comparative freeze-fracture stud of y perialgal and digestive vacuole in … · 2005. 8. 22. ·...

J. Cell Sd. 71, 121-140 (1984) 121Primed in Great Britain © The Company of Biologists Limited 1984

COMPARATIVE FREEZE-FRACTURE STUDY OFPERIALGAL AND DIGESTIVE VACUOLES INPARAMECIUM BURSARIA

RENATE MEIER'*, MARCELLE LEFORT-TRAN2, MONIQUEPOUPHILE2, WERNER REISSER3 AND WOLFGANG WIESSNER1

1 Abteilung fur Experimentelle Phykologie, Institut fur Pflanzenphysiologie, UniversitatGottingen, Untere Karspule 2, D-3400 Gottingen, Federal Republic of Germany2Laboratoire de Cytophysiologie de la Photosynthese, CJJJi.S., F-91190 Gif-sur-Yvette,France3 Fachbereich Biologie, Universitat Marburg, Lahnberge, D-3550 Marburg, FederalRepublic of Germany

SUMMARY

In the endosymbiotic unit of Paramecium bursaria (Ciliata) and Chlorella sp. (Chlorophyceae)algae are enclosed individually in perialgal vacuoles, which do not show acid phosphatase activityand thus differ from digestive vacuoles. Both types of vacuoles have been studied by freeze-fracture.

Perialgal vacuoles are nearly spherical; their membrane always fits tightly to the algal surface.The vacuole size and shape do not vary much. During division of the algal cell into four autosporesthe vacuole diameter only doubles. After autospore formation the vacuole invaginates around thealgal daughter cells and divides. Newly formed perialgal vacuoles remain in intimate contact andexhibit characteristic attachment zones before final separation. The two fracture faces of perialgalvacuole membranes are homogeneously covered with intramembranous particles (IMPs) but rarelyshow signs of vesicles pinching off or fusing with the membrane, except during vacuole division.The P-faces bear more IMPs (3164 ± 625 IMP//mi2) than the E-faces (654 ± 208 IMP//im2). Therange of IMP density on both faces is enormous, suggesting that the membrane is not static.Membrane changes are supposed to occur simultaneously with the enlargement of the vacuole andto be caused by fusion with cytoplasmic vesicles, as the fractured necks on vacuole membranes mayindicate.

Digestive vacuoles in P. bursaria show significant variations in size, shape, membrane topographyand IMP density, as well as signs of endocytic activity. Different vacuole populations are presentin P. bursaria according to different feeding conditions: ciliates fed for a long time have smallvacuoles with few IMPs (322 ± 198 IMP//m2 on the E-faces, 1438 ± 458 IMP//im2on the P-faces),which are probably condensed digestive vacuoles, whereas organisms fed for a short time have largervacuoles with highly paniculate faces (680 ± 282 IMP//*m2 on the E-faces, 2701 ± 503 IMP//im2

on the P-faces) and thus are supposed to be older vacuoles. The digestive vacuole membrane changescontinuously.

Compared to digestive vacuoles perialgal vacuoles are characterized by small size combined withhigh IMP density on the two fracture faces. Their IMP densities resemble those of old digestivevacuole membranes. However, it would be premature to conclude that membranes of perialgal andold digestive vacuoles are identical. Membranes of old digestive vacuoles are mainly derived fromlysosomal material, which presumably does not contribute to the formation of perialgal vacuolemembranes as is indicated by the small vacuole diameter; fusion with lysosomes would considerablyenlarge it. The conclusions from these results are discussed in relation to the formation of perialgalvacuoles during the establishment of a stable symbiotic unit after infection of alga-free P. bursariawith the Chlorella species exsymbiotic from green paramecia.

•Author for correspondence.

122 R. Meter and others

INTRODUCTION

Paramecium bursaria contains two types of vacuoles. Digestive vacuoles areformed at the cytopharynx when food particles are engulfed and then pass through aseries of morphological and physiological changes. During this cyclosis food particlesare digested and the remaining material is defecated at the cytoproct, as was reportedfor other Paramecium species (Allen, 1978; EsteVe", 1970; Fok, Lee & Allen, 1982;Jurand, 1961; Mast, 1947; Muller & Toro, 1962; Schneider, 1964). The secondvacuole population consists of several hundred perialgal vacuoles, which harboursymbiotic algae (Chlorella sp., Chlorophyceae) and do not occur in other Parameciumspecies. Thin-section electron-microscopical studies, as well as cytochemical charac-terization of these vacuoles (Karakashian & Karakashian, 1973; Karakashian,Karakashian & Rudzinska, 1968; Meier, Reisser & Wiessner, 1980; Reisser, 19766;Vivier, Petitprez & Chive1, 1967), revealed distinct differences between perialgal anddigestive vacuoles: unlike the latter, perialgal vacuoles: (1) are mainly located in theperipheral cytoplasm of the Paramecium; (2) they always contain only one algal cell,since chlorellae and the surrounding vacuoles divide synchronously; and (3) they lackacid phosphatase activity so that the enclosed symbionts are not digested. The reasonsfor these differences between the two vacuole types are unknown.

