attempted to detect a possible minimal unitaryreaction, that is, single exocytotic events coupled with singleprotrusion processes in the pseudopodial region of the cell.Because of the exceptional optical properties, we employedagar-overlay methods (Kamiya and Kuroda, 1965; Naib-Majani et al., 1983; Fukui et al., 1987) combined with video-enhanced microscopy, which allowed us to visualize a singleexocytosis process of vesicles containing slime, the extracel-lular matrix, in live Physarum plasmodia. We found that cyto-plasmic protrusion in the pseudopodial region occurred inter-mittently and immediately after exocytosis of individualslime-containing vesicles. The protrusion was mostly filledwith actin microfilaments and contained few membraneorganelles, which is characteristic of pseudopods. Quantita-tive analysis revealed that newly inserted membrane matchedwell with pseudopods in terms of their surface area, timingand location. These results strongly suggest the presence offunctional coupling of the single exocytotic events to the indi-vidual protrusions during pseudopod formation.

MATERIALS AND METHODS

Culture and agar-overlay methodThe plasmodium of Physarum polycephalum (Carolina BiologicalSupply Co., Burlington, USA) was fed with oatmeal and housed ondamp towel paper set on glass Petri dishes in a plastic case (Camp,1936).

For light microscope observation, we employed the agar overlaymethod (Kamiya and Kuroda, 1965; Naib-Majani et al., 1983; Fukuiet al., 1987). Spread plasmodia were developed from the endoplas-mic drops (Achenbach et al., 1979; Achenbach and Wohlfarth-Bottermann, 1986; Ogihara and Sesaki, 1992). In brief, theplasmodia were cut at the surface with a razor blade. Because of thepositive inner pressure, the endoplasm poured out to form a drop-like lump at the site of the incision. With electron microscopy, weconfirmed that the plasma membrane was resealed within 10seconds. Using sharp forceps, endoplasmic drops were transferredcarefully onto coverslips coated with 1.5% plain agar as a substrate.They were then covered with agar films of the same size as thecoverslip. Agar films of 3% in concentration and of about 0.5 mmin thickness were used. These sandwiched plasmodia were allowedto spread in a moist chamber kept dark for about 20 hours beforeobservation.

Video microscopy and image analysis Plasmodia were viewed by video-enhanced differential interferencecontrast (DIC) microscopy using inverted Olympus IMT-2 with a×40, 0.55 NA Olympus Plan objective, or Nikon Optiphoto-II witha ×40, 0.7 NA Nikon Plan objective. For time-lapse recordings,images were collected with a charged-coupled device camera (FCD-10; Ikegami Inc., Tokyo, Japan) and stored on an s-VHS videotape.Recorded images were averaged using successive three videoframesthrough an image processor (∑-III; Nippon Avionics Co., Tokyo,Japan), and digitized with a video image capture board (VideovisionStudio; Radius Inc., San Jose, USA) and Adobe Premiere (AdobeSystem Inc., Mountain view, USA) installed in a Macintosh Quadra950 (Apple Computer, Cupertino, USA).

For quantitation of the digitized images, NIH Image version 1.54was used. Diameters of vesicles were determined by measurements oftheir cross sectional length. Their surface areas of the vesicle werecalculated from their diameter, with an assumption that the vesiclesare true spheres. The position of the vesicle in a cell was determinedby measuring the minimum distance between the vesicle and the celledge. 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 measuringthe distance from their tip to base. The base was assumed to corre-spond to the relatively flat cell edge of the neighboring region. Sinceprotrusions have the plasma membrane on both the ventral and dorsalsides, and since the thickness of protrusion was negligibly smallcompared with the length, the surface area of protrusions was calcu-lated by doubling the protruded area that was determined by manuallyoutlining on digitized images.

