Specific proteolysis regulates fusion between endocytic compartments in Xenopus oocytes

Cell, Vol.

51, 557-566,

November 20, 1987, Copyright 0 1987 by Cell Press

Specific Proteolysis Regulates Fusion between Endocytic Compartments in Xenopus Oocytes Lee K. Opresko and Ruth A. Karpf Department of Pathology Center for the Health Sciences University of Utah Salt Lake City, Utah 84132

Summary We examined the role of proteolytic ligand modification in endosomal targeting using vitellogenin (VTG) uptake by Xenopus oocytes as a model system. Noncleavable VTG is internalized, but does not appear in yolk platelets. We identified two inhibitors of VTG processing into the yolk proteins: the ionophore monensin and pepstatin A, a specific inhibitor of cathepsin D. Pepstatin neither affected ligand binding and internalization, nor inhibited the degradation of nonspecifically incorporated proteins, whereas monensin inhibited all of these processes. Inhibiting VTG processing prevented its deposition into yolk platelets by strongly interfering with endosome-yolk platelet fusion. Monensin treatment resulted in morphologically abnormal endosomes, while pepstatin only inhibited VTG cleavage and the subsequent fusion of endosomes with yolk platelets. Since VTG cleavage is initiated prior to its deposition in platelets, we postulate that ligand proteolysis could be necessary for normal endosomal targeting. Introduction Endocytic vesicles contain materials derived from both the cell surface and the extracellular environment. Virtually all of the externally derived materials contained in these vesicles eventually will be transferred to lysosomes, while most surface-derived material will be recycled back to the intracellular pool of membrane components (Goldstein et al., 1979, 1985; Anderson and Kaplan, 1983). The mechanisms responsible for both the sorting and the targeting of endosomes are unknown. However, owing to the specificity of this process, some kind of lysosome-endosome recognition molecules would seem to be implicated. Unfortunately, very little information is available concerning the mechanisms of vesicle-vesicle recognition in postendocytic compartmentation. A good model system in which to study endosomal targeting is the Xenopus oocyte. These cells grow to a large size primarily via the receptor-mediated uptake of vitellogenin (VTG), the yolk precursor protein. VTG is synthesized by the liver and sequestered from the maternal blood stream by oocytes in the vitellogenic stages of development (stages Ill-V; Dumont, 1972). Endocytic activity is so pronounced in these cells that >80% of the protein in a terminal size oocyte is derived from VTG (Callen et al., 1980). VTG undergoes only limited proteolysis after internalization, resulting in the formation of the four classes of

yolk proteins (Bergink and Wallace, 1974; Wiley and Wallace, 1981). These proteins are stored in a crystalline complex that forms the core of the membrane-bound, intracellular inclusions called yolk platelets. During oogenesis, yolk platelets increase in both size and number. These increases occur because a portion of the incoming, VTGcontaining endosomes fuse with preexisting yolk platelets, while the remainder are directly converted into new platelets (Brummett and Dumont, 1977; Wall and Patel, 1987; Danilchik and Gerhart, 1987). The stability of the yolk platelet population in oocytes (they are degraded only during embryogenesis) and the large numbers of endocytic vesicles found in these cells make it possible to study those factors that contribute to endosomal targeting. At first glance, the oocyte may seem to be an unusual cell type. However, its enormous size is due to relatively minor modifications of the normal endocytic pathway. Although oocytes possess large numbers of VTG receptors (up to 3 x lOi1 per 1 mm cell; Opresko and Wiley, 1987) their density on the cell surface is within the range described for some mammalian cells. The VTG receptor itself seems to be a standard, type 2 (Kaplan, 1981) calcium-dependent receptor. The tubular membrane network that has been observed to be involved in the VTG internalization pathway (Dumont, 1978) is structurally, and probably functionally, very similar to the CURL compartment (Geuze et al., 1984) described in mammalian cells. Transitional yolk bodies (terminal density endosomes; Opresko et al., 1980a) have recently been identified as multivesicular bodies (Wall and Patel, 1987), which have the same function of transporting materials to lysosomes in a variety of cell types. Yolk platelets seem to be modified lysosomes (Pasteels, 1973), and at least a subpopulation of them do contain limited subsets of proteolytic enzymes (Wall and Meleka, 1985). As is the case with lysosomes, yolk platelets are the destination of all internalized material in oocytes, including both protein and nucleic acids (Wallace and Hollinger, 1979; Opresko et al., 1979, 1980a). However, oocytes do not have typical lysosomes, although they are able to remove and degrade nonspecifically incorporated material from both “light” (protease-containing) and “heavy” (protease-negative) yolk platelets (Opresko et al., 1979, 1980a). The inability of oocytes to effectively degrade or remove VTG from platelets is apparently responsible for its accumulation in these organelles. We recently found that the proton ionophore monensin will block the terminal transfer of VTG to the yolk Platelets (Opresko and Wiley, 1987). Monensin also prevents endosome-lysosome fusion in mammalian cells (Merion and Sly, 1983). The ability of monensin to block VTG transfer was very interesting to us because one of monensin’s more pronounced effects in other cell types is to inhibit the degradation of internalized ligands (Basu et al., 1981; Wiley et al., 1985). Monensin and other compounds that raise intravesicular pH (e.g., methylamine and ammonium chloride) inhibit the proteolytic processing of internalized ligands, presumably as a secondary effect of altered pH