In the natural habitat perialgal vacuoles are always present in P. bursaria. Theciliate and its chlorellae form a stable symbiotic unit ('green Paramecium') withmutual benefit to each partner (Karakashian, 1963; Pado, 1965, 1967; Pringsheim,1928; Reisser, 19766; Weis, 1974). Only under laboratory conditions is it possible toseparate ciliates and algae from each other and to culture them independently (Prings-heim, 1928; Reisser, 1975, 1976a; Wichterman, 1943). Alga-free ciliates can be easilyreinfected with the Chlorella species exsymbiotic from P. bursaria. During suchinfection experiments algae are phagocytosed and many of them pass through cyclosisand are digested. But some algae succeed in re-establishing the symbiotic associationby an unknown mechanism, resulting in the formation of perialgal vacuoles (Bomford,1965; Hirshon, 1969; Karakashian & Karakashian, 1965; Reisser, Meier & Wiessner,1980; Siegel, 1960; Weis, 1978). The mechanism by which perialgal vacuoles areformed in green paramecia is also not completely understood.

The present freeze-fracture study describes the morphological characteristics ofperialgal and digestive vacuoles of P. bursaria and compares the freeze-fractureparameters of their membranes in order to test whether the differences between thetwo vacuole types are based on a different organization of their membranes and to gaininsight into the origin and formation of perialgal vacuoles.

MATERIALS AND METHODS

Culture conditionsGreen and alga-free Paramecium bursaria Ehrbg. were cultured in bacterized lettuce medium at

22 °C and a light/dark regime of 14: 10 h (Philips fluorescent tubes TL4OW-1/32 and TL40W-1/55; illuminance, 14-9 Wra"z).

Vacuoles in Paramecium bursaria 123

Primary treatment of paramecia

Perialgal vacuoles were investigated in starved green paramecia that were free of digestivevacuolea as well as in organisms that had been fed with bacteria for 4h before cryofixation.

Digestive vacuoles were studied in alga-free P. bursaria fed with bacteria for 4 h and in ciliatesthat had been provided with food for 12 min only, thereafter washed twice in sterile lettuce mediumand then transferred for 30 min into sterile medium before freezing.

Rapid freezing and freeze-fracturing

For cryofixation cells were harvested by gentle centrifugation, sandwiched between two copperfoils and frozen at — 180cC without prior fixation and glycerol treatment using a double liquidpropane-jet device (Balzers Cryo-Jet QFD 101). Samples were fractured at — ISO °C without etchingand replicated using a Balzers double-replica device in a Balzers BA 360 M freeze-etch unit equippedwith electron guns and a quartz crystal monitor. Cleaned replicas were mounted on uncoated coppergrids and examined with a Hitachi HU 11A electron microscope at 75 kV or a Philips EM400 at80 kV.

Calculations of intramembranous particle (IMP) density and vacuole size

The IMP density was measured on selected micrographs of membrane faces with minimal tiltenlarged to x 100 000. On each micrograph IMPs were counted in ten squares of 0-04/im2 each andthe average IMP density on each membrane face was calculated for 1-0fjxn1. Data were obtainedfrom 30 E and 30 P-faces of different perialgal and digestive vacuoles, respectively, and are givenas mean IMP density ± the standard error of the mean (S.E.M.).

The vacuole diameter was determined from micrographs of those vacuoles that had been used forIMP density measurements.

RESULTS

Membrane fracture faces of perialgal and digestive vacuoles

During freeze-fracturing membranes are split internally (Branton, 1966), thusexposing two fracture faces, the protoplasmic face (P-face) and the complementaryexoplasmic face (E-face). According to the definition of Branton et al. (1975) the P-face of vacuole membrane3 shows concave, and the E-face convex, curvature.

