THE JOURNAL OF BIOLOGICAL CHEMISTRY Q 1993 by The Amancan Society for Biochemistry and Molecular Biology, Inc. Vol. 288, No. 9, Issue of March 26, pp. 6742-674’7.1993

Printed in U. S. A.

A Post-lysosomal Compartment in Dictyostelium discoideurn”

(Received for publication, September 22,1992)

Harish Padh, Juhyun Ha, Malti Lavasa, and Theodore L. SteckS From the Department of Biochemistry and Mokclllcrr Bwbgy, University of Chicago, Chicago, Illinois 60637

Fluorescein isothiocyanate (F1TC)-dextran and pyr- anine were fed to the social amoeba, Dictyosteliurn discoideum. These membrane-impermeable, pH-sensi- tive fluorophores initially entered a =neutral endocytic compartment. They encountered maximal acidity (pH = 6) about 16 min after ingestion, in what appeared to be digestive vacuoles (lysosomes). The environment of the probes returned to near neutrality by 30 min. At that time, the probes accumulated in a decreasing num- ber of vacuoles of increasing size; ultimately, there were only a small number of vacuoles per cell with diameters of up to 3 pm. The late vacuoles sedimented more rapidly than did proton pumps, acid hydrolases, and recently ingested cargo. Unlike the vacuoles har- vested immediately after the cells were fed FITC-dex- tran, the late vacuoles were not acidified by MgATP in vitro. Egestion of ingested FITC-dextran com- menced after a lag of -46 min. A similar lag was observed for the resurfacing of two endocytosed bi- layer-intercalated fluorophores.

These results suggest that, in Dictyostelium, undi- gested endocytic cargo accumulates in and is returned to the cell surface through a distinctive compartment of large and nearly neutral post-lysosomal vacuoles. It will be important to determine the degree to which internalized plasma membrane components follow this post-lysosomal pathway.

In endocytosis, there must be a homeostatic balance be- tween the internalization and the return to the cell surface of both the ingested plasma membrane and the enclosed aqueous compartment. In many animal cells, such restoration proceeds rapidly at the pre-lysosomal level (1-7). It is generally thought that unassimilated ingested matter becomes sequestered in terminal residual bodies, where it may remain indefinitely (see Refs. 8 and 9). Presumably, post-lysosomal egestion is minimal in the cells of higher organisms, which are not burdened with the indigestible remains of ingested microbes. Nevertheless, various animal cells expel undigested soluble markers from large, late compartments with time constants of several hours (1, 2, 4, 5, 7). The exocytic path from lysosomes back to the cell surface is not well characterized in animal cells.

Pre-lysosomal processing of receptor-bound ligands is not prominent in protozoa. Rather, ingested microbes and

BE-21 and National Science Foundation Grant MCB-9113366. The * This research was supported by American Cancer Society Grant

costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked “aduertisenent” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

$ To whom correspondence should be addressed Dept. of Biochem- istry and Molecular Biology, University of Chicago, 920 E. 58th St., Chicago, IL 60637. Tel.: 312-702-1329; Fax: 312-702-0439.

aqueous solutes are promptly conveyed to lysosomes; indiges- tible residues are continually egested from a post-lysosomal station (10, 11). The cell surface sites for endocytosis and exocytosis and the intervening digestive pathway are anatom- ically well defined and controlled in ciliates (12) and flagel- lates (13, 14).

In amoebae, like many animal cells, surface sites for endo- cytosis and exocytosis are usually not fixed. Early studies ascribed egestion to large, late vacuoles lacking the acidic pH and acid hydrolases characteristic of lysosomes (10, 15, 16). The goal of the present study was to characterize the late vacuolar limb of the endocytic pathway in Dictyostelium dis- coideum.

EXPERIMENTAL PROCEDURES

Materials and Cells“. discoideum strain Ax-3 was cultured ax- enically as described (17). Pyranine,’ octadecylrhodamine B, and TMA-DPH were obtained from Molecular Probes; FITC-BSA, FITC- dextran (M, 70,000), and Triton WR-1339 (Tyloxapol) were from Sigma. All other reagents were prepared as reported (17-19) unless indicated otherwise.

Endocytosis-Cells were harvested in mid-exponential phase, washed, and used immediately (17). Prior to feeding probes, washed cells were allowed to recover at room temperature for 10-20 min. Incubations were conducted at room temperature with the cells swirl- ing continuously at 200 rpm. Centrifugation of cells in time courses was in a microcentrifuge.

Fluorescence-Fluorescence was quantitated in a Perkin-Elmer LS-50 spectrophotofluorimeter. Total fluorescence was determined in 5 mM sodium glycinate (pH 8.5) containing 0.2% Triton X-100. Excitation and emission wavelengths for FITC were 495 and 520 nm; for rhodamine B, 540 and 570 nm; and for pyranine, 403 and 510 nm.

