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© The Rockefeller University Press,
0021-9525/2000//901 $5.00
The Journal of Cell Biology, Volume 149, Number 4,
, 2000 901-914
Original Article |
Distinct Membrane Domains on Endosomes in the Recycling Pathway Visualized by Multicolor Imaging of Rab4, Rab5, and Rab11
sonnichs{at}embl-heidelberg.de
Two endosome populations involved in recycling of membranes and receptors to the plasma membrane have been described, the early and the recycling endosome. However, this distinction is mainly based on the flow of cargo molecules and the spatial distribution of these membranes within the cell. To get insights into the membrane organization of the recycling pathway, we have studied Rab4, Rab5, and Rab11, three regulatory components of the transport machinery. Following transferrin as cargo molecule and GFP-tagged Rab proteins we could show that cargo moves through distinct domains on endosomes. These domains are occupied by different Rab proteins, revealing compartmentalization within the same continuous membrane. Endosomes are comprised of multiple combinations of Rab4, Rab5, and Rab11 domains that are dynamic but do not significantly intermix over time. Three major populations were observed: one that contains only Rab5, a second with Rab4 and Rab5, and a third containing Rab4 and Rab11. These membrane domains display differential pharmacological sensitivity, reflecting their biochemical and functional diversity. We propose that endosomes are organized as a mosaic of different Rab domains created through the recruitment of specific effector proteins, which cooperatively act to generate a restricted environment on the membrane.
Key Words: endocytosis • transferrin recycling • Rab proteins • EEA1
© 2000 The Rockefeller University Press
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Introduction |
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Considering the structural and functional properties of endosomes, several problems arise for the distinction of these two membrane pools by cargo accessibility and morphological criteria. First, the flow of cargo through endosomes becomes increasingly asynchronous with time. Second, the spatial separation between early and recycling endosomes is not apparent in all cell types. For example, transferrin-accumulating membranes are more concentrated around the microtubule organizing center in CHO cells than in A431 cells. Third, endosomal membranes are extremely pleomorphic (Griffiths et al. 1989) and their morphology varies between cell types (Hopkins et al. 1990; Yamashiro and Maxfield 1984). Carrier vesicles for recycling cargo have not been characterized. Consequently, by the so far applied criteria, tubules and vesicles of the recycling endosome cannot be distinguished from transport intermediates.
A more accurate definition of endosomal organelles must therefore include their molecular machinery. Early and recycling endosomes are at the crossroad of several transport routes. They continuously exchange membrane with late endocytic compartments, the plasma membrane, and the TGN (Ghosh et al. 1998), and, in addition, undergo homotypic fusion and fission reactions. All these trafficking events tend to average the biochemical composition over time. How can endosomes then maintain their functional identity?
A group of regulatory molecules localized to distinct subsets of membranes along the secretory and endocytic pathway are members of the Rab family of small GTPases (Pfeffer 1994; Novick and Zerial 1997). In the GTP-bound state Rab proteins recruit specific effector proteins to the membrane. These have multiple functions, ranging from membrane budding (McLauchlan et al. 1998) or docking (Christoforidis et al. 1999a; Guo et al. 1999) to interactions with the cytoskeleton (Echard et al. 1998; Nielsen et al. 1999). Interference with their GTPase cycle provokes drastic morphological and functional changes in the target organelles, suggesting that Rab proteins contribute to the organization of membranes. One of the best-studied Rab proteins on endosomes is Rab5. It is required for the delivery of material from the plasma membrane to early endosomes as well as homotypic endosome fusion (Gorvel et al. 1991; Bucci et al. 1992; McLauchlan et al. 1998). We recently identified a surprisingly large number of cytosolic proteins binding specifically to the activated form of Rab5 (Christoforidis et al. 1999a). Rather than serving completely distinct functions, these Rab5 effectors appear to act cooperatively. Upon activation by the Rabaptin-5/Rabex-5 complex (Horiuchi et al. 1997), Rab5 can recruit phosphoinositol-3 kinases (Christoforidis et al. 1999b), including hVPS34 (Schu et al. 1993), ensuring the local generation of phosphoinositol-3-phosphate (PI(3)P). An environment of PI(3)P and active Rab5 is essential for the binding of the docking protein EEA1 (early endosomal antigen 1) to the endosomal membrane (Patki et al. 1997; Mills et al. 1998; Simonsen et al. 1998; Christoforidis et al. 1999a). EEA1 itself directly interacts with the SNARE syntaxin13 in a dynamic, oligomeric complex (McBride et al. 1999).