Perialgal vacuoles can be easily identified in freeze-fracture replicas of green P.bursaria. They appear nearly spherical and parts of the endosymbiotic chlorellae areexposed when the fracture plane has passed through the vacuole lumen (Figs 1,2).Most strikingly, this perialgal vacuole lumen seems to be less extensive than wasformerly assumed on the base of thin-section electron micrographs (Karakashian etal. 1968). Our freeze-fracture pictures clearly show that the vacuole membrane andthe algal cell wall are closely associated, leaving a distance of 0-05 [im at most betweenthem. Since the material had not been pretreated chemically for the freeze-fracturetechnique, the formerly observed larger distance between membrane and algal surfacemust be regarded as an artifact resulting from fixation and embedding procedures.

Sometimes two, three or four perialgal vacuoles are found to be continuous witheach other or to be linked together (Figs 3—6). Although the dynamics of vacuolemovement are not visible in freeze-fracture replicas, it can be assumed that thesemicrographs document division stages of perialgal vacuoles, because the symbioticChlorella is known to divide into two or four autospores, each of which is enclosed

R. Meier and others

Figs 1-2


again in an individual perialgal vacuole (Karakashian et al. 1968; Vivieret al. 1967),whereas simultaneous fusion of four perialgal vacuoles is improbable. Using the freeze-fracture technique it was possible to gain further insight into the mode of vacuoledivision. The perialgal vacuole starts to invaginate around the algal daughter cells aftertheir release from the mother cell. Thereby the close contact of the vacuole membraneto the algal surface is always maintained (Fig. 3B). Remnants of the algal mother cellwall can be seen in the lumen of the dividing perialgal vacuole (Fig. 3B) . During divisionthe daughter vacuoles are arranged linearly (Fig. 3A) or circularly (Fig. 4A) . Frequentlydaughter vacuoles are linked together, revealing characteristic circular attachmentzones between them (Figs 4-6). The continuation of the fracture faces in the centre ofthe attachment zone of neighbouring daughter vacuoles, as well as the alternation of Eand P-faces in that zone (see Fig. 4B) , indicates that two distinct vacuole membranes arepresent, i.e. that division of the vacuole is already complete. However, the membranesof neighbouring daughter vacuoles remain in intimate contact before final separation.They exhibit corresponding depressions and projections, which confine the zone ofattachment (Figs 4-6). The diameter of this zone varies, possibly indicating differentstages of separation of the new perialgal vacuoles. Thus, the vacuoles seen in Fig. 4Bhave a large attachment zone and therefore seem to be less separated from one anotherthan those in Figs 5 and 6. Stages of this final separation of daughter vacuoles can befound more frequently than stages of the initial invagination of a vacuole (Fig. 3).Probably the final separation needs more time than the early steps of vacuole division.

The vacuole topography is generally smooth and there is almost no endocytic activityassociated with the membrane (Figs 1,2). Only a few small circular depressions on theE-faces, similar projections on the P-faces and fractured necks might indicate someinitial contact of vesicles, fusion processes or membrane blebbing (Figs 4,5). They arefound more frequently on membranes of dividing perialgal vacuoles.

Abbreviations used in figures: Ep^ and Ppm,, E and P-face of the perialgal vacuole mem-brane ; Epm and Ppm , E and P-face of the algal plasma membrane; Ea, and POT , E and P-faceof the outer algal chloroplast envelope membrane; Ed and Pa , E and P-face of the inneralgal chloroplast envelope membrane; Ejv and Pdv , E and P-face of the digestive vacuolemembrane; cw, cell wall of the endosymbiotic Chlorella sp.; mctu, mother cell wall of thealgal cell; dv, digestive vacuole; b, bacterium; v, vesicle. Bare, 1 /im.

Figs 1, 2. Perialgal vacuole and enclosed endosymbiotic Chlorella cell.Fig. 1. A large portion of the convex E-face of the perialgal vacuole membrane is

exposed. This fracture face is homogeneously covered with IMPs but reveals no furtherstructures, e.g. signs of endocytic activity (see also Fig. 2). Under the cross-fractured cellwall of the enclosed alga a part of the particle-rich P-face of its plasma membrane is visible.The two algal chloroplast envelope membranes are fractured in a characteristic pattern,exposing the E-face of the outer chloroplast envelope membrane with several 'windows'through which the P-face of the inner chloroplast envelope membrane can be seen. Notethe very small distance between the algal surface and the perialgal vacuole membrane.X27 600.

Fig. 2. The fracture plane has exposed the successive concave membrane faces: theparticle-rich P-face of the perialgal vacuole membrane, the E-face of the algal plasmamembrane, which is poor in particles, the P-face of the outer chloroplast envelope mem-brane and detached portions of the E-face of the inner envelope membrane. The Chlorellacell wall is seen in cross-fracture close to the perialgal vacuole membrane. X32 600.