Cell 558

2 HOUR PULSE

PULSE + 2 I HOURCHASE

-VTG - LV-I

iFv’-2

1

PVTs

LEU CON

I Figure

PEP CHY

2

3

MON ANT

4

1. A Survey

5

APR

6

7

of Inhibitors

CON ANT APR CHY LEU PEP MON

8

9

IO

II

12 13

14

INHIBITOR LANE No.

of VTG Cleavage

Oocytes were pretreated for 2 hr in 200 pg/ml of the indicated inhibitor and then placed into a 1.0 mglml solution of [s*P]VTG containing the same inhibitor. After 2 hr, the cells were washed and a sample was placed into ice cold acetone. The remaining cells were cultured for an additional 21 hr in the indicated inhibitor plus 2.5 mglml unlabeled VTG. All cells were then processed for gel electrophoresis as described in Experimental Procedures. The photograph is of the autoradiogram of the original gel. The abbreviations are as follows: CON, control; LEU, leupeptin; CHY, chymostatin; PEP pepstatin; ANT, antipain; MON, monensin (100 PM); and APR, aprotinin. The proteins indicated on the gel are LV-1, lipovitellin 1; PV, phosvitin; LV-2, lipovitellin 2; and PVTs, phosvettes.

(Tartakoff, 1983; Haigler et al., 1985). It has widely been assumed that the accompanying inhibition of endosomelysosome fusion brought about by these agents was also a secondary effect of a relatively high intravesicular pH. However, we had previously observed that if VTG could not be cleaved, it was not transferred to the terminal density yolk platelets. This initial proteolytic processing of VTG preceded the appearance of any VTG-derived material in yolk platelets (Opresko et al., 1980a). It thus seemed possible that the inability of VTG-containing endosomes to fuse with yolk platelets in monensin-treated oocytes could be due to an inhibition of VTG proteolytic processing and not to an altered endosomal pH. We decided to critically test the hypothesis that ligand proteolysis is required for the targeting of VTG-containing endosomes to yolk platelets. We first identified a specific inhibitor of the initial VTG cleavage step and then determined the effect of this inhibition on the subsequent transfer of VTG to yolk platelets. We also developed a fluorescent technique that allows us to unambiguously observe endosome-yolk platelet fusion. The results of our experiments confirm that there is an absolute correlation between the initiation of VTG cleavage and the normal targeting of endosomes to yolk platelets. Results Evaluating Inhibitors of VTG Processing To determine whether the initiation of VTG cleavage is required for the fusion of VTG-containing endosomes with yolk platelets, we first sought to identify an inhibitor of this

process. VTG cleavage is initiated in the endosome prior to the appearance of VTG or its derivatives in the heavy yolk platelets (Opresko et al., 1980a). Previous studies had shown that internalized VTG, which could not be proteolytically processed, was not transferred to the yolk platelets. However, this noncleavable VTG was produced by partial denaturation. The lack of VTG transfer could thus have been due to the physical alteration of the molecule rather than its nonproteolyzed state. To differentiate between these possibilities, it was necessary to inhibit VTG cleavage by another method and then examine the effect of this inhibition on the translocation of the internalized molecule. We therefore tested a number of lysosomal protease inhibitors for their ability to inhibit VTG cleavage by oocytes. The inhibitors were primarily selected on the basis of their reported specificity and lack of secondary metabolic effects on treated cells (Umezawa, 1972, 1982). Oocytes were incubated with a high concentration of the different inhibitors (200 pg/ml) for 2 hr and then pulsed with 1.0 mglml [s2P]VTG plus inhibitor for an additional 2 hr. This was followed by a 21 hr chase in unlabeled VTG plus inhibitor. Samples were taken immediately following the pulse and after the chase and analyzed by SDS gel electrophoresis and autoradiography. As shown in Figure 1, most of the label from the pulse cells migrated with the same electrophoretic mobility as VTG. In most cases very little of this high molecular weight protein is seen following the 21 hr chase. Instead, the label appeared in the yolk protein products lipovitellin 1 (Mn,, 115,000), lipovitellin 2 (M,, 32,000), phosvitin (M,, 35,000), and the smaller phosphoproteins phosvettes (M,, 15,000-21,000) (Sergink and Wallace, 1974; Wiley and Wallace, 1981). However, oocytes treated with the cathepsin D inhibitor pepstatin A, as well as with monensin (Figure 1, lanes 11 and 13, respectively), still retained a large amount of unprocessed VTG after the 21 hr chase. In a number of studies we found that the only lysosomal protease inhibitor that was effective in preventing VTG processing was pepstatin A (results not shown). As expected, monensin also blocked VTG processing, presumably by raising intravesicular pH. Surprisingly, monensin was also effective in eliminating all preliminary processing of VTG that is observed immediately following the 2 hr pulse (Figure 1, compare lanes 1 and 6). To determine the minimal pepstatin concentration that was effective in preventing VTG proteolysis, oocytes were pretreated for 2 hr in concentrations of pepstatin ranging from O-100 pglml. After a pulse in labeled VTG plus the inhibitor, the cells were chased for an additional 21 hr using unlabeled VTG and pepstatin. After solubilization, the oocyte-associated VTG was evaluated by gel electrophoresis and autoradiography as shown in Figure 2. Total inhibition of VTG cleavage required a relatively high dose of 100 pg/ml. As is the case with monensin (Opresko and Wiley, 1987) oocytes probably require a high concentration of the hydrophobic pepstatin molecule because of the abundance of lipid inclusions in the cell. The necessary high concentration of pepstatin could possibly cause a general inhibition of proteolysis of all materials internalized by oocytes. To determine whether this was the case, we measured the rate of degradation

Proteolytic 559

Regulation

of Endosome

Fusion

0 Time Figure

0 Figure

Dose

of Pepstatin

on VTG Uptake

Oocytes were incubated in 2.5 mglml [32P]VTG plus or minus 100 uglml pepstatin. Samples of 10 cells each were removed at the indicated times, and the amount of acid-precipitable radioactivity in the cells was determined. Control, filled squares; pepstatin, open squares.