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

Tetramethylrhodamine-phalloidin labelingTo label surface protrusions with tetramethylrhodamine-conjugatedphalloidin, 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 furtherfixed for an additional 20 minutes in the 1:2 diluted fixative. Per-meabilization was carried out in the 1:2 diluted fixative supple-mented with 0.5% Triton X-100 for 10 minutes. Autofluorescenceof glutaraldehyde was quenched with 1 mg/ml NaBH4 in TPBS(PBS containing 0.1% Tween-20) for 15 minutes. Specimens werewashed in TPBS for 30 minutes, and nonspecific binding wasblocked for 45 minutes in TPBS with 1% BSA. Samples werestained for 100 minutes with 200 nM tetramethylrhodamine-phal-loidin (Sigma Chemical Co., St Louis, USA) dissolved in TPBS, andthen washed with TPBS over a period of 60 minutes. Specimenswere mounted in PBS containing 50% glycerol and 0.5% β-mer-captoethanol. Fluorescence images were acquired with a NikonOptiphoto-II using a ×40, 0.7 NA Nikon Plan objective on Kodak T-max film.

Electron microscopySpread plasmodia without an agar overlay on a 1.5% plain agar filmwere fixed in 2.5% glutaraldehyde, 1% OsO4 and 50 mM Na-caco-dylate buffer at pH 7.0 for 1 hour at room temperature. Alternatively,plasmodia were fixed in 2% glutaraldehyde, 0.2% Triton X-100, 2mM MgCl2, 0.2% tannic acid, and 60 mM Na-cacodylate at pH 7.3for 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 orderto preserve the fine structure of actin microfilaments. They werewashed three times in water, en bloc stained for 15 minutes in 2%uranyl acetate and carried though a series of dehydration steps inethanol and propylene oxide. Specimens were embedded in a 50:50Epon-araldite mixture. After ultrathin sectioning, sections werestained with uranyl acetate followed by lead citrate, and observedwith a JEM-1200EX electron microscope at the acceleration voltageof 80 kV.

For fixation of agar-overlaid plasmodia, another fixative wasrequired to keep their overall morphology intact. Agar sheetscovering the plasmodia decreased the efficiency of fixative penetra-tion. Peeling the agar sheet off the plasmodia affected the morphol-ogy 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 Na-cacodylate; (3) 4% glutaraldehyde, 0.4% Triton X-100, 2 mMMgCl2, 0.4% tannic acid, and 60 mM Na-cacodylate; (4) 4% glu-taraldehyde, 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

811Pseudopod formation coupled with exocytosis

60 mM Na-cacodylate. We placed these fixatives for agar-overlaidplasmodia on the agar film and observed their morphology using theDIC optics. Among them, we found that the no. 5 fixative fixedplasmodia most quickly without significant changes in their overallmorphology. Although preservation of actin filaments was compro-mised by this fixation method, membrane structures were wellpreserved.

For morphometrical analysis, EM negatives were digitized with ahigh-resolution drum scanner (DT-S1030AI; Dainippon Screen Co.,Kyoto, Japan) at the maximum resolution of 5200 dpi. The digitalimages were stored on a 128 MB magneto-optical disk with aMacintosh 660 AV (Apple Computer, Cupertino, USA). Using NIHImage, mean diameter of the vesicles was estimated from their sizedistribution in the ultrathin sections according to a standard stereo-logical method of Weibel (1979).

Fig. 1. Ultrastructuralcharacterization of the slime-containing vesicle. (A-E) Slime-containing vesicles and apossible sequence duringexocytosis. (A) Slime-containingvesicle in the cytoplasm.(B) Slime-containing vesicledocked on the plasmamembrane. (C and D) Vesiclesthen fuse with the plasmamembrane to form omega-shaped concavities. (E) Theopenings of the concavitiesgradually become larger andflattened. Slime filamentsbecome conspicuouslycoagulated in D, and the parallelarray of the filaments is seen inE. Asterisks in C-E indicate theagar substrate. (A and C) Coatedvesicles were frequentlyobserved on the cytoplasmicsurface of the slime-containingvesicle (arrows). (Inset in F)Dark field image of a locomotingplasmodium on the agarsubstrate. (F) Slime-containingvesicles in the apical region of alobopod of the plasmodium.Locomoting plasmodia on agarsubstrate were fixed, processedfor electron microscopy, andsectioned perpendicular to thesubstrate. The upper and thelower sides of the micrographcorrespond to the dorsal and theventral side of the plasmodia,respectively. Asterisks indicatethe slime-containing vesicle.Arrowheads show omega-shapedconcavities on the plasmamembrane. The contour lengthof the flattered concavity isroughly equal to the vesicleperimeter length. Arrowsindicate contractile vacuoles.Bars: 1 µm (A-E); 5 µm (F);3 mm (inset in F).