126 R. Meier and others

The digestive vacuole system in Paramedum caudatum was thoroughly studied byAllen (1976) and Allen & Staehelin (1981). As these authors pointed out, shape andmembrane topography of digestive vacuoles change during cyclosis. Accordingly, a

Fig. 3. Division stage of a perialgal vacuole. A. After division of the Chlorella cell into fourautospores the perialgal vacuole starts to invaginate around them (arrows), the developingdaughter vacuoles are still continuous with each other. X11 600. B. Enlargement of the twodaughter vacuoles at the left of A. The fracture plane has passed through the vacuole lumenexposing parts of two algal daughter cells and remnants of the algal mother cell wall. Theclose contact of the vacuole membrane to the algal surface is maintained even duringdivision of the perialgal vacuole. X45 000.

Vacuoles in Paramecium bursaria II?

Fig. 4. Division stage of a perialgal vacuole. A. The four daughter vacuoles (1—4) arecircularly arranged (contrary to the linear arrangement seen in Fig. 3A). TWO vesicles areattached to the vacuole (arrows). Several circular depressions on the membrane face(arrowheads) arise from the connection of vesicles to the vacuole membrane, probablybefore fusion. X 17 000. B. Enlarged view of the lower left of A showing the circularattachment zone of two neighbouring daughter vacuoles (arrowheads) characterized bycorresponding depressions and projections. Since the fracture plane has passed from theP-face of one to the E-face of the adjacent vacuole, which is continuous also in the centreof the attachment zone, it can be assumed that division of the perialgal vacuole has beenfinished already and that the individual daughter vacuoles are linked together before theirfinal separation. X45 000.


pm pav

-pav

Figs 5,6. Attachment zone of neighbouring daughter vacuoles after division of the perialgalvacuole.

Fig. 5. The much smaller diameter of the attachment zone compared to that in Fig. 4Bmight indicate an advanced stage of vacuole separation. Circular depressions on the E-faceindicate contact of vesicles with the vacuole membrane. X45 000.

Fig. 6. Detached patches of the P-face of the vacuole below (arrows) are attached to theE-face of the vacuole above, thus indicating intimate contact between the two perialgalvacuoles. X45 000.

variety of digestive vacuoles is also observed in P. bursaria, ranging from smallspherical vacuoles, which are very condensed and model the enclosed bacteria (Figs7, 8), to large and irregularly shaped ones. The vacuole membranes are either smooth,i.e. without signs of endocytic activity (Figs 7, 8) or they reveal signs of membraneblebbingor fusion with vesicles (Figs 9—11). The vesicles seen in Fig. 9, surroundingthe vacuole and fusing with it, might be primary lysosomes. Similar structures havebeen observed by Allen & Staehelin (1981). On the other hand, the vesicles seen incontinuity with a cross-fractured digestive vacuole (Fig. 10) might result from

Figs 7, 8. Small digestive vacuole. The vacuole membrane closely surrounds the ingestedand tightly packed bacteria so that their imprints are visible on the fracture faces (arrows).The vacuole membrane does not show signs of endocytic activity.

Fig. 7. E-face with few particles. X35 700.Fig. 8. Particle-rich P-face and portion of the condensed vacuole contents. X 38 500.


1/

dv

Figs7-8


Figs 9-11


membrane blebbing towards the cytosol, which is reported by Allen & Staehelin(1981) to occur in old digestive vacuoles. The partial digestion of the enclosed bac-teria, indicating the advanced age of this vacuole, supports this interpretation.Numerous fractured necks on the E-face of a vacuole (Fig. 11) are also signs of theendocytic activity of that vacuole. We conclude from the observed variations invacuole morphology that digestive vacuoles of different age are circulating in P.bursaria under the given feeding conditions.

Size ofperialgal and digestive vacuoles

Vacuoles are characterized not only by shape but by size. Although measurementsof vacuole diameter must be interpreted with caution because of the impossibility ofdetermining whether a vacuole is fractured through its centre, the data are useful forcomparative studies.

The diameters of perialgal vacuoles (Table 1) vary only a little. Thus the smallestvacuole that was measured in replicas of green paramecia had a diameter of 2*5 £im,while the largest perialgal vacuole was almost twice this diameter. For division intofour daughter vacuoles the surface area must enlarge fourfold while the vacuolediameter only doubles.