The pH within endocytic vacuoles containing FITC-dextran was read from a reference curve; this was constructed using cells bearing the ingested probe and made permeable to protons with 0.1 mM oleic acid suspended in a buffer of 50 mM MES adjusted to the indicated pH with acetic acid or NaOH. Cellular fluorescence was determined using emission at 520 nm and excitation at both 450 and 495 nm. The ratio of the fluorescence intensity of the former to the latter reading was plotted as a function of pH (20, 21). We obtained comparable calibration curves with cells loaded with FITC-dextran and fixed with formaldehyde (22).

To calibrate the pH in vacuoles bearing ingested pyranine, the cells were washed into 50 mM histidine adjusted to several pH values with acetic acid or Tris base, and oleic acid was added to 0.1 mM. Pyranine fluorescence was determined with excitation at 403 and 453 nm; emission was at 510 nm. The ratio of the first fluorescence value to the second reflected pH (Fig. 4A).

The pH dependence of the fluorescence of endocytic FITC-dextran and pyranine was not significantly different from that of the probes in free solution. Furthermore, the spectral properties of these probes were not altered by 1 h of internalization. In all experiments where data were expressed as relative fluorescence or fluorescence ratios, the absolute readings were robust (i.e. comparable to that shown in Fig. lA).

The abbreviations used are: pyranine, 8-hydroxypyrene-1,3,6- trisulfonic acid; TMA-DPH, trimethylammonium-diphenylhexa- triene; FITC, fluorescein 5-isothiocyanate; BSA, bovine serum albu- min; MES, 4-morpholineethanesulfonic acid.

6742

A Post-lysosomal Compartment in D. discoideum 6743

Analytical Procedures-The activities of acid hydrolases and the vacuolar H'-ATPase were determined as described (19).

RESULTS

Kinetics of Uptake and Release of FITC-Dextran-Vegeta- tive amoebae took up pinocytic probes with a nearly linear time course for a period of over 1 h in the experiment shown in Fig. 1A and up to 2 h in other experiments (see Fig. 2 and Refs. 23 and 24). Release of FITC-dextran following the ingestion of a 5-min pulse commenced after a lag of approxi- mately 45 min in the experiment shown (Fig. lB, curve 1 ) and no less than 30 min in several similar experiments. The egestion of 80% of the fluophore was accompanied by the release of less than 3% of lysosomal N-acetylglucosaminidase (Fig. lB, curve 2), acid phosphatase (Fig. 1B, curve 3), and 8- galactosidase activities (not shown).

Kinetics of Digestion of Endocytic FITC-BSA-Cells took up this probe with a nearly linear time course for more than 2 h (Fig. 2A, curve 1 ) . The digestion of the probe to acid- soluble fragments commenced after a lag of no more than 15 min (Fig. 2A, curve 2). (The quantitation of fragment accu- mulation here was imprecise because secretion of fragments into the extracellular space was not evaluated. The fact that the pool of intracellular acid-soluble fragments reached an early plateau presumably signifies such an exocytic process.)

I I

0 w o

w 0 60 120 MINUTES

FIG. 1. Endocytosis and exocytosis in Dictyoetelium. Panel A , endocytosis. Cells were washed, resuspended in growth medium to 5 X 10' ml-', and swirled to allow recovery. FITC-dextran was added to 2 mg ml-'. Aliquots of the swirling suspension were taken period- ically; the cells were washed three times with ice-cold 5 mM glycine- NaOH (pH 8.5) containing 100 mM sucrose and resuspended to 3.3 X 10' ml-' in this buffer containing 0.2% Triton X-100. Fluorescence was recorded in arbitrary units with excitation at 495 nm and emission at 520 nm. Panel B, exocytosis. Cells were fed FITC-dextran for 5 min as in p a n e l A, washed thrice, resuspended in growth medium, and swirled. Aliquots were centrifuged at intervals and the fractional release of markers determined. Curve 1 (open circles), FITC fluores- cence, determined as in panel A. Curve 2 (triangles), N-acetyl-,% glucosaminidase activity. Curve 3 (closed circles), acid phosphatase activity.

w 0 Z

0 v) w CY 0.4 0 3 J

0.8

Ir. 0.0

0 ..-., rc 0 2 4 0.0 0.5 1.0 1.5

HOURS FIG. 2. Digestion of ingested FITC-BSA. Panel A, continuous

endocytosis. Cells were prepared as in Fig. L4. FITC-BSA was added to 2 mg ml" and the suspension swirled. Aliquots were taken period- ically; the cells were washed with ice-cold buffer and dissolved in buffer containing 0.4% Triton X-100. Curve 1, total cellular fluores- cence determined as in Fig. lA. Curve 2, fluorescence of cell-associated probe soluble in 5% trichloroacetic acid. To facilitate precipitation, BSA was added to 0.8 mg ml-'; the acidity was then offset by reading small aliquots in 300 mM NaH2P0,, pH 8.0. Panel B, pulse-chase mode. Cells were fed FITC-BSA for 10 min as in p a n e l A, washed, resuspended in 22 mM KPi (pH 6.5) containing 2 mM MgCIz, and swirled as in Fig. 1B. At intervals, uncentrifuged aliquots were treated with trichloroacetic acid and the acid-soluble fluorescence deter- mined. Ordinate values were normalized to unity.