RESULTS

Vesicles containing the slime in PhysarumplasmodiaFig. 1A-E shows representative ultrastructures of vesicles con-taining the slime, the mucous material covering the surface ofthe plasmodia, originally described by Stiemerling (1970). Thediameter of the vesicles was 3.3±0.6 µm (average ± s.d.,n=187). The vesicle often appeared to be closely attached tothe plasma membrane (Fig. 1B), suggesting the presence of thedocking process as shown in synaptic vesicles (Rothman,1994). Slime in the unfused vesicle showed a heterogeneousdistribution, with dense and sparse regions in places (Fig.1A,B). In the sparse region, filamentous structures were clearly

812 H. Sesaki and S. Ogihara

Fig. 2. Identification of the slime-containing vesicle in liveplasmodia. Agar-overlaid plasmodia were observed both with videomicroscopy (A and B) and electron microscopy (D). The slime-containing vesicles (circles) translocated in live plasmodia for 50seconds as shown in A and B, and the tracks of displacementbetween A and B are shown in C (filled circles) with 10 second-intervals. Four contractile vacuoles (asterisks in A and B) werevirtually immobile (C; open circles). (D) Ultrastructure of the liverecorded region of the plasmodium. Immediately after lightmicroscope observation in B, the plasmodium was fixed, andsectioned parallel to the substrate. Circles in D indicate slime-containing vesicles observed in videoimages. Asterisks in D indicatecontractile vacuoles which match those in C. Bars, 10 µm.

seen, and the individual filaments were 5.7±0.95 nm (n=100)in average thickness. The filaments appeared to be intertwinedinto a condensed network, and the extent of such condensationvaried from region to region. The slime filaments observedinside the omega-shaped concavities, which have a remarkablysimilar degree of curvature to that of the vesicle, were distrib-uted with more spatial heterogeneity than the vesicle contentsto show numerous aggregations (Fig. C,D). The staining of theslime filaments in the concavities was more dense in electronmicroscopy than the filaments in the vesicle. In relativelyflattened concavities, coagulation of the slime filaments wasconspicuous (Fig. 1E).

Plasmodia locomoting on the agar substrate showed fan-shaped morphology with well-developed lobopods (largecylindrical pseudopods; Taylor and Condeelis, 1979) andbranched strand portions (Fig. 1F, inset). A conspicuousamount of secreted slime was found at the leading edge (Fig.1F). In addition, the leading edge contains a lot of the slime-containing vesicles (Fig. 1F, asterisks). Most of the vesicleswere localized beneath the plasma membrane of lobopods. Onthe plasma membrane at the ventral surface of lobopods, manyomega-shaped structures were seen (Fig. 1F, arrowheads),suggesting that the exocytosis of the slime-containing vesiclesoccurs primarily on the ventral side of lobopods.

Direct observation of exocytotic processes of theslime-containing vesicle in live plasmodiaTo identify the vesicle that contains the slime in live cells, weobserved the plasmodia with video microscopy and then fixedthem for electron microscopy (Fig. 2). Referring to thepositions of contractile vacuoles (Fig. 2D, asterisks), we iden-tified the slime-containing vesicle in ultrathin sections (Fig.2D), which had corresponding vesicular morphologies in thevideoimages (Fig. 2B). As shown in Fig. 2C, these vesiclesmoved at about 0.2 µm/second in the cytoplasm, faster thancontractile vacuoles that were not significantly translocatedduring the same observation period. Consistent with electronmicroscope observation, the diameters of the slime-containingvesicles were 3-4 µm in the videoimages. No other organelleswith such a large diameter were found in the cytoplasm exceptfor the nuclei and the contractile vacuole. Although the size of

Fig. 3. Shrinkage of a slime-containing vesicle. (A) Time-lapse images of the slime-containing vesicle during itsdisappearance. The vesicles in agar-overlaid plasmodiawere recorded with video-enhanced microscopy.Recording starts at the upper left image and proceeds at1/6-second intervals to the lower right image.(B) Representative time course of the changes in diameterof the slime-containing vesicle. Inset shows the diameterchanges of a contractile vacuole during representativecontraction. (C) Average rate of the changes in diameterof the slime-containing vesicle and the contractilevacuole. Error bars indicate s.d. (n=7). Bar in A, 10 µm.

contractile vacuoles was similar to that of the slime-containingvesicle, they could be readily distinguished due to their char-acteristic ultrastructures such as irregular contour of theirmembrane surface and the absence of visible contents, and totheir unique behavior repeating cycles of contraction andswelling (see details in the next paragraph).