I 3 IO 30 60 100 PEPSTATIN CONCENTRATION (pg/mL)

2. Pepstatin

4. Effect

(hr)

Response

5

rated materials at a relatively low rate (Opresko and Wiley, 1987) the cells must be exposed to 1251-labeled proteins for longer periods of time. Loss of acid-insoluble radioactivity from v2P]casein could indicate either a protease or phosphatase activity. However, we have previously demonstrated that [s2P]VTG does not become dephosphorylated after internalization (Wallace et al., 1972; Opresko et al., 1980a; Opresko and Wiley, 1984). The data obtained using [32P]casein are shown in Figure 3. As can be seen, pepstatin-treated cells actually degraded casein at a slightly higher rate than control cells. As expected, monensin markedly inhibited the rate of casein degradation. Similar results were also obtained with both 12sl-labeled bovine serum albumin and ovalbumin (data not shown). Therefore, pepstatin seems to specifically inhibit VTG processing and not simply prevent the proteolysis of all internalized materials. It still seemed possible that pepstatin could inhibit other aspects of ligand internalization in oocytes, as was the case with monensin. Monensin treatment results in the disappearance of surface VTG receptors and a concomitant lowering of ligand uptake over time (Opresko and Wiley, 1987). We therefore measured the rate of VTG uptake in cells treated with pepstatin. The results from this experiment are shown in Figure 4. As can be seen, there is no difference in the VTG uptake rate of cells exposed to pepstatin. Since the net rate of VTG uptake is a product of the number of VTG receptors as well as their ability to bind and internalize VTG, pepstatin does not appear to affect any of these parameters.

Oocytes were incubated for 2 hr in [32P]casein containing either 100 uM monensin, 100 pglml pepstatin, or 1% DMSO. After the pulse, 10 oocytes were removed from each group, dissolved in formic acid, and precipitated in 10% TCA. The remaining cells were placed into saline containing nonradioactive casein plusor minus pepstatin or monensin. At the indicated times, 10 cells were removed from each group and processed. The values at each point represent the average of the two groups of 5 cells.

Cell Fractionation of Oocytes Treated with Different Inhibitors We next investigated whether VTG was able to be transferred to the yolk platelets in oocytes treated with pepstatin or other inhibitors. This was done by comparing the time-dependent distribution of radioactivity in cells using sucrose gradient fractionation. We have shown previously that this technique accurately reflects the transfer of mate-

Oocytes were pretreated for 2 hr with varying concentrations of pepstatin and then pulsed for 2 hr in [32P]VTG containing the indicated pepstatin concentrations. After washing, the cells were incubated for 21 hr in unlabeled VTG and pepstatin. The chased cells were fixed in acetone and processed for gel electrophoresis. This figure is the autoradiogram of the gel. The placement of the various proteins is the same as the Figure 1.

of nonspecifically incorporated [32P]casein in cells treated with monensin or pepstatin. Casein, a general substrate for proteases, was phosphorylated using Xenopus protein kinase and ATP rather than iodinated because oocytes possess an efficient deiodination activity (Opresko et al., 1980b). Since oocytes internalize nonspecifically incorpo-

I

0 Figure

3. Effect

1 of Pepstatin

’

I

1

I

0

I

I

2 3 Chase Time (hr)

4

and Monensin

Degradation

on Casein

-1

Cell 560

4

j

Control

ii+y”-

I(

YPS

%

”

0

lb

2b

3b

40

0

Fraction No. Figure 6. Effect of Monensin Yolk Platelets

0

10

20

30

40

50

60

Fraction No. Figure 5. Effect Platelets

of Protease

Inhibitors

on Transfer

of VTG to the Yolk

Oocytes (35 per group) were pretreated for 2 hr in the various inhibitors and then pulsed for 1 hr in [32P]VTG plus inhibitor. After washing, the cells were chased in unlabeled medium plus inhibitor for 8 hr. They were then homogenized and placed on 190/o-50% sucrose gradients and centrifuged as described. The resulting gradients were fractionated, and the fractions were analyzed for contained radioactivity. The dotted lines on the figure Indicate the position of the terminal density endosomes and the yolk platelets (YPs). Oocyte plasma membrane and newly formed endosomes sediment between fractions 20 and 28 (Opresko and Wiley, 1987). The sucrose concentration increases from fraction 1 to fraction 60. (The two peaks of radioactivity between fractions 1 and 10 are [s*P]VTG released during homogenization. The large, unprocessed VTG molecule sediments at the more dense region of the gradient, while processed material IS centered around the less dense peak. The relative distribution of label at the top of the gradrent is thus a convenient marker for the relative extent of VTG processing.)