Rab5 can therefore be seen as a regulatory anchor for factors that create a restricted, fusion competent domain on the early endosome (McBride et al. 1999). This working model can explain how endosomes might be organized to sustain extensive membrane exchange without compromising their structural and functional integrity. If generalized for other family members, the model would predict that Rab proteins with their specific set of effectors can contribute to a compartmentalization within a continuous membrane structure. A functional arrangement of machineries controlling vesicle targeting, protein sorting or cytoskeletal interactions into domains would help to increase the efficiency and coordination of these processes.
Apart from Rab5, two other Rab proteins have been implicated in the recycling pathway, Rab4 and Rab11. Rab4 was suggested to play a role in fast recycling from early endosomes back to the plasma membrane (Van der Sluijs et al. 1992, Sheff et al. 1999). Rab11 has been localized to the recycling endosome, the TGN and specialized storage membranes of regulated secretory pathways (Urbe et al. 1993; Ullrich et al. 1996; Jedd et al. 1997; Calhoun et al. 1998). Mutants of Rab11 effect transferrin recycling (Ullrich et al. 1996; Ren et al. 1998) and transport of vesicular stomatitis virus glycoprotein (VSV-G) to the plasma membrane (Chen et al. 1998). Our model would predict that Rab4 and Rab11 should be enriched in membrane domains distinct from that of Rab5, but that can reside on the same endosome. Could different arrangements of these domains account for morphologically and kinetically distinct transferrin recycling structures?
To test this hypothesis we have visualized Rab4, Rab5, and Rab11 in combination with endosomal cargo. Our results provide evidence for the proposed domain organization of endosomes.
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Materials and Methods |
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Biochemical Transferrin Recycling Assay
Iron-saturated human transferrin and iron-saturated biotin-labeled transferrin were purchased from Sigma-Aldrich.
A431 cells were serum starved for 2 h to deplete endogenous transferrin. Biotinylated transferrin was internalized at 10 µg/ml for 2 min at 37°C.
Unbound and surface-bound transferrin was removed by washing the cells with ice-cold PBS containing 1 mg/ml unlabeled transferrin followed by low pH buffer (150 mM NaCl, 10 mM acetic acid, pH 3.5, and 1 mg/ml unlabeled transferrin). This procedure resulted in the removal of 95–98% of surface-bound ligand. To measure recycling, cells were returned to 37°C in the presence of 1 mg/ml of unlabeled transferrin. At each time point the medium was collected, cells were washed with ice-cold PBS and homogenized in 1 ml of lysis buffer (PBS containing 2% Triton X-100 and 0.4% SDS). 5 µl of the supernatant (recycled biotinylated transferrin), or 5 µl of the lysate (cell-associated biotinylated transferrin) were quantified as described previously (Zacchi et al. 1998). Results were expressed as percent of the total signal.
Internalization of Fluorescently Labeled Transferrin and Immunofluorescence
A431 cells were grown on glass coverslips and serum starved for 1 h before incubation with 60 µg/ml fluorescently labeled transferrin (Molecular Probes). The probe was chased in the presence of a fivefold excess of nonlabeled transferrin.
Immunofluorescence labeling was performed according to standard procedures. Antibodies against transferrin receptor and TGN46 were diluted 1/100, EEA1 antiserum was used at 1/500, Rab11 antibody at 1/200 dilution. Cells were mounted in ProLongTM Antifade (Molecular Probes).
Incubations with 5 µg/ml brefeldin A (Sigma-Aldrich) or 50–100 nM wortmannin (Sigma-Aldrich) were performed in serum-free medium.
Microinjection
Plasmid DNAs encoding the Rab proteins were injected into the cell nucleus using an Eppendorf micromanipulator and transjector. Compared with transient transfections, we found that this technique ensured a better control over expression levels in general and a constant ratio of the two fusion proteins in particular. Expression of the CFP was generally lower compared with the YFP fusion proteins. A mixture of 7 ng/µl YFP and 20 ng/µl CFP plasmid DNA when monitored 6–14 h after injection resulted in similar expression levels as seen in the stable cell lines.