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

813Pseudopod formation coupled with exocytosis

Fig. 5. Protrusion contains actin microfilaments. Agar-overlaidplasmodia were observed with video-enhanced microscopy: (A) 0,(B) 4, and (C) 11 seconds. The same plasmodia were fixed at 18seconds (D) and stained with tetramethylrhodamine-conjugatedphalloidin (E). The arrowhead in A indicates the slime-containingvesicle before exocytosis. Bar, 10 µm.

They gradually shrank in diameter while keeping theroundness in shape, and finally became undetectable. Asshown in Fig. 3B and C, the diameter shortened linearly andthe shortening rate was 1.1±0.3 µm/second (s.d., n=7). Afterdisappearance, the vesicles were not found in the other focalplanes at the same spot. This indicates that vertical transloca-tion of the vesicles did not occur. On the other hand, theaverage rate of contraction of contractile vacuoles was 12±5.0µm/second (n=7), ten times as fast as the shrinkage rate of theslime-containing vesicle (Fig. 3C). The slime-containingvesicles never swelled up again after they disappeared once,whereas contractile vacuoles repeated contraction and swelling(Fig. 3B, inset). It is, therefore, probable that the disappear-ance of the slime-containing vesicle is accounted for by exo-cytosis. Consistent with this notation, disappearance ofsecretory vesicles during exocytosis has been reported in chro-maffin cells (Edwards et al., 1984; Terakawa et al., 1991), andsalivary grand acinar cells (Segawa et al., 1991). Interestingly,the slime-containing vesicles occasionally fused with eachother in the cytoplasm before exocytosis. The fused vesiclesbecame larger in diameter than unfused vesicles (data notshown). Fusion between secretory vesicles has been reportedpreviously in mast cells (Lawson et al., 1977).

Cell surface protrusion after exocytosis of the slime-containing vesicleWhen the slime-containing vesicles disappeared at the celledge, a part of the neighboring cell surface protruded (Fig. 4).We observed a total of 246 cases of exocytosis, and found thatthe onset of such protrusion was within 5 seconds of vesicledisappearance in 134 cases. The protruded region showedlamella-like morphology, and did not appear, in most cases, tocontain intracellular granular organelles visible by video-enhanced DIC optics. The rate of protrusion was ~1µm/second, which is quite similar to the rate of pseudopodformation in plasmodia previously reported by Ueda andKobatake (1978).

Since one of the most important characteristics of thepseudopod is the presence of actin microfilaments (Condeelis,1993), we examined microfilament distribution in newlyformed protrusions. Plasmodia were fixed immediately afterprotrusion following exocytosis, as shown in Fig. 5A-D. Flu-

Fig. 4. Surface protrusion after exocytosis of the slime-containingvesicle in agar-overlaid plasmodia. The sequence starts at the upperleft image and proceeds at 0.6-second intervals to the lower rightimage. The arrowhead indicates the slime-containing vesicle, and thearrow indicates the onset of protrusion. Bar, 10 µm.

orescent phalloidin staining demonstrates that the cytoplasmicprotrusion contains actin microfilaments (Fig. 5E). Theintensity was relatively weak in this protrusion compared toneighboring regions, probably reflecting the difference inthickness between the protrusion and the remaining cell body.