rial into yolk platelets (Opresko et al., 1980a). Oocytes were pretreated for 2 hr with or without inhibitors, pulsed with [32P]VTG for 1 hr, and then chased for 8 hr. In the case of monensin, cells were exposed to a 100 FM concentration only during the chase period (to prevent premature loss of the VTG receptors). The distribution of labeled ligand in these cells is shown in Figure 5. In untreated oocytes or those treated with either leupeptin or chymosta-

and Pepstatin

on Transfer

of VTG to the

Three groups of oocytes, control (pretreated for 2 hr in 1% DMSO). monensin (no pretreatment), and pepstatin (pretreated for 2 hr in 100 uglml pepstatin), were pulsed for 1 hr in [3zP]VTG plus or minus pepstatin. The cells were then chased for 8 hr in media containing unlabeled VTG and either 100 uglml pepstatin, 100 uM monensin, or 1% DMSO. After the chase, the cells were processed for sucrose gradient fractionation as described in Figure 5.

tin, there is a bimodal distribution of radioactivity between the heavy yolk platelets and the most dense endosomes (light yolk platelets). However, in pepstatin or monensintreated cells, little VTG appears in the heavy platelet region of the gradient (see Figures 5 and 6). Thus, both agents that prevented proteolytic processing of VTG prevented the endosomes from acquiring the density of yolk platelets. (Owing to the use of wild-type animals as oocyte donors for these experiments, the results illustrated in Figures 5 and 6 are not identical.) A limitation of our cell fractionation technique is that we are only indirectly measuring the event of interest (fusion). Sucrose gradients can indicate alterations in the density of VTG-containing vesicles but cannot directly demonstrate how the alterations are occurring. To conclusively demonstrate that proteolysis is required for endosome-yolk platelet fusion, we needed a procedure that would unambiguously differentiate one yolk platelet or endosome from another. This cannot be done biochemically, since organelles of the same type have a uniform composition. It can, however, be done visually by using fluorescent VTG and microscopic sections of oocytes.

Proteolytic 561

Regulation

Table 1. The Effect Uptake by Oocytes

of Endosome

of Fluorescent

Fusion

Labeling

% Inhibition VTG Uptake

Sample Rhodamine Rhodamine Rhodamine Rhodamine Rhodamine Dansylaziridine Dansylaziridine

0’ 5 10 20 40

2 HOURPULSE (before cleoroqe)

on VTG

PI HOURCHASE (after cleovo9e)

4

b

of

68 61 59 41 12 0 (6 mol per VTG dimer)

53 52

Oocytes were incubated in a set concentration of radiolabeled VTG to which were added VTG preparations exposed to different molar ratios of fluorescent dyes. The ability of the different fluorescent VTG preparations to compete for uptake with the radiolabeled VTG was then determined. The more competitive the preparation, the greater the observed degree of inhibition of uptake and the less harmful the modification of the VTG molecule. a The numbers listed in the sample column are the molar ratios of dye molecules to VTG monomer; there are 133 lysines per VTG monomer (Wiley and Wallace, 1981).

Preparing Fluorescent VTG to Visualize Endosomal Fusion Directly Most fluorescent dyes are coupled to proteins via e-amino groups of lysine. Since previous studies had indicated that modification of lysine residues in VTG can affect its binding to oocytes (Opresko et al., 1980b), we modified VTG to different degrees with several different fluorescent dyes and then examined its ability to compete with unmodified VTG for uptake by oocytes. Shown in Table 1 are the results obtained with rhodamine and dansylaziridine. Even a low level of modification by rhodamine resulted in some loss of the ability of the labeled molecule to compete for uptake (higher levels of inhibition were obtained with VTG coupled to either lucifer yellow or fluorescein; data not shown). However, modification of VTG with dansylaziridine had no effect on its uptake. This dye reacts with free sulfhydryl groups rather than amino groups, making it possible to couple only 4-6 dye molecules to each VTG dimer (Scouten et al., 1974). Due to this low level of dye substitution, it was difficult to use dansylaziridine-VTG as a probe for endosomal translocation. However, dansylaziridine-VTG was visible in oocyte sections following lengthy incubations and did provide us with a noncompromised standard for evaluating the intracellular distribution of other labeled VTG preparations. Having determined that fluorescent VTG was recognized by oocytes, we next examined its internalization and processing into the yolk proteins. This was done by incubating oocytes for 2 hr with VTG labeled with both fluorescent dye and 32p followed by a 21 hr chase in medium containing unlabeled VTG. As an internal control we also incubated cells in [32P]VTG preparations that were exposed to the same conditions required for fluorescent labeling, but to which no dye was added. The degree of proteolytic processing of the different preparations was then evaluated by SDS gel electrophoresis and autoradiography. As shown in Figure 7, there was no difference be-

CON AZR-CCM AZR RH-CON RH

Figure 7, Fluorescent Processing

Labeling

CONAZR-CC4 AZR RH-CON

RH

of VTG Does Not Alter Its Intracellular

VTG was labeled either with rhodamine isothiocyanate (RH) using a 10x molar excess of dye to vT(; monomer or with dansylaziridine (AZR) using a 3x molar excess of dye to VTG monomer. Duplicate VTG samples were also incubated under similar conditions in each case but without dye (RH-CON and AZR-CON). The resulting preparations were then labeled with 32P, and an additional VTG sample was also radiolabeled (CON). Oocytes were incubated in the different labeled ligands for 2 hr, a sample was removed from each group, and the remaining cells were maintained for 21 hr. The cells were processed as described in Figure 1 and electrophoresed. The resulting dried gel was exposed for autoradiography.