The minimum value of the diameter of digestive vacuoles (Table 1) is of the sameorder of magnitude as that of perialgal vacuoles, but contrary to the latter digestivevacuoles enlarge up to sixfold and are almost twice as large in diameter as a dividing

Table 1

Vacuole type

. Diameter of perialgal

Mean*

and digestive vacuoles in

Diameter (^m)

Smallest

P. bursaria

Largest

PerialgalDigestive^

(a) 4h(b) 12min

3-6±0-5

5-1 ±1-05-7±3-0

2-5

2-12-5

4-8

7-212-9

• Mean values are given ± S.E.M. ; for determination see Materials and Methods,f Digestive vacuoles from paramecia frozen after 4h of feeding (a) or after 12min of feeding

followed by a 30-min transfer into sterile medium (b).

Fig. 9. Digestive vacuole surrounded by vesicles, probably lysosomes. One vesicle isattached to the vacuole membrane (arrow), which also shows a fractured neck (arrowhead)indicating contact with another vesicle. X27 000.

Fig. 10. Cross-fractured digestive vacuole. Many small vesicles lie next to the vacuole orare continuous with it, either fusing with the vacuole or pinching off from it (arrows). Thepartial digestion of the vacuole contents (arrowhead) hints at the advanced age of thevacuole. X27000.

Fig. 11. Condensed digestive vacuole modelling the enclosed bacteria (see also Figs 7, 8).A. Numerous fractured necks are scattered on the E-face. x27 000. B. Enlargement of thecentral part of the vacuole seen in A. The fractured necks (arrows) point into the cytosolindicating membrane blebbing of the vacuole or fusion with cytoplasmic vesicles.X 60 000.


perialgal vacuole. The great variation in size between different digestive vacuolesaccounts for the large standard errors of the mean. Paramecia that have been fed withbacteria for 4 h have considerably smaller vacuoles than ciliates that have been fed for12 min and transferred afterwards into sterile medium for 30 min. It can be assumedthat, depending on feeding conditions, digestive vacuoles of different size are presentin P. bursaria.

Intramembranous particle (IMP) density on perialgal and digestive vacuole mem-brane fracture faces

The membrane fracture faces are covered with IMPs, which represent internalmembrane proteins. Their number and distribution characterize each membrane andits functional differentiations (Satir & Satir, 1974).

Both E and P-faces of perialgal vacuole membranes are homogeneously coveredwith IMPs. On the E-face their number ranges from 390 to 1353//im2, on the P-facefrom 1883 to 4300//im2. It is evident that there is an enormous variation in IMPnumbers on both faces, which also accounts for the large standard errors of the meanIMP density (Table 2). Nevertheless, these calculations can be used for comparativestudies. The great diversity of IMP density on different vacuole membranes alsoindicates that perialgal vacuole membranes are not static. The P-face of perialgalvacuole membranes bears significantly more particles than the E-face (Table 2). Thisunequal distribution of IMPs between the complementary faces is typical for mostbiological membranes and is also expressed by the partition coefficient (Kp, Table 2),which gives the ratio of IMPs on P and E-faces (Satir & Satir, 1974).

In order to determine the distribution of IMP densities on E and P-faces of theperialgal vacuole population, the number of faces observed with different IMPdensities was plotted (Fig. 12). The distribution on E-faces shows a significant peak,which corresponds closely to the mean IMP density on E-faces (Table 2). On theother hand, there is no prevailing IMP density on P-faces of the perialgal vacuolepopulation of P. bursaria.

Fracture faces of digestive vacuole membranes are also homogeneously coveredwith IMPs. The P-face always bears more IMPs than the E-face; the partition co-efficient is similar to that of perialgal vacuole membranes (Table 2).

Table 2. IMP density and partition coefficient of perialgal and digestive vacuolemembranes in P. bursaria

Vacuole type

PerialgalDigestivef

(a) 4h(b) 12 min

f For explanation

Mean

rE-face

654 ± 208

322 ±198680 ±282

see legend to Table

IMP

1.

density (IMP//W)*A

P-face

3164 ±625

1438 ±4582701± 503

Total

3818

17603381

Partitioncoefficient

Kp

4-8

4-54-0


(Att)U

avc

40-

30

20

c

I 10

cn

vE

cID

CJID

40-

30

10-

-0Lru

0 1000IMP density (IMP//mi2)

1000 2000 3000 4000

IMP density (IMP/^m2)

1000 2000 30001000IMP density (IMP//mi2)

Fig. 12. Distribution of IMP densities on E and P-faces of perialgal (A) and digestive (B)vacuole membranes in P. bursaria. The IMP densities are grouped into classes of200 IMP//imz. Percentages of membrane faces in each IMP density class are plotted foreach vacuole type and fracture face. Paramecia with digestive vacuoles were frozen after4 h of feeding (black) and after 12 min of feeding followed by a 30-min transfer into sterilemedium (white).