As seen in pulse-chase experiments (Fig. 2B), acid-soluble fragments had already accumulated following 10 min of feed- ing and <lo min of washing. In view of the possibility that acid hydrolases are dispersed along the pre-lysosomal endo- cytic pathway (25), this experiment did not distinguish be- tween endosomal and lysosomal digestion.

Time Course of Acidification of Ingested FITC-Dextran- Cells were fed a 10-min pulse of FITC-dextran, washed, and their fluorescence followed as an indicator of pH in the tagged vacuoles (Fig. 3A). The apparent pH value of this endocytic space was initially close to neutral, fell to approximately 5 in 15 min, and then rose sharply to a plateau value of pH >6.5, where it remained after 30 min. Similar measurements were made during continuous feeding (Fig. 3B). Maximum acidity was observed at -20 min as in Fig. 3A. That the fall and rise of vacuolar pH in Fig. 3B was less acute than in Fig. 3A is attributed simply to the lack of synchrony among the vacuoles in the cells continuously fed.

Control experiments showed that this pH reversal was not dependent on: (a ) the endogenous fluorescence of the cells, ( b ) release of the probe to the extracellular fluid, (c) the concentration of FITC-dextran, (d) the ligand bearing the FITC, (e) the cleavage of FITC from its carrier, or (f) its leakage from the vacuoles. Comparable pH values were ob- tained when rhodamine-dextran was employed as an internal reference in the dual fluorescence ratio mode (26). Finally, the fluorescence time course was nullified by treating the cells with the ionophore, nigericin (Fig. 3, curve 2 in panels A and B).

Time Course of Acidification of Ingested Pyranine-Pyran- ine is another good indicator of vacuolar pH (Fig. 4 4 ; see also Ref. 27). It gave results comparable to FITC-dextran with respect to the time course of its uptake as well as its acidifi- cation and deacidification in the pulse mode (Fig. 4B, curve 2) and when fed continuously (Fig. 4C, curve I). The pyranine fluorescence time course was also sensitive to nigericin (solid circles in Fig. 4, B and C ) . Pyranine was always entirely vacuolar (see Fig. 5).

Znhibitwn of Endocytic Activity Arrests the Progress of the

6744 A Post-lysosomal Compartment in D. discoideun

I Q

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5.3 0

A

L 0 30 60 90

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0.8 ; 0 m z 0

0.2 m

al P "1

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MINUTES FIG. 3. Time course of acidification of FITC-dextran in-

gested by Dictyoetefium. Panel A, pulse-chase mode. Cells were fed FITC-dextran for 10 min, washed, resuspended in growth medium at 5 X lo' ml-', and swirled, all as in Fig. 1B. At intervals, aliquots were centrifuged, the cells resuspended to 6.7 X lo6 ml" in 50 mM MES-NaOH (pH 7.0) containing 10 mM KCl, and cellular fluores- cence determined for curue 1 (see "Experimental Procedures"). Ni- gericin was then added to 10 p~ and fluorescence redetermined for curue 2. Plotted against time is the ratio of the fluorescence values at 520 nm excited at 450 and 495 nm (right axis) and the apparent vacuolar pH read from a calibration curve (Zejt axis). Panel B, Con- tinuous mode. Washed cells were fed FITC-dextran. At the indicated times, aliquots were washed and resuspended to 6.7 X 10' ml" in 50 mM MES-NaOH (pH 7.0) containing 10 mM KCl. Cellular fluores- cence was determined before (curve 1 ) and after nigericin addition (curue 2), as described above. In both panels, the quantity of fluores- cence detected was similar to that shown in Fig. L4.

Acidification Time Course-Incubation on ice inhibits endo- cytosis and exocytosis in Dictyostelium. In the experiment shown in Fig. 4B, cells loaded with a pulse of pyranine and chased at 22 'C were not only read directly (curue 2) but duplicate samples were also taken at each time point and held on ice for 1 h prior to fluorimetry (Fig. 4B, curve 1 ). Clearly, arresting endocytic flow on ice immediately arrested the pro- gression of the pH in the environment of the endocytic probe. The same result was obtained with FITC-dextran.