tween the VTG labeled with the different fluorescent dyes either immediately following the pulse or after the 21 hr chase. In addition, further studies showed that rhodamine-VTG (Rh-VTG) was correctly transferred to the yolk platelets as determined by sucrose gradient fractionation (results not shown). Thus, the coupling of rhodamine to the VTG molecule does not significantly alter its intracellular processing or transfer to the yolk platelets. Effect of Inhibitors on Endosome-Yolk Platelet Fusion With the successful completion of our preliminary studies we could critically test our hypothesis that inhibitors of VTG cleavage would inhibit the fusion of endosomes with yolk platelets. We pretreated cells for 2 hr either with or without the different lysosomal protease inhibitors and then pulsed them for 2 hr with Rh-VTG in the presence of inhibitors. Some oocytes were fixed in glutaraldehyde immediately following the pulse, while others were chased for 8 and 24 hr in the presence of inhibitors and unlabeled VTG. The oocytes were embedded in plastic and sectioned at 1.5-2.0 pm. The results are shown in Figures 8A-8F. All oocytes looked very similar after the 2 hr pulse with Rh-VTG. The endosomes appear as very bright fluorescent spots (vesicles) at the cell periphery (see also Figure 9A). Very little endosome-platelet fusion was observed. However, by 8 hr the fusion of endosomes with yolk platelets could be clearly seen in the control cells as crescents of fluorescent material at the surface of some

Cell 562

Proteolytic 563

Regulation

of Endosome

Fusion

platelets (Figures 8C and 8E; Figure 9B, arrowheads). The same pattern was seen with oocytes treated with any of the protease inhibitors except for pepstatin. In pepstatintreated oocytes very few platelets contained Rh-VTG, although the endosomes had moved deeper into the oocyte interior. By 24 hr, some endosomes had fused with platelets in pepstatin-treated cells (Figure 8F). However, the fused material did not take the shape of a crescent, but instead remained in a more or less discrete spot. In contrast, a large percentage of the platelets in control cells (or those treated with other protease inhibitors; the cell shown in Figure 8E was treated with chymostatin and has an appearance identical to that of a control cell) contain Rh-VTG, which by this time is more evenly distributed around the entire surface of the platelet. This indicates either that the platelets have fused with multiple endosomes, or that the newly incorporated Rh-VTG has been redistributed throughout the crystalline yolk platelet surface. An examination of VTG cleavage in pepstatin-treated cells using SDS gel electrophoresis showed that most of the Rh-VTG was intact after the 24 hr chase, while virtually all had been processed by control cells (data not shown). It thus appears that the inhibition of VTG cleavage by pepstatin strongly inhibits, but does not totally prevent, the fusion of endosomes with yolk platelets. However, VTG cleavage did seem essential for the redistribution of the protein into the crystalline structure of the growing platelets. In the above studies, we only examined the effect of lysosomal protease inhibitors. To determine whether monensin had a similar effect on endosome-yolk platelet fusion, we also examined oocytes exposed to Rh-VTG plus 100 FM monensin for 2 hr. Cells could not be pretreated with monensin because of its inhibitory effect

Figure

6. Uptake

and Compartmentation

of Rh-VTG

in Control

on VTG incorporation. As shown in Figure lOA, endosomes in monensin-treated oocytes appeared quite different from those in either control or pepstatin-treated cells. Not only were there far fewer endosomes, but they were also much larger (compare with the section in Figure 106 of a control cell after a 2 hr pulse in Rh-VTG). In addition, the penetrance of these large endosomes into the interior of the cell was much more limited than those in either control or pepstatin-treated cells. We have previously shown that the density distribution of endosomes from monensintreated oocytes is not significantly different from untreated cells, and thus their greater size is probably not due to simple osmotic swelling. Instead, it appears more likely that monensin treatment reduces the number of forming endosomes with a consequent increase in the amount of ligand that each contains. The inhibition of vesicle movement could also reflect a general perturbation of endosomal formation and targeting similar to that observed in mammalian cells. Discussion We have tested the hypothesis that ligand processing can regulate the fusion of endosomes with the terminal endocytic compartment in oocytes. This was done by examining the effects of inhibitors of VTG cleavage on the postendocytic compartmentation of the ligand. We found that the cathepsin D inhibitor pepstatin A was able to prevent ligand processing and did indeed inhibit the fusion of VTG-containing endosomes with yolk platelets. Protease inhibitors that did not block the cleavage of the molecule had no effect on the fusion process. The efficacy of pepstatin in inhibiting both VTG cleavage and endosomal fusion was surprising since this inhibitor is one of the most

and Pepstatin-Treated

Cells

(A) Section through an oocyte following a 2 hr incubation in Rh-VTG (magnification, x1600). (6) Section through an oocyte pretreated with 100 rig/ml pepstatin for 2 hr and then incubated in pepstatin plus Rh-VTG for 2 hr (magnification, x2000). (C) Section through an oocyte incubated 2 hr in Rh-VTG and then chased for 6 hr in culture medium containing unlabeled VTG (magnification, x1725). (D) Section through an oocyte pretreated with pepstatin, pulsed with Rh-VTG plus pepstatin, and then chased for 6 hr in culture medium plus pepstatin (magnification, x1600). (E) Section from an oocyte pretreated for 2 hr in 200 nglml chymostatin, incubated in Rh-VTG plus chymostatin for 2 hr, and then chased for 24 hr in culture medium plus chymostatin (magnification, x1600). (F) Section from an oocyta pretreated with pepstatin, pulsed with Rh-VTG plus pepstatin And chased for 24 hr in culture medium plus pepstatin (magnification, x2100). N indicates a nucleus.