Depending on feeding conditions, digestive vacuoles have different ranges of IMPdensities on both E and P-faces (Fig. 12). The vacuole E-faces of ciliates that havebeen fed for 4h have 145-1000 IMP/jan2, while their P-faces have 720-2765 IMP//im2. Digestive vacuole E-faces of paramecia that have been fed for 12min and trans-ferred afterwards into sterile medium for 30 min have IMP numbers ranging from 388to 1230/jUm2, while complementary P-faces have IMP densities between 2138 and3738 IMP//im2. Obviously, the IMP density of digestive vacuole membranes ofparamecia fed for short times is nearly twice as high as that of vacuoles of organismsfed for 4 h. The frequency of E and P-faces plotted against the IMP density (Fig. 12)shows clear peaks. We therefore assume that two nearly homogeneous but differentvacuole populations are present in the paramecia under the given feeding conditions.

DISCUSSION

In endosymbiosis research the question of the special character of perialgal vacuolesis of basic interest. Perialgal vacuoles with enclosed symbiotic chlorellae are presentnot only in green Paramedum bursaria but also in other endosymbiotic associationssuch as are formed by Climacostomum virens, Euplotes daidaleos, Stentor poly-morphus, Spongilla lacustris, Vorticella sp., Hydra viridis and Anthopleuraxanthogrammica (Reisser & Wiessner, 1984). According to ultrastructural andcytochemical investigations the perialgal vacuoles of, at least, green Climacostomum(Reisser, Meier & Kurmeier, 1983), Hydra (Hohman, McNeil & Muscatine, 1982;O'Brien, 1982) and Anthopleura (O'Brien, 1980) display features similar to those ofP. bursaria, i.e. they always enclose a single Chlorella cell only and do not show acidphosphatase activity, thus differing from digestive vacuoles. But the vacuole mem-branes have never been characterized, although they may account for the observedfunctional differences between both vacuole types. Therefore, our comparative studyof perialgal and digestive vacuole membranes in/3, bursaria provides further evidenceof differences between the two vacuole types and may be of general importance alsofor other symbiotic associations.

For this investigation we have chosen the freeze-fracture technique because by itlarge areas of individual membranes are fractured, exhibiting their IMP populationfor a detailed study of membrane ultrastructure and enabling dynamic and localizedactivities, such as fusion events, to be seen. The best preservation of membranearchitecture and dynamic processes was achieved by the preparation of cells with anadvanced rapid-freezing technique (Moor, Kistler & Miiller, 1976; Miiller, Meister& Moor, 1980). This method avoids artifacts caused by glutaraldehyde fixation andglycerol treatment of the material during conventional freezing. It thereforeapproaches the conditions in vivo and guarantees a high 'spatial and time resolution'of fixation (Plattner & Bachmann, 1982).

Preceding a comparison of perialgal with digestive vacuoles and in order to deter-mine specific features of perialgal vacuoles, digestive vacuoles will be characterizedfirst. Our freeze-fracture study shows that digestive vacuoles in P. bursaria differ insize, shape, membrane topography and IMP density. Previous investigations on


Tetrahymena pyriformis (Batz & Wunderlich, 1976; Kitajima & Thompson, 1977),Acanthamoeba castellanii (Bowers, 1980) and P. caudatum (Allen, 1976; Allen &Staehelin, 1981) also demonstrated that digestive vacuole membranes are not staticbut undergo several changes according to vacuole age. In their timed study of vacuoledifferentiation in P. caudatum, Allen & Staehelin (1981) described three stages ofdigestive vacuoles, named DV-I, DV-II and DV-III, differing in age and freeze-fracture parameters. DV-I are reported to be large young (not more than 6min old)vacuoles with 4575 ± 733 IMP//im2 on their E-face and 1142± 365 IMP/^m2 ontheir P-face. DV-II are 3—25 min old, condensed vacuoles with 60 ± 44 IMP//wn2 ontheir E-face and 1402 ± 203 IMP/jUm2 on their P-face. DV-III are expanding andmature vacuoles, more than 10 min old and reveal a characteristic IMP pattern of1144 ± 338 IMP/Vm2 on the E-face and 3367 ± 508 IMP//mi2 on the P-face.