To characterize the ultrastructural basis of the protrusion, wefixed cells for electron microscopy immediately after protru-sion had occurred (Fig. 6). In the protruded region (Fig. 6G),most of the membrane organelles were excluded, suggestingthat their main constituent is actin microfilaments, takentogether with phalloidin staining as shown above. The protru-sions displayed ultrastructures that showed continuous texturefrom the cortex to the protrusion. The cortical region containsthe actin microfilament meshwork (Naib-Majani et al., 1983),although OsO4 contained in the fixative we used destroyed finedetails of microfilament structures, as reported previously(Small, 1981). The presence of actin filaments in protrusionsat the cell edge was confirmed by the use of another fixativewith different constituents as follows: 2% glutaraldehyde,0.2% Triton X-100, 2 mM MgCl2, 0.2% tannic acid, and 60mM Na-cacodylate at pH 7.3 followed by post-fixation with1% OsO4 and 40 mM Na-cacodylate at pH 6.0. Plasmodia werenot agar-overlaid. Actin microfilaments were excellentlypreserved and their organization was clearly demonstrated tobe meshwork filling the entire space of the protrusions (Fig. 7).By contrast, in the region between the cortex and the granularcytosol, a distinct pattern of ultrastructure running parallel tothe cell edge was seen (Fig. 6G, arrows) and it probably rep-resents microfilament bundles (Ishigami et al., 1981). Theseobservations indicate that the protrusion contains an actinmicrofilament meshwork without conspicuous membraneorganelles.

Among the ultrathin sections of the newly protruded portionof the cell, the section close to the substrate contained theproximal part to the distal part of the protrusion (Fig. 6H).Judging from the overall morphology, the distal portion of theprotrusion with the round shape in Fig. 6H appears to corre-spond to the protrusion in Fig. 6G. This means that theproximal part of this protrusion is hidden by the overlayingcytoplasm in Fig. 6G. Since the vesicle in live plasmodia dis-appeared 4.6 µm from the cell edge, and since this distanceroughly corresponded to the length of the proximal part of theprotrusion, it is suggested that the protrusion started where thevesicle disappeared. Interestingly, the protrusion appears topush the secreted slime outward, judging from the stretchedappearance of the slime around the tip of the protrusion.

The slime-containing vesicles invaded the hyaline layer (Fig.

814 H. Sesaki and S. Ogihara

Fig. 6. Ultrastructuralcharacterization of theprotrusion. Agar-overlaidplasmodia were observed withvideo microscopy (A-E) andthen fixed for electronmicroscopy (F-H). (A-D) Theslime-containing vesicle(arrow in A) was exocytosedand surface protrusion (arrowsin B-E) followed. Note thehyaline appearance of theprotrusion. (F-H) Ultrathinsections of the sameplasmodia sectioned parallelto the substrate. Fixation wasdone about 5 seconds after E.Three sections are chosen sothat the one in F is furthest tothe substrate, the one in H isthe closest, and the one in G isbetween. The cell edgemarked with a bar in Froughly corresponds to such aregion in G and the protrusionin H. Black asterisks in E andF indicate unidentifiedvesicular structures observedboth in the videoimage andelectron micrograph. Thewhite asterisk indicates theslime-containing vesicle. Inthe region between the cortexand the granular cytosol,distinct pattern ofultrastructure which probablyrepresents microfilamentbundles (Ishigami et al., 1981)runs parallel to the cell edge(arrows in G). Bars: 5 µm(E,F,G); 2 µm (H).

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

Quantitative relationships between exocytosis andprotrusionTo determine spatial and temporal relationships between thedisappearance of the slime-containing vesicle and the protru-sion of the cell surface, quantitative analyses were performed.Fig. 9A shows a representative time course of the vesicle dis-appearance and surface protrusion. Protrusion started immedi-

ately 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 timelags ranging from 0 to 5 seconds after disappearance of theslime-containing vesicle (Fig. 9B).

If the membrane of a newly formed protrusion was suppliedby fusion of the slime-containing vesicle with the plasmamembrane, the surface area of the protrusion would be corre-lated to the surface area of the vesicle. To test for this possi-bility, the surface area of individual vesicles was calculatedfrom their diameter assuming that their shapes are true spheresin the cytoplasm. Surface area of protrusions was determinedby doubling the protruded area. Fig. 9C shows a virtually linearrelationship between the surface area of the protrusion and thatof the slime-containing vesicle. Furthermore, the ratio of

815Pseudopod formation coupled with exocytosis

Fig. 7. Organization of actin microfilaments in the protrusion. Spreadplasmodia without an agar overlay were fixed for electronmicroscopy. See Materials and Methods for details in the fixative. Arandom meshwork pattern of actin microfilaments is well preserved.Bar, 0.5 µm.