Cell 564

Figure

10. A Comparison

of Endosome

Size and Mobility

(A) Section from a cell pulsed for 2 hr in Rh-VTG (magnification, x950). N Indicates a nucleus.

in Control

and Monensin-Treated

plus 100 NM monensin

specific protease inhibitors we tested (Umezawa, 1972, 1982). In other studies on the inhibition of ligand processing, it is generally leupeptin that is most inhibitory (Ascoli, 1979; Savion et al., 1980; Wiley et al., 1985; Berg et al., 1985). Leupeptin, however, will inhibit the action of both serine and cysteine proteinases, while pepstatin is very specific for aspartic proteinases (Umezawa, 1982). The major intracellular protease in this class is cathepsin D, and pepstatin binds to cathepsin D with very high affinity (Rich, 1986). This binding is so specific that derivatives of pepstatin are used as histochemical reagents to localize cathepsin D in mammalian cells (Mathews et al., 1981a, 1981b; Yamato et al., 1984). A trivial explanation for our results would be that we were preventing VTG processing by inhibiting the fusion of the endosomes with platelets and not visa versa. However, this explanation is very unlikely for the following reasons. First, we have previously established that the initial proteolysis of VTG occurs in the endosomes prior to yolk platelet fusion (Opresko et al., 1980a). Data from other investigators have also indicated that VTG processing occurs in discrete phases, with the first steps occurring prior to the time that internalized VTG is found in platelets (Jared et al., 1973; Wail and Patel, 1987). A reexamination of the timing of the different VTG cleavage events has revealed that cleavage of the carboxyl terminus of VTG (the LV2 domain) is completed between 1 and 2 hr after internalization. In comparison, only 30% of the VTG is transferred to the heavy yolk platelets by 2 hr (Wiley and Wallace, submitted). Only the final processing of the phosphorylated regions of VTG seem to unambiguously occur in the platelet compartment. Significantly, the site of the initial cleavage (between the phosvitin and LV2 domains of VTG) is next to an appropriate cleavage site for cathepsin D (Byrne et al., 1984; Beynon and Bond, 1986). Sec-

(magnification,

Cells x950).

(B) Sectlon

from a cell pulsed

for 2 hr in Rh-VTG

ond, pepstatin treatment does not affect the processing of other, nonspecifically incorporated proteins. If pepstatin treatment prevented other protease-containing compartments from fusing with endosomes, we should have seen a general inhibition of the degradation of internalized materials. Our assay for nonspecific degradation of proteins was apparently effective since it accurately showed that monensin is a general inhibitor of endosomal and lysosomal proteolysis. Diment and Stahl (1985) recently described the degradation of mannose-BSA in mammalian macrophages by a pepstatin-sensitive protease localized to a prelysosomal, endosomal compartment. It is therefore possible that the processing of VTG takes place via a similar enzyme in oocytes. Third, we have shown that partial denaturation of VTG will block both its processing and its subsequent transfer to platelets. It is difficult to envision how an alteration in the VTG molecule itself could affect any other compartment besides the endosomes. Finally, the eventual transfer of uncleaved VTG to the yolk platelets in pepstatin-treated cells (less than 20% after a 24 hr chase) and its lack of subsequent processing (as determined by SDS gel electrophoresis; data not shown), indicate that endosome-platelet fusion does not in itself result in VTG processing. The simplest explanation for all of the above observations is that VTG proteolysis within the endosomes is required to direct the efficient fusion of those structures with yolk platelets. Ligand proteolysis could regulate endosome fusion and targeting by several different mechanisms. Proteolysis could generate a specific regulatory fragment of VTG that may confer fusion competency to the vesicle. However, the cleavage step that unambiguously occurs in endosomes produces two large polypeptides (32,000 M, and >150,000 M,). It seems unlikely that they could possess regulatory properties significantly different from the par-