We have performed experiments such as those in Fig. 3 and 4 both on the amoeba, Acanthamoeba castellani, and the ciliate, Tetrahymena pyriformis. The time courses for acidifi- cation and deacidification were similar to those for Dictyos- telium.

Pyranine Is a Cytological Indicator of the pH in the Endo- cytic Pathway-Cells loaded with pyranine for 30 min and viewed through a fluorescein filter set showed a few large vacuoles, 1 to 3 pm in diameter (Fig. 5A). When such cells were permeabilized to buffer at pH 5, the pyranine fluores- cence was abolished (panel B). In cells permeabilized at pH 7 (panel C) and at pH 9 (panel D), a multitude of vacuoles of small to large size was observed. A violet fluorescence filter set, which visualizes pyranine at all pH values (27), gave images similar to those at pH 9 for unpermeabilized cells. The

5.21 ' 0 20 40 60

MINUTES FIG. 4. Time course of acidification of pyranine ingested by

Dictyostelium. Panel A, calibration curve. Cells fed pyranine for 30 min were washed and resuspended in buffers of different pH values containing 0.1 mM oleic acid. The fluorescence was determined at excitation of 403 and 453 nm and emission of 510 nm, and the fluorescence ratio plotted on a log scale as a function of pH (see "Experimental Procedures"). Panel B, pulse-chase mode. Washed cells were fed 5 mM pyranine for 10 min as in panel A . The loaded cells were then washed and resuspended in 22 mM potassium phos- phate buffer (pH 6.5) containing 2 mM MgC12 and chased as in Fig. 1B. At intervals, duplicate samples were collected. One set was placed on ice for 1 h, and the other was immediately centrifuged. In trace 2, the pelleted cells were resuspended in 50 mM MES-NaOH (pH 7.0) containing 10 mM KC1 and their fluorescence recorded. The right axis gives the fluorescence ratio; the left axis gives the apparent pH read frompanel A. The fluorescence ratios were redetermined on each sample after addition of 10 pM nigericin (trace 3 ) . For trace 1, the aliquots held on ice for one hour were centrifuged and processed as in trace 2. Panel C, continuous feeding. Cells were prepared and fed as in Fig. 3B except that the probe was 5 mM pyranine. For trace I , aliquots were washed at intervals and the fluorescence ratio deter- mined as in panel B . For trace 2, 10 p~ nigericin was added to each sample in trace I and the fluorescence ratio redetermined.

probe was, in all cases, entirely vacuolar and not cytosolic (Fig. 5).

Within a few minutes of ingestion, vacuolar pyranine be- came increasingly visible in the fluorescence microscope. The first vacuoles to appear were of moderate sue. Apparently, the smallest endocytic vacuoles did not become deacidified before delivering their pyranine to larger ones.

Acidification in Vitro Is Specific for Early Endocytic Vacu- oles-Cells were fed FITC-dextran for 10 min, washed, chased for 0, 20, and 60 min, and homogenized in the presence of M$+, which preserves their capacity for vacuole reacidifica- tion. The native acidity in the organelles was discharged and the re-acidification of the FITC-dextran space followed in vitro (19). Re-acidification of the FITC-dextran compartment was strong in cells that had not been chased (Fig. 6, curue 3 ) .

A Post-lysosomal Compa

FIG. 5. Visualization of endocytic vacuoles containing pyr- anine. Cells were fed 5 mM pyranine for 30 min and washed as described in Fig. 4. The cells were resuspended in 50 mM histidine- acetate at pH 5.0 or 50 mM histidine-Tris base at pH 7.0 or 9.0. 0.1 mM oleic acid was added to the samples for panels B-D. Cells were photographed through fluorescein filters as described (17). Panel A, pH 7.0 without oleic acid. Panel B, pH 5.0 plus 0.1 mM oleic acid. Panel c, pH 7.0 plus 0.1 mM oleic acid. Panel D, pH 9.0 plus 0.1 mM oleic acid. Bar = 10 pm.

w u z

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I Y Y 1

1 I O 2 4

MINUTES FIG. 6. Acidification of endocytic FITC-dextran in vitro.

Cells loaded with FITC-dextran for 10 min as in Fig. 1 were washed and further incubated in growth medium for 0, 20, or 60 min. The cells were then pelleted and homogenized in 5 mM sodium glycinate (pH 8.5) plus 100 mM sucrose containing 2 mM M&l2 (17, 18). The unfractionated homogenate was centrifuged and the pelleted organ- elles washed, purged with nigericin, and analyzed for ATP-dependent (starting at 1 min) and nigericin-reversible (starting a t 4 min) quench- ing of FITC-dcxtran fluorescence as described (19). Curues 1-3 show the fluorescence time course in homogenates prepared after 60, 20, and 0 min of chase, respectively. Because a small amount of the probe was egested during the chase, the maximal fluorescence in each trace was normalized to 100; traces 1 and 2 were shifted upward for visibility.