In contrast to different membrane types investigated so far (Dempsey, Bullivant &Watkins, 1973; Lefort-Tran, Pouphile, Freyssinet&Pineau, 1980; Lefort-Tran e< a/.1978) the IMP densities on both digestive and perialgal vacuole membranes in P.bursaria and their partition coefficients are not altered by fixation and glycerinationprocedures. This is evident from a comparison of the data presented here with thoseobtained from conventionally frozen P. bursaria (Meier, Reisser, Wiessner & Lefort-Tran, 1980). Therefore, it is admissible to compare our results with those of Allen &Staehelin (1981) obtained from conventionally frozen/3, caudatum. This comparisonshows that none of the digestive vacuoles found in P. bursaria is young enough tocorrespond to a DV-I stage, which is clearly characterized by its particle-rich E-faceand reverse membrane polarity. But the range of IMP density, from 145 to 1230 IMP/^mz on the E-faces and from 720 to 3738 IMP/^m2 on the P-faces of digestive vacuolemembranes in P. bursaria overlaps that reported for DV-II and DV-III. Moreover,assuming that 50% of DV-II and 50% of DV-III are present in P. caudatum, theresulting mean particle density can be calculated to be 602 IMP//zm2 on the E-faceand 2385 IMP/jtim2 on the P-face, giving a total of 2987 IMP//im2 and a partitioncoefficient of 4-0. These hypothetical values correspond closely to the IMP densitiesreported by us for digestive vacuole membranes in P. bursaria that have been fed for12 min. We therefore suggest that in these organisms vacuoles resembling DV-II andDV-III are present in equal numbers. On the other hand, ciliates that have been fedfor 4h have more vacuoles with fewer IMPs on their fracture faces. Fewer IMPs arecharacteristic for DV-II. These paramecia also have smaller vacuoles than ciliates fedfor the shorter time. This observation also hints at a greater number of vacuolesresembling DV-II, since Allen & Staehelin (1981) reported DV-II to have a diameterof only 3-5 /xm whereas the diameter of DV-III is up to 16*4/im. The occurrence oflarge vacuoles (up to 12-9 jxm) in P. bursaria that have been fed for 12 min thusconfirms the suggestion that those paramecia contain a considerable number of oldvacuoles resembling DV-III.

Our experiments reveal that P. bursaria not only has distinct vacuole stages suchas DV-I, -II and -III but that transition stages also exist (see Fig. 12). Digestivevacuoles apparently change continuously in size and IMP density when they passthrough cyclosis, even though some stages seem to occur more frequently or to exist


for a longer time. To simplify the identification of digestive vacuoles in the replicas,we used alga-free P. bursaria throughout our experiments, thus avoiding confusionwith perialgal vacuoles. But control investigations of digestive vacuoles of greenorganisms (data not shown) confirm the results obtained from alga-free ciliates.Obviously, symbiont-containing ciliates do not differ from alga-free P. bursaria as faras the digestive vacuole system is concerned. From the comparison of our freeze-fracture data of digestive vacuoles in P. bursaria with those of P. caudatum and fromthe observed similarities between both species we conclude that formation of digestivevacuoles in P. bursaria and the following events during cyclosis are very similar tothose described for P. caudatum by Allen and co-workers (Allen, 1974, 1976; Allen& Fok, 1980, 1983a,6; Allen & Staehelin, 1981; Allen & Wolf, 1974).

Unlike digestive vacuoles, perialgal vacuoles in P. bursaria do not show dramaticvariations in size, shape and membrane topography. Their membrane always fitstightly to the algal surface. This fact may be of great importance for the exchange ofnutrients between both partners (Reisser, 1981) and for the ciliate-alga recognitionphenomena (Reisser, Radunz & Wiessner, 1982). It also implies that changes in sizeof perialgal vacuoles depend directly on growth of the enclosed Chlorella cell andexplains the slight variations in size of perialgal vacuoles. Under given culture con-ditions, ciliates divide once a day. If one algal cell divides into four autospores, onlyone third of the chlorellae have to divide for maintenance of a constant number ofintracellular algae. Since perialgal vacuoles must enlarge simultaneously with growthand division of the algae, this low division rate of the endosymbiotic Chlorellaaccounts for the small changes in size of perialgal vacuoles, which are less dramaticthan the extensive changes in digestive vacuoles occurring within even a few minutes.