Proteolytic 565

Regulation

of Endosome

Fusion

ent molecule, since most regulatory components derived from proteolytic processing of large precursors are small peptides (Plow et al., 1982; Edington et al., 1985; Seya and Nagasawa, 1985). VTG-derived fragments of similar size are not produced until after the endosomes have fused with platelets (Wiley, 1979). Another possibility is that the VTG-cleaving enzyme degrades a specific inhibitor of endosomal fusion. Thus, once cleavage is initiated, the fusion inhibitor is also hydrolyzed. This hypothesis is also unlikely since we previously found that partial denaturation of VTG inhibits endosome-platelet fusion. If the VTG-cleaving enzyme were actually acting upon a second protein, it seems unlikely that a partially denatured ligand should block this process. An alternate hypothesis is that VTG cleavage removes the ligand from its receptor and that the absence of occupied receptors render endosomes capable of fusing with platelets. This seems plausible because there should be relatively few occupied VTG receptors in terminal stage endosomes. These vesicles undergo continuous condensation and self-fusion prior to attaining a terminal density. Concomitantly, membrane is retrieved and there is an overall reduction in both surface area and receptor number per vesicle. By the time VTG processing is initiated, almost all endosomal VTG is condensed in the vesicle lumen. However, because of the extremely high ligand concentration in these vesicles, all remaining VTG receptors should be occupied. These remaining receptors could become unoccupied by a combined process of VTG proteolysis, a low pH, and additional VTG condensation and membrane retrieval. The resulting loss of the remaining occupied receptors would make the vesicles “competent” for fusion with platelets. Since pepstatin would only prevent proteolysis, occupied receptors could eventually become unoccupied via the remaining two processes. This would explain why there is a low level of endosomal fusion after 24 hr in pepstatin-treated cells despite the lack of VTG processing. There are several aspects of the last hypothesis that make it most attractive. First, it is compatible with our previous observations on partially denatured VTG. Second, if the state of VTG receptors directs vesicle fusion, this could confer the appropriate specificity to the incoming endosomes with regard to their contents. Third, the entire compartmentation pathway would be geared to the maximal recovery of the recycling VTG receptors. Although the majority would be retrieved during the condensation and endosome-endosome fusion portion of the pathway, the remaining receptors could be retrieved when ligand proteolysis severed the ligand-receptor complex. Under this scenario, the majority of the endosome membrane and receptors would have to be removed prior to fusion with platelets and lysosomes, which would contain the truly destructive proteases. These would be responsible for the further processing of the VTG molecule. Previous studies indicate that proteases do exist within the yolk platelets, but the exact nature or specificity of these enzymes is presently unknown (Opresko et al., 1979; 1980a). The regulation of compartment fusion by proteases may be more widespread than previously suspected. Pro-

teases are known to be involved in many translocation processes. For example, signal peptidase is involved in the correct addressing of secretory proteins in both eukaryotic and prokaryotic cells (Walter et al., 1984; Dalbey and Wickner, 1985,1987). In addition, many secretory proteins are synthesized in a precursor form (Blobel and Dobberstein, 1975). These proproteins are often found in specific cellular locations (Orci et al., 1986), and many also contain regulatory portions that are liberated via the action of specific proteases (Douglass et al., 1984). A more direct demonstration of the involvement of proteolysis in the fusion between membrane-bound compartments has been provided by Mundy and Strittmatter (1985). These authors have shown that a metalloendoprotease activity is required for the fusion of secretory granules with the plasma membrane in mast cells and chromaffin cells. Agents that disrupt intravesicular pH gradients affect both proteolytic processing and protein compartmentation. For example, the proton ionophore monensin inhibits the processing of secretory protein precursors as well as their secretion (Orci et al., 1984; Steiner et al., 1984). It also prevents ligand proteolysis and the transfer of ligands to the lysosomes (Merion and Sly, 1983; Berg et al., 1985). Monensin elevates intravesicular pH by catalyzing the exchange of cytoplasmic Na+ or K+ for H+. This can in turn affect multiple vesicular parameters. These include the activity of “lysosomal” proteases, the dissociation of ligands from their receptors, the ionic composition of the vesicle lumen, as well as the conformation of transmembrane proteins. It is difficult to determine which of these parameters are involved in specific aspects of endosomal function. Mutant cells that are defective in the transfer of material to lysosomes have been described (Haigler et al., 1985). It has been proposed that this defect is due to an inability to properly acidify the endosomal compartment (Merion et al., 1983). However, measurement of vesicular pH has shown that there is very little difference between control and mutant cells (Merion et al., 1983; Robbins et al., 1984). Yet, the difference in both endosomal, and especially lysosomal, ligand degradation is very large. Rather than the mutant cells being defective in endosomal acidification, they may have a defect in endosomal protease function. This would result in no preprocessing of ligand, which could in turn inhibit ligand transfer. Although we have postulated here that proteolysis can regulate endosomal targeting, proteolysis per se is probably not the essential aspect of this process. Rather, it is more likely that the state of the receptors or other transmembrane proteins are the actual regulatory entities. The association of VTG with its receptor is not particularly pHsensitive (Opresko and Wiley, 1987) so that ligand proteolysis may play a central role in dissociating the ligandreceptor complex. However, it is essential to define those molecules whose function is critically dependent on the appropriate environment and how their function translates itself into vesicle action. Our evidence suggests that intravesicular proteases may be centrally involved in some of these actions. How widespread the role of proteases is in vesicular targeting remains an open question at this time, since very few studies have been directed toward this

Cell 566

topic. However, further investigation on the role of cathepsin D in endosomal targeting in oocytes should provide us with valuable insights. Experimental