However, the capacity for re-acidification was greatly reduced by a 20-min chase (curve 2) and was minimal after a 1-h chase (curve I).

Sedimentation Velocities of Acidosomes, Lysosomes, and Early and Late Endocytic Compartments-Cells were fed rho- damine B-dextran for 100 min, washed, and then fed FITC- dextran for 20 min. The labeled cells were homogenized, and four markers were analyzed for their clearance by differential centrifugation (Fig. 7). The most sedimentable marker was

.rtment in D. discoideum 6745

* I i k > I- o U W > t- U

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-

- W

1 o3 1 o4 g-mln

1 os 10'

FIG. 7. Differential centrifugation of homogenates contain- ing short term and long term endocytic cargo. Cells were fed rhodamine B dextran (M, 70,000) a t 2 mg ml" for 100 min and washed as in Fig. 1. The cells were then fed FITC-dextran at 2 mg ml" for an additional 20 min. The cells were washed and homoge- nized; the homogenates were diluted in 5 mM glycine-NaOH (pH 8.5) containing 100 mM sucrose to 1 X 10' cell equivalents m1-I. The suspension was centrifuged in a Sorvall SS-34 rotor. The supernatant fraction was centrifuged again at a higher time-integrated force. This process was repeated serially and the successive pellets analyzed. Curue 1 (solid triangles), rhodamine B dextran. Curve 2 (solid circles), acid phosphatase activity. Curue 3 (open triangles), FITC-dextran. Curve 4 (open circles), vacuolar H+-ATPase activity. Control experi- ments demonstrated no significant energy transfer between the fluo- phores.

the first-fed cargo, rhodamine B-dextran (curve 1 ). The least sedimentable marker was the vacuolar H+-ATPase (curve 4 ) , which in Dictyostelium is almost entirely associated with acidosomes, a separate proton-pumping organelle (19). Acid phosphatase (curue 2) and the recently ingested cargo (FITC- dextran; curve 3) had identical, intermediate sedimentation velocities. Two other acid hydrolases, @-galactosidase and N- acetylglucosaminidase, invariably coincided with acid phos- phatase activity. These data concur with cytochemical evi- dence that acid hydrolases in Dictyostelium are located in small, not large, vacuoles (28).

Endocytosis and Exocytosis of the Plasma Membrane- TMA-DPH is a cationic fluorescent dye that partitions into the outer leaflet of the plasma membrane bilayer without entering the cytoplasm (29, 30); it thus serves as a probe for tracing plasma membrane endocytosis and exocytosis (31). Intact Dictyostelium bound TMA-DPH within 1 min. When endocytosis was inhibited, the bound dye was entirely re- moved by washing cells with a non-disruptive lipoidal deter- gent, Triton WR-1339. Under standard conditions of endo- cytosis, TMA-DPH increasingly accumulated in vacuoles vis- ible in fluorescence microscopy. As it became internalized, the fraction of the probe removed by Triton WR-1339 (hence, that at the cell surface) declined. The half-time of such internalization was about 5 min at room temperature. Inter- nalization and re-externalization (see below) were both blocked at 0 "C and by the presence of 150 mM NaCI, two inhibitors of endocytic flow (not shown). At steady state, the amount of probe inside the cell was approximately 1.5 times greater than that extractable from the cell surface; this is the same fraction of plasma membrane internalization found using [3H]galactose conjugates (23).

The time course of resurfacing of TMA-DPH, which had been internalized for a period of 5 min, was followed as the extractability of cellular fluorescence (Fig. 8A). Initially, there was a rapid loss of -20% of the probe, conceivably due to exocytosis from a small, early endocytic compartment and/or reflux from contractile vacuoles, which apparently fuse inter-

6746 A Post-lysosomal Compartment in D. discoideum

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100

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FIG. 8. Return of endocytic TMA-DPH to the cell surface. Cells were washed and resuspended to 1 X 10' cells ml-I in 50 mM glucose, 20 mM KPi (pH 7.0), 2 mM MgClz containing 10 PM TMA- DPH. At this cell concentration, virtually all of the probe partitioned into the plasma membranes (not shown). The suspension was swirled for 5 min (panel A ) or 60 min (panel B ) to allow endocytosis, at which times aliquots were diluted into 10 volumes of the same buffer containing 2% Triton WR-1339, previously shown to remove the probe exposed at the extracellular surface. The suspensions were swirled and, at the indicated times, aliquots collected. These were washed once with ice-cold buffer containing 2% Triton WR 1339 and twice more with ice-cold buffer containing 150 mM NaCl and 1 mM NaN8 (previously shown to inhibit endocytosis and exocytosis). The cells were resuspended in the same buffer and cellular TMA-DPH fluorescence determined (excitation, 360 nm; emission, 430 nm). Fractional cellular fluorescence is plotted against chase time.

mittently with the plasma membrane. There followed a lag of about an hour, with little or no exocytosis. Resurfacing of the probe then resumed and exhibited roughly first-order kinetics over a few hours.