The lack of acid phosphatase activity (Karakashian & Karakashian, 1973) andespecially the absence of thorotrast label (Karakashian & Rudzinska, 1981) in perial-gal vacuoles have been taken as a sign of the inhibition of fusion between lysosomesand perialgal vacuoles. Our measurements of vacuole diameter support this assump-tion, since fusion with primary lysosomes should result in an enormous enlargementof the vacuole as has been reported by Allen & Staehelin (1981) for DV-II in P.caudatum: the diameter of these vacuoles enlarges 4-7-fold after fusion withlysosomes and the concomitant change to DV-III. Corresponding changes in sizehave never been observed in perialgal vacuoles of green Paramecium.

The two fracture faces of perialgal vacuole membranes show an enormous range ofIMP density, most likely because the internal membrane organization is not static.These IMP density changes may depend on growth and division of the enclosedsymbiotic alga. Fractured necks, depressions and projections on perialgal vacuolemembranes, appearing especially during vacuole division, indicate that vesicles maybe involved in the process of division. Such vesicles may contribute new membranematerial to a perialgal vacuole, thus giving rise to its enlargement and simultaneousalteration of IMP density. As discussed above, these vesicles must differ fromlysosomes. In addition they may be formed by new membrane material, since egestionof symbiotic algae and concomitant recycling of perialgal vacuole membrane materialwas not observed by us.


A comparison between the ranges of IMP density on perialgal and digestive vacuolemembranes (see Fig. 12) shows that the ranges on E and P-faces of perialgal vacuolemembranes overlap those on corresponding membrane faces of those digestivevacuoles that are supposed to resemble DV-III after fusion with primary lysosomes.We therefore conclude that the membranes of perialgal and old digestive vacuoles (DV-III) are similar. Moreover, IMP size distributions, obtained from preliminarymeasurements of the diameter of IMPs on E and P-faces of perialgal and digestivevacuole membranes (unpublished results), do not differ significantly. However, itwould be premature to conclude that membranes of perialgal and old digestive vacuolesare identical, since a predominant part of the latter is lysosome-derived (Allen &Staehelin, 1981), whereas we suppose that lysosomes do not contribute membranematerial to perialgal vacuoles. Hence our freeze-fracture results show that perialgalvacuoles are characterized by the combination of a high IMP density on the two frac-ture faces and a small diameter and in this regard differ from digestive vacuoles.

The freeze-fracture characterization of perialgal and digestive vacuoles in P.bursaria gives new insight into events during infection of alga-free P. bursaria withchlorellae suitable for symbiosis formation. During infection algae are taken up byphagocytosis but then escape digestion by triggering the formation of perialgalvacuoles, which may be formed either at the cytopharynx or after enclosure of chlorellaeinto digestive vacuoles. Since the IMP density on perialgal vacuole membrane faces andthe membrane polarity differ fundamentally from those reported for the cytopharynxmembrane (Allen & Staehelin, 1981), the first hypothesis implies the existence of aspecial type of vesicle, which fuses with the cytopharynx membrane in the course ofalga—ciliate recognition events and thus encloses algae by a new, i.e. perialgal, type ofmembrane. According to the second hypothesis, suitable algae enter the digestive cycleat first, i.e. they are enclosed in digestive vacuoles, which is probably achieved by fusionof discoidal vesicles with the cytopharynx membrane as has been reported by Allen &Staehelin (1981) for formation of digestive vacuoles in P. caudatum. Perialgal vacuoleformation is then assumed to occur at either of two different steps of cyclosis, i.e. beforefusion of the young digestive vacuole (D V-I) with acidosomes, a type of vesicles knownto cause the acidification of digestive vacuoles inP. caudatum (Allen & Fok, 1983c), orbefore fusion of the condensed DV-II with lysosomes, since perialgal vacuoles do notshow acid phosphatase activity (Karakashian & Karakashian, 1973) and are small indiameter. Since the IMP density on perialgal vacuole membrane faces does not resem-ble the density on D V-I or DV-II membrane faces, the membrane must change duringperialgal vacuole formation. If fusion only with lysosomes is prevented, the pH also,reported to be 3*0 in DV-II (Fok et al. 1982), should rise again, since symbioticchlorellae survive in a medium with a pH lower than 4-5 only for a short time (Reisser,1975). Both membrane transformation and increase in pH might be achieved by fusionof special, possibly symbiosis-specific vesicles, or by selective removal of membranematerial during the process of perialgal vacuole formation.

We thank Roland Boyer for excellent photographic assistance. This research was supported bya grant from the Stiftung Volkswagenwerk (Az 35/392) to Professor Dr W. Wiessner.


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(Received 27 April 1984-Accepted 14 May 1984)

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