Procedures

Materials Large, female Xenopus laevis were purchased from the South African Snake Farm (Fish Hoek. Cape Province). They were maintained in running, dechlorinated tap water and fed twice weekly with shredded beef supplemented with vitamins (100x MEM vitamins, GIBCO). Chemicals were from Sigma Chemical Co., with the exception of HEPES buffer (Ultrol grade) from CalBiochem and acrylamide from BioRad. [szP]ATP (2500-3000 Cilmmol) was from Amersham or New England Nuclear. Culture medium was from GIBCO, and calf serum was from KC Biological. Protein Labeling VTG was phosphorylated in serum obtained from estrogen-stimulated animals (2-4 mg 8-estradiol per animal) using Xenopus protein kinase as previously described (Opresko and Wiley, 1984). After labeling, the VTG was dialyzed against saline solution O-R2 (Wallace et al., 1973) and stored at 0%. Each batch of VTG was used within 72 hr of labeling. The average specific activity of the labeled VTG preparations was 100 nCi per mg of VTG. Casein was also phosphorylated with Xenopus protein kinase and dialyzed against O-R2 prior to use. Rh-VTG was prepared by dissolving rhodamine isothiocyanate in DMSO prior to its addition to the VTG. The stock rhodamine solution was formulated such that less than 5% of the total volume of the reaction mixture was composed of the dye. Rhodamine was added at various molar concentrations ranging from 5x to 40x the molar concentration of VTG. The rhodamine coupling buffer was 100 mM phosphate buffer (pH 8.5). Labeling was carried out in the dark for 4 hr at room temperature. Dansylaziridine-VTG was formulated using a 3:l molar ratio of dansylaziridine to VTG monomer. The dansylaziridine was dissolved in 100% methanol, and the buffer used was 100 mM Tris (pH 9.0). After labeling, the fluorescent VTGs were dialyzed against water. The dialyzed samples were then used in the formulation of the standard oocyte culture medium. Oocyte Isolation and Incubation All cells within a single experiment were derived from the same ammal. The ovary was excised under sterile conditions and placed into sterile solution O-R2. The cells were removed by manual dissection with watchmaker’s forceps and were measured using an ocular micrometer mounted in a Wild M-5 dissecting microscope. Cells, 0.88-1.00 mm in diameter, were placed into 50% Liebovitz-15 medium containing 5.4 x 10-s M VTG but lacking any hormones (Wallace and Misulovin, 1978). The cells were cultured overnight, and any morphologically abnormal cells were discarded prior to experimental measurements. To ensure that the results were comparable between different experimental groups, the composition of cell sizes among the different groups was kept constant. The degree of denaturation of the different fluorescent VTG preparations was determined by their ability to compete for uptake by oocytes with [32P]VTG. Oocytes were placed into 5.4 x lo-’ M [32P]VTG plus or minus a 3- to 5-fold molar excess of test VTG. The amount of VTG Incorporated after l-2 hr was then determined and compared with the amount Internalized by oocytes incubated in the same concentration of untreated, competing VTG. Each determination was made using 10 oocytes per point. After incubation, the cells were washed 3 times in saline and dissolved, in groups of 5, in 100 nl of 88% formic acid. The dissolved samples were placed onto Whatman 3MM filter paper disks along with an 80 nl rinse, dried, and immersed in ice cold 10% TCA containing 10 mM sodium pyrophosphate. After 18 hr at 4%, the disks were rrnsed once in 10% TCA, in 2 changes each of alcohol ether (3:1), and in ether and then air dried. The dried disks were counted on a Packard scintillation counter The effects of various protease Inhibitors on VTG cleavage and protein degradation were determined using cells pretreated with the desired inhibitor for 2 hr. Cells were sampled Immediately following a l-2 hr incubation in [32P]VTG (2.17 x 10e6M) plus inhibitor and

processed for either scintillation counting or gel electrophoresis. The remaining cells were placed in nonradioactrve medium contarning the inhibitor for various lengths of time. A similar regrmen was followed for obtaining samples for sucrose gradient fractionation. In this case, the cell samples were homogenized after washing and then placed on top of a 19%-50% gradient After 12 hr at 37,500 rpm, the gradients were fractionated and processed as previously described (Opresko et al., 198Oa). Gel Electrophoresis Control and test cells were immersed in ice cold acetone for 15 hr, rinsed in 1 change of acetone, and air dried. They were then borled for 2 min in 2% SDS, 1 mM EDTA, 10% glycerol, bromphenol blue, 20 mM Tris (pH 6.8). After boiling, the samples were kept overnight at room temperature, reboiled for2 min, reduced with 10 mM dithiothreitol, and boiled for another min. The samples were diluted and placed on 5%-15% gradient polyacrylamide gels and electrophoresed until the tracking dye was 1 cm from the bottom of the gel. After fixing, staining, and destaining, the gels were dried and exposed for autoradiography at -70% using Kodak XAR-5 X-ray film and a Cronex Lightning Plus intensification screen. Histological Processing Cells Incubated in fluorescent VTG were washed and fixed in 3% glutaraldehyde. 100 mM cacodylate buffer (pH 7.2) for 2 hr at room temperature. After rinsing in buffer, the cells were dehydrated through 100% alcohol and embedded in JB-4 resin (Polysciences) or Historesin (LKB). Sectrons were cut on an LKB HistoRange microtome at 1.5-2.0 nm and examined with a Zeiss universal microscope equipped with epifluorescence. Micrographs were taken using either Tri-X pan or Ektachrome film (Kodak). Acknowledgments The authors wish to thank Christina Samathanam for excellent technical assrstance with the fluorescent labeling of VTG and H. Steven Wiley for helpful discussions and critical review of this manuscript This work was supported, in part, by grant GM32992 from the National Institutes of Health to L. K. 0. The costs of publication of this article were defrayed rn part by the payment of page charges. This article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to Indicate this fact. Recerved

April 28, 1987; revised

September

8, 1987.

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