In contrast, cells that had taken up TMA-DPH for 60 min released the endocytic probe without a lag (Fig. 8B). Not only were these findings quite reproducible, but preincubation times between 5 and 60 min gave lag periods intermediate between those shown in panels A and B of Fig. 8. Our failure to remove all of the ingested probe is ascribed to the efficient re-ingestion of emergent probe molecules by ongoing endo- cytosis.

The lag seen in Fig. SA could reflect the transit time required for a wave of internalized plasma membrane mole- cules to traverse an endocytic pathway prior to its return by exocytosis to the cell surface. The 1-h preincubation in Fig. 8B presumably allowed the probe to spread through the entire endocytic pathway, thereby obviating the lag.

We have performed similar experiments using octadecyl- rhodamine B, a fluorescent probe of utility similar to TMA- DPH (32). Its exit from cells was similar to that shown in Fig. 8.

We cannot infer whether the path through the cell of TMA- DPH and octadecylrhodamine B is the same as that of in- gested aqueous solutes or plasma membrane proteins. It is clear, however, that the internalized bilayer-intercalated probes return to the cell surface only after a prolonged period in the cell interior.

DISCUSSION

These results suggest the following picture of the endocytic circuit in Dictyostelium, at least with respect to the probes employed. Ingested solutes enter a compartment of roughly

neutral pH. Acidification commences promptly (Figs. 3 and 41, apparently by transient associations of the endocytic com- partments with a separate proton-pumping organelle (Fig. 6; see Refs. 19,33, and 34). At this point, endocytic probes reside in numerous small and acidic vacuoles (Fig 5; see also Ref. 17). These are presumably digestive (Fig. 2). At approximately 20 min, probe molecules abruptly leave that acidic environ- ment (Figs. 3 and 4) for nearly neutral vacuoles, which de- crease in number and increase in size over time (Figs. 5 and 7). The differential in pH between the young and old vacuoles (Figs. 3 and 4) suggest that their contents do not mix rapidly. The reciprocal relationship of their size and number suggests that the large vacuoles incorporate the smaller ones. (If, however, young vacuoles merged with old ones, the endocytic probes would appear in the largest ones from the start, which we do not observe. Thus, young endocytic vacuoles may repeatedly merge with cohorts of the same age.) The largest vacuoles are low in both acid hydrolase activity (Fig. 7) and acidity (Fig. 5 and Ref. 17). On these grounds, they are taken to be post-lysosomal. Egestion of soluble endocytic probes begins 30-45 min after ingestion (Fig. 1B ), about the time at which the probes enter the large, nearly neutral vacuoles. That the late vacuoles are mostly dissipated by fusion with the plasma membrane is also suggested by the lag of >30 min in the resurfacing of the internalized cell surface markers TMA-DPH (Fig. 8) and octadecylrhodamine B.

Since some acid phosphatase activity is present in large cytoplasmic vacuoles in Dictyostelium (35), it may be that acid hydrolases are gradually retrieved from post-lysosomal vacuoles as they age. That the post-lysosomal vacuoles are normally entirely purged by the time they meet the plasma membrane is suggested by a very low rate of secretion of acid hydrolases (Fig. 1B). Secretion of mature acid hydrolases can be induced in Dictyostelium under special conditions (36); enzyme retrieval may thus be regulated, perhaps by recycling from post-lysosomal vacuoles to active lysosomes (12, 37). If so, the latter could be viewed as a sorting organelle potentially comparable to the well studied endomembrane systems in animal cells (38).

The reversal of the acidification of endocytic vacuoles in Dictyostelium, demonstrated here in Figs. 3, 4 and 7, was recently documented with membrane impermeable 31P probes and NMR (39). The mechanism underlying this interesting phenomenon is presently obscure. Because the acidic vacuoles are impressively proton-tight in vitro (19, 33), we expect that the discharge observed in situ is more than passive permea- bility. Proton symport of digestion products into the cyto- plasm could be involved (see Refs. 40-42). Another factor is the dissociation of endocytic vacuoles from proton pumps at the time they become deacidified (Fig. 6). On the other hand, lysosomal digestion should itself maintain vacuolar acidity through the deprotonation of weak acid hydrolysis products (e.g. amino acids, fatty acids, and phosphate esters). Given that the sharp fall in endocytic acidity seen in Figs. 3 and 4 reflects the average of a vast number of imperfectly synchro- nized vacuoles, the deacidification of any one of them may be an exceedingly brief, triggered event.