Live Cell Imaging
Cells were grown on life cell imaging dishes (MatTek Corp.). Imaging was performed in CO2-independent medium (GIBCO BRL). Time-lapse series were acquired at 37°C on an inverted Olympus IX70 microscope, equipped with a 100x oil immersion objective, NA 1.35, and a 12-bit Till Imago CCD camera (Till Photonics). The temperature was controlled by a climate box covering the set up. Filters (AHF Analysentechnik) allowed for simultaneous detection of CFP and YFP or GFP and rhodamine, respectively. Switching between the excitation wavelengths with a Polychrome II multiwavelength illumination system was controlled by the TillvisION 3.3 software (Till Photonics). Exposure times were 200–300 ms for each channel followed by a 60-ms readout delay, resulting in time-lapse sequences of roughly two frames per second, which were merged as RGBs using TILLvisION. Series were exported as single TIFF files, processed in Adobe Photoshop 5.0 or converted into QuickTime movies using IPLab 3.2 (Scanalytics Inc.).
Confocal Microscopy, Image Processing, and Quantitation of Signal Overlap on Fixed Cells
Images were acquired on the Compact Confocal Camera (CCC) as described by White et al. 1998, using a 63x 1.4 NA Plan-Apochromat III DIC objective (Carl Zeiss). Fluorescent dyes were imaged sequentially in frame-interlace mode to eliminate cross talk between the channels. CFP was excited with a 430-nm laser line (Directly Doubled Diode/D3; Coherent) and imaged through a combination of 440–505-nm bandpass and 525-nm longpass emission filters. YFP was exited with the 514-nm Argon laser line and imaged through a 525-nm longpass emission filter. Texas red was excited with the 594-nm Helium Neon laser line and imaged through a 610-nm longpass emission filter.
Serial sections were acquired satisfying the Nyquist criteria for sampling and processed on a multiprocessor SGI Unix computer using the Huygens System 2.2 (Scientific Volume Imaging BV). A maximum likelihood estimation (MLE)-based algorithm was used for image reconstruction. Z-stacks of images were exported as TIFF files, and individual sections were analyzed for fluorescent signal overlap by visual inspection. Using Adobe Photoshop 5, a 5 x 5-cm grid was projected onto the image of the reference channel, and in every second grid square all fluorescent structures were marked on a separate, superimposed layer. Signals were referred to as individual structures if they comprised of a continuous patch of intensity values >50 (in a range of 0–255). This layer was then projected onto the corresponding images for the other two channels, and the underlying image was analyzed for fluorescent signal at the marked position. At least two sections per cell were counted, ensuring that peripheral and perinuclear structures were equally taken into account. 5–10 cells were analyzed, counting 150–200 fluorescent structures per cell.
Electron Microscopy
A431 cells stably expressing Rab4- or Rab5-GFP were labeled with BSA-gold (5 nm) at an OD(520) of 0.5. Cells were washed extensively and harvested with a cell scraper. After homogenization (3 mM imidazole, pH 7.4, 250 mM sucrose, 2 mM MgCl2) with a 22.5-gauge needle, unbroken cells and nuclei were sedimented at 1,000 g for 10 min. The resulting PNS was supplemented with an ATP regenerating system (10 mM ATP, 80 mM creatine phosphate, 0.4 mM creatine kinase) and 1 mM GTP and incubated with carbon-coated copper grids at room temperature for 15 min. After fixation with 2% (wt/vol) paraformaldehyde for 10 min, unspecific binding sites and remaining fixative were blocked with 0.8% (wt/vol) BSA, 0.1% (vol/vol) fish skin gelatin, and 20 mM glycine in PBS for 10 min. Antibodies against EEA1, GFP, and Rab11 were diluted in blocking solution at 1:50, 1:200, and 1:30, respectively. Protein A conjugated to 10-nm gold was diluted according to the manufacturer's instructions (Slot and Geuze, University of Utrecht). The first antibody/protein A complex was fixed with 1% (vol/vol) glutaraldehyde in PBS for 10 min. After quenching of the fixative with 20 mM glycine in PBS, the immunolabeling procedure was repeated with the second antibody and protein A conjugated to 15-nm gold. Finally, the material was stained with 2% methyl cellulose and 3% uranyl acetate in a mixture of 1:3 for 5 min at 4°C. Specimen were observed on a Philips CM120 BioTwin transmission electron microscope.
For quantitation, random images of membrane-containing fields were taken at 27,000 magnification. Endosomes were identified by the internalized BSA-gold. 50–100 endosomes on ten images were analyzed for immunogold labeling, and the results were expressed as a percentage of the total identified endosomes.
Online Supplemental Material
All videos were acquired and processed as described previously in Live Cell Imaging. They comprise 40–60 frames, animated at 10 frames/s (
10x acquisition speed). The videos can be found at http://www.jcb.org/cgi/content/full/149/4/901/DC1.