Fig. 8. Invasion of the slime-containing vesicle into thehyaline layer before exocytosis.(A-E) Videoimages showing thatthe slime-containing vesicle(arrows) invades the hyalinelayer on the cell edge of agar-overlaid plasmodia. The vesicledisappeared in the hyaline layer,and protrusion (arrowheads) wasobserved at the site ofexocytosis. Images were takenduring a period of 81 seconds.(F) Electron micrograph showingthe presence of the slime-containing vesicle (asterisks)near the cortical region justbeneath the plasma membrane.The one on the right has invadedthe cortical regions, the one onthe left is found in thegranuloplasm, and the oneinbetween lies in theintermediate zone. Agar-overlaid plasmodia locomoting on the agar subsparallel to the substrate. Note that vesicular organelles other than the slim10 µm (E); 2 µm (F).

protruded area to vesicle surface area showed values rangingfrom 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 aprotruded surface area became. Collectively all these datasupport the idea that membrane materials of the slime-con-taining vesicle were used as the plasma membrane of protru-sions during exocytosis.

DISCUSSION

We visualized single exocytotic events, and correlated them tothe process of cell surface protrusion. Exocytosis was observedwith video-enhanced microscopy combined with agar-overlaymethods applied to live plasmodia (Figs 3, 4, 5, 6 and 8). Theexocytotic vesicle contains and secretes slime, as revealed byelectron microscopy (Fig. 1). Slime is the extracellular matrixthat consists of D-galactan with an average degree of poly-merization of 560 (Henney, 1982). Preceding exocytosis, theslime-containing vesicle selectively invaded the hyaline layerwhich 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-containingvesicles, which enables the vesicles to access to the plasmamembrane before fusion. Cell surface in the neighboring regionprotruded at the site where exocytosis occurred (Figs 4, 5, 6and 8). The protrusion rate was quite similar to the rate ofpseudopod extension previously reported (Ueda and Kobatake,1978). Protruded regions then formed a new hyaline layer.They contained actin microfilaments (Fig. 5). The organizationof microfilaments was suggested to be meshwork of randomlyoriented filaments rather than straight bundles (Figs 6 and 7).These results, taken together, suggest that Physarum plasmodiaextend individual pseudopods after single exocytotic events ofthe slime-containing vesicle. Such protrusion, coupled with

trate were fixed, processed for electron microscopy, and sectionede-containing vesicles are densely confined in the granuloplasm. Bars:

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Fig. 9. Quantitative analysis of the relationship between exocytosisand protrusion. (A) Representative timecourse of the slime-containing vesicle shrinkage and protrusion. Filled circles indicatediameter of the vesicle. Open circles indicate length of theprotrusion. (B) Time lag between exocytosis and protrusion isvirtually proportional to the position of the slime vesicle from thecell edge. The time lags were determined using videoimages with0.25 second intervals. (C) Surface area of protrusions is proportionalto that of the slime-containing vesicle. (D) Ratio of surface areasbetween protrusion and vesicle depends on the position of vesiclesfrom the cell edge.

exocytosis of the slime-containing vesicle, may be the minimalprocess which underlies the pseudopod formation, and anassembly of many protrusions could lead to formation of alarge pseudopod called a lobopod (Taylor and Condeelis, 1979)observed in Physarum plasmodia.

While surface protrusion that is coupled with exocytosis ofthe slime-containing vesicle showed the characteristics ofpseudopods as discussed above, it is not clear whether or notthese protrusions are directly involved in locomotion of theplasmodium. For example, it has been shown that not allpseudopods that extend from the cell surface lead to cellmigration (Condeelis, 1993). Also a rearward transport processof the surface structure (Abercrombie et al., 1970; Heath andHolifield, 1991), if present in the Physarum plasmodium,would considerably reduce the contribution of the surface pro-trusion to the net migration of the plasmodium. There is alsoa possibility that the protrusion functions merely in the cellspreading process, not tightly associated with the net migrationwhich obviously requires the coupling with retraction of theposterior part of the plasmodium.