Residual bodies in animal cells might be the homologues of post-lysosomal vacuoles in protozoa; they too tend to be large in size, small in number, neutral in pH, of diminishing acid hydrolase activity, and capable of exocytic fusion with plasma membranes (8,9,46). However, endocytic markers in animal cells typically remain in acidic compartments for hours (20, 43). Macrophages are unusual in that their newly formed phagosomes are transiently alkalinized (44). In some cells, endocytosed transferrin leaves an early acidic compartment

A Post-lysosomal Compartment in D. discoideum 6141

for a neutral space within 5 min of uptake (45); however, this phenomenon seems to represent recycling of receptor-ligand complexes to the cell surface through a pre-lysosomal path- way.

The post-lysosomal system described here is common among protozoa rather than divergent in Dictyostelium. In Acanthumoeba, endocytic cargo is acidified and deacidified with a time course similar to that shown for Dictyostelium? In addition, aging phagosomes in Acanthumoeba are poor in acid phosphatase activity (35). The late vacuoles in Acan- thumoeba are few in number but, unlike Dictyostelium, not notably large in size (47).

Entamoeba histolytica similarly manifests prolonged filling of an extensive compartment of large vacuoles prior to the onset of exocytosis (48). These vacuoles are not acidic and could be distinguished from a more numerous population of small and acidic lysosomal vacuoles.

Endocytic processing in amoebae is comparable in many ways to that seen in ciliates despite the fact that, in Para- mecium and Tetrahymena, the pathway begins at an anatom- ically defined cytopharynx, ends with exocytosis at a cyto- proct, and creates endocytic compartments from specialized cytoplasmic membranes rather than from internalized plasma membrane (12). As in Dictyostelium, the nascent digestive vacuoles in Paramecium become rapidly acidified through association with accessory acidosomes; their pH drops to a minimum after about 5-10 min; they then acquire acid hydro- lases; the resulting digestive vacuoles lose their acidity by about 20 min and, by about 30 min, lose various acid hydrolase activities as well; at that time, post-lysosomal vacuoles begin to expel their wastes by exocytosis (12). The ciliate Tetrahy- mena shows a similar behavior (49, 50). Moreover, FITC- dextran and pyranine ingested by T. pyriformis report a time course of pH change comparable to that for Dictyostelium.'

In conclusion, the similar functional plans for endocytosis in amoebae and ciliates may reflect an archetypal eukaryotic pattern. Post-lysosomal sorting may have preceded pre-lyso- soma1 sorting in evolution, and the post-lysosomal pathway in protozoa could share mechanisms mediating organelle traffic with the endomembrane systems in animal cells.

Acknowledgments-We are grateful to Alan A. Finegold and Su Zheng for their help in initiating this project.

REFERENCES 1. Besterman, J. M., Airhart, J. A,, Woodworth, R. C., and Low, R. B. (1981)

2. Adams, C. J., Maurey, K. M., and Storrie, B. (1982) J. Cell Biol. 93,632- J. Cell Biol. 9 1 , 716-727

GRII 3. &%man, R. M., Mellman, I. S., Muller, W. A., and Cohn, Z. A. (1983) J.

4. Daukas, G., Lauffenburger, D. A., and Zigmond, S. (1983) J. Cell Bbl. 96, Cell BioL 96 , l -27

1642-1650

H. Pa&, J. Ha, M. Lavasa, and T. L. Steck, unpublished obser- vations.

5. Swanson, J. A., Yirinc, B. D., and Silverstein, S. C. (1985) J. Cell Biol.

6. Thilo, L. (1985) Biochim. Biophys. Acta 822,243-266 7. Buktenica, S., Olenick, S. J., Salgia, R., and Frankfater, A. (1987) J. Biol.

Chem. 262,9469-9476 8. Daems, W. T., W~sse, E., and Brederoo, P. (1969) in Lysosomes in Biology

and Pathology (Dingle, J. T., and Fell, H. B., eds) pp. 64-112, North- Holland Publishing Company, Amsterdam

100,851-859

9. Holtzman, E. (1989) Lysosomes, Plenum Press, New York

11. Stockem, W. (1977) in Mammalian Cell Membranes (Jamieson, G. A., and 10. Kitching, J. A. (1956) Protoplnsmatologia 3.D.3b, 1-54

12. Fok, A. K., and Allen, R. D. (1990) Int. Reu. Cytol. 123,61-94 13, Vickerman, K., and Preston, T. M. (1976) in Bwlogy of the Kznetoplastidn

(Lumsden, W. H. R., and Evans D. A., eds) Vol. 1, pp. 35-130, Academic

Robinson, D. M., e&) Vol. 5, pp. 151-195, Butterworth, Boston

Press, New York 14. Webster, P. (1989) Eur. J . Cell Biol. 49,295-302 15. Chapman-Andresen, C. (1973) in The Biology of Amoeba (Jeon, J. W., ed)

pp. 319-348, Academic Press, New York 16. Ryter, A., and de Chastellier, C. (1977) J. Cell Biol. 76,200-217 17. Padh. H.. Lavasa. M.. and Steck. T. L. (1989) J. Cell Biol. 108,865-874 18. Padh; H.; Lavasa; M.,'and Steck,'T. L. (1989)'Biochem. Biophys; Acta 9 8 2 ,

271-278 19. Padh, H., Lavasa, M., and Steck, T. L. (1991) J. Biol. Chem. 2 6 6 , 5514-

5520 20. Ohkuma, S., and Poole, B. (1978) Proc. Natl. Acad. Sci. U. S. A. 76,3327-

3331 21. Ohkuma, S. (1987) in Lysosomes: Their Role in Protein Breakdown (Glau-

mann, H., and Ballard, J. J., eds) pp. 115-148, Academic Press, Orlando, FL

22. Tycko, B., and Marfield, F. R. (1982) CeU 28,643-651 23. de Chastellier, C., Ryter, A., and Thilo, L. (1983) Eur. J. Cell Biol. 30,233-

24. Lenhard, J. M., Mayorga, L., and Stahl, P. D. (1992) J. Biol. Chem. 2 6 7 ,

25. Casciola-Rosen, L. A. F., and Hubbard, A. L. (1991) J. Biol. Chem. 266,

26. Murphy, R. F., Powers, S., and Cantor, C. R. (1984) J. Cell Biol. 98,9717-

243

1896-1903

4341-4347

1763 27. Stia&nger, R. M., Papahadjopoulos, D., and Hong, K. (1990) Biochemistry

28. Bush, J. M., and Cardelli, J. A. (1989) J. Biol. Chem. 264,7630-7636 29. Bronner, C., Landry, Y., Fonteneau, P., and Kuhry, J. G. (1986) Biochem-

29,4929-4939

istrv 26.2149-21 30. L& e, Y., Swaisgood, M. H., Ramos, B. V., and Steck, T. L. (1989) J. Biol.

31. Illinger, D., Poindron, P., Fonteneau, P., Modollel, M., and Kuhry, J . 4 .

32. Hoekstra, D., de Boer, T., Klappe, K., and Wilschut, J. (1984) Biochemistry

33. Padh, H., Lavasa, M., and Steck, T. L. (1991) J. Biol. Chem. 266,12123-

34. Nolta, K. V., Padh, H., and Steck, T. L. (1991) J. Biol. Chem. 266,18318-

35. Ryter, A., and Bowers, B. (1976) J. Ultrastruct. Res. 6 7 , 309-321 36. Cardelli, J. A., and Dimond, R. L. (1988) in Protein Transfer and Organelle

Biogenesis (Das, R., and Robbins, P., eds) pp 363-399, Academic Press,

37. Lagunoff D. (1964) Corn t Rend. Trau. Lab. Carkberg 34,433-446 New York

38. Klausne; R. D. (1989) &li 67,703-706 39. Brenot, F., Aubry, L., Martin, J.-B., Satre, M., and Klein, G . (1992)

40. Mancini, G. M. S., de Jonge, H. R. Galjaard, H., and Verheijen, F. W.

42. Idriss, J.-M., and Jonas, A. J. (1991) J. Biol. Chem. 266,943-1 41. Jonas, A. J., and Jobe, H. (1990) J. Blol. Chem. 266,1745-49

43. Jensen, M. S., and Bainton, D. (1973) J. Cell Biol. 66, 379-388 44. Geisow, J. M., Hart, P. D., and Young, M. R. (1981) J. Cell Biol. 89,645-

45. Sipe, D. M., Jesurum, A., and Murphy, R. F. (1991) J. Biol. Chem. 2 6 6 ,

" ~ - I ~-~~ -~

Cfem. 264,3786-3793

(1990) Biochim. Biophys. Acta 1030,73-81

23,5675-5681

12126

18323

Biochimie 74,883-895

(1989) J. Bwl. Chem. 284,15247-i5254

652

3469-3474 46. Roederer, M., Mays, R. W., and Murphy, R. F. (1989) Eur. J . Cell. Biol.

40.37-44 47. Bowks, B., and Olszewski, T. E. (1983) J. Cell Biol. 97,311-322 48. Aley, S. B., Cohn, Z. A., and Scott, W. A. (1984) J. Exp. Med. 180, 724-

49. Nilsson, J. R. (1979) in Biochemistry and Ph siology of Protozoa (Levan- 737

dowsky,, M., and Hutner, S. H., eds) 2nd Ed., Vol. 2, pp. 502-507, Academic Press, New York

50. Fok, A. K., and Shockley, B. U. (1985) J. Protozool. 32.6-9