The shortening rate of the diameter of the slime-containingvesicle during exocytosis was ten times as slow as the con-traction rate of the contractile vacuole (Fig. 3D), although theirdiameter was quite similar. Such a rate difference between the

exocytotic vesicle and the contractile vacuole revealed in thisarticle strongly supports a proposal for different mechanismsunderlying these two types of membrane deformation. Con-traction of the contractile vacuoles could be driven by an ATP-dependent active process involving actin microfilaments andmyosin I, as reported recently (Doberstein et al., 1993). On theother hand, in exocytosis, resumption of the planar nature ofthe plasma membrane after fusion of vesicles and formation ofthe omega shape as an intermediate state of the exocytosiscould be a passive process probably due to the intrinsic natureof the membrane including surface tension and/or to positiveinner pressure of the cytoplasm (Kamiya, 1959). Alternatively,the slow rate of the slime-containing vesicle shrinkage couldbe explained by the viscosity of the contents to be extruded.Slime is a branched filamentous structure as shown in theelectron micrograph in Fig. 1 (Wolf et al., 1981). Biochemicalcharacterization (Henney, 1982) confirms its branchedstructure and hence its viscous nature. Consistent with thisnotion, the slow rate of exocytosis in general has beenexplained by a difference in the diffusibility of the vesiclecontents in acinar cells and goblet cells which secrete theviscous mucus (Segawa et al., 1991).

A series of experiments has raised the hypothesis that exo-cytosis is required for pseudopod formation in locomoting cells(Bretscher, 1983, 1984; Bergmann et al., 1983; Bretscher andThomson, 1983; Singer and Kupfer, 1986; Kupfer et al., 1987).This hypothesis would predict some quantitative relationshipsbetween exocytosis and cell surface extension, if we assumethat insertion of the new membrane materials via exocytosisdirectly allows the cell surface to expand. One of the criticalquestions to be addressed has been whether or not the amountof membrane materials supplied by exocytosis can explain theexpansion of the plasma membrane at the site of thepseudopod. But this question has not been so far directlyexamined, because the size of the exocytotic vesicle isgenerally too small to be correlated to pseudopod formation.Here we directly correlated single exocytosis to pseudopodformation in a quantitative manner, taking advantage of the factthat the slime-containing vesicles are extremely large andhence visible with light microscopy. Also the proximity of theexocytosis site and the protrusion site is an advantage of theplasmodium. Protruded areas were substantially proportionalto the surface area of the vesicle (Fig. 9C) and did not exceedthe membrane area that was supplied by exocytosis (Fig. 9D).Calculation of the surface areas of the slime vesicle and theprotrusion based on their morphologies could be exact enoughto obtain accurate values, firstly because the slime-containingvesicle is almost a complete sphere with a smooth membranecontour, and secondly because the protrusion has negligiblethickness and a smooth membrane contour. These resultsstrongly indicate that the slime-containing vesicle supplied thepre-existing plasma membrane with new membrane materialsto expand its surface during pseudopod formation.

It is of interest to understand the geometrical relationshipbetween exocytosis and protrusion. Do protrusions really startat the site of exocytosis? To answer this question, we tried tocorrelate the site of the exocytosis to that of the protrusion.Some protrusions started obviously at the region where theslime-containing vesicle disappeared, i.e. in which exocytosisoccurred at the very edge of the plasmodia. In the other casesin which the slime-containing vesicles disappeared at sites rel-

817Pseudopod formation coupled with exocytosis

atively distant from the cell edge, not all the sites of protrusionwere distinct. Since protrusion at the cell edge occurred imme-diately after the exocytosis (Fig. 9A), the observed time lagsin exocytosis distant from the cell edge allowed us to estimatehow 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), andcan be converted to distance in a range from 0 to 5 µm, takingthe protrusion rate to be 1 µm/second (Fig. 9A). These valuesmatched well with the distance between where the slime con-taining vesicle disappeared and where protrusions came outfrom the cell edge. In addition to these quantitative analyses,electron microscopy suggests that these time lags could beexplained by three-dimensional topography of the cell edgewhich renders protrusions invisible in a top view using the lightmicroscope until the protrusion comes out from the edge aftera certain delay. In fact, the proximal part of the protrusion ishidden by overlying cytoplasm, when exocytosis occurredaway from the cell edge (Fig. 6). Collectively, it is highlyplausible that protrusion started at the site where exocytosisoccurred.

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

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(Received 28 May 1996 - Accepted 27 January 1997)