Movies 1–3.
Rab4, Rab5, and Rab11 occupy subdomains on transferrin-labeled endosomes.
Movie 1.
A431 cell expressing GFP-Rab5 after 10 min internalization of rhodamine transferrin. Note that transferrin (red) frequently enters tubules that do not contain Rab5 (green), but seem to be connected with the more globular Rab5 structures. In the center, a segregation event of Rab5 from a larger transferrin-filled endosome can be observed.
Movie 2.
A431 cell expressing GFP-Rab4 after 10 min internalization of rhodamine transferrin followed by a 5-min chase. Note the tubules and small vesicles extending from larger structures that contain both Rab4 (green) and transferrin (red).
Movie 3.
A431 cell expressing GFP-Rab11 after 10 min internalization of rhodamine transferrin followed by a 30-min chase. Transferrin (red) and Rab11 (green) containing structures have accumulated in the pericentriolar area. Note though, that also on transferrin-labeled structures located more in the periphery Rab11 occupies vesicular and tubular subdomains.
Movies 4–6.
Rab4, Rab5, and Rab11 occupy subdomains that can be located on the same endosome.
Movie 4.
A431 cell coexpressing YFP-Rab4 and CFP-Rab5. Three populations of membranes can be observed: two, which have either only Rab4 (green) or only Rab5 (red), and a third containing both, as shown by the yellow color in the overlapping regions. Note that Rab4 and Rab5 do not completely overlap; in contrast, they are often segregated into different domains on these structures.
Movie 5.
In A431 cells coexpressing CFP-Rab5 and YFP-Rab11 little overlap is observed between the small vesicular and tubular Rab11 structures (green) and the bigger globular Rab5 endosomes (red). However, both kinds of membranes are often located in close proximity.
Movie 6.
In A431 cells coexpressing YFP-Rab4 and CFP-Rab11 a significant population of endosomes contains both Rab4 (red) and Rab11 (green). However, again little yellow overlap is observed, both proteins occupy mostly separate domains on the same membrane (compare video 4).
Movies 7–9.
Differential response of Rab4/5 and Rab4/11 endosomes to treatment with brefeldin A.
Movie 7.
A431 cell expressing GFP-Rab4 (green) after internalization of rhodamine transferrin (red) for 10 min followed by incubation with 5 µg/ml BFA for 10 min. Note the dynamics of the extensive tubular network containing both Rab4 and transferrin, but also the remaining globular endosomes that seem to have transient interactions with the network.
Movie 8.
In A431 cells coexpressing YFP-Rab4 and CFP-Rab11 both proteins are found in the tubular network generated after 10 min of BFA treatment. Note though, that a significant population of Rab4 (red) remains in globular endosomes that do not contain Rab11.
Movie 9.
A431 cells coexpressing YFP-Rab4 and CFP-Rab5 demonstrate that endosomes containing Rab5 (red) do not enter the tubular network caused by treatment with BFA. Rab4 (green) can be found on Rab5 structures and on tubules that seem to form transient connections with the globular Rab4/Rab5 endosomes.
Movie 10.
A431 cell coexpressing YFP-Rab4 and CFP-Rab5. Rab5 endosomes (red) tubulate in response to treatment with 100 nM wortmannin for 30 min.
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Transferrin Travels through Rab5-, Rab4-, and Rab11-labeled Domains on Endosomes
We then followed the passage of transferrin through endosomes with respect to the machinery represented by Rab4, Rab5, and Rab11. In cells expressing GFP-Rab4, Rab5, or Rab11, rhodamine transferrin was internalized for 2 min at 37°C and chased in the presence of nonlabeled transferrin (Fig. 2 A). After fixation, images were acquired and overlap of fluorescent signal was quantified (see Materials and Methods). Early after internalization, the major population of transferrin-labeled structures (70%) colabeled with Rab5 (Fig. 2a and Fig. b). This association was transient and decreased rapidly after the first minutes. The number of transferrin-labeled endosomes containing Rab11 increased slowly from 20 to 55% during a 30-min chase (Fig. 2a and Fig. b), in line with an involvement of Rab11 in late stages of recycling (Ullrich et al. 1996; Ren et al. 1998). Rab4 has been implicated in the fast route of recycling (Van der Sluijs et al. 1992; Sheff et al. 1999). Consistently, >90% of transferrin-labeled membranes contained Rab4 after a 5-min chase (Fig. 2a and Fig. b). Surprisingly, 70% remained positive for Rab4 at late time points (15 and 30 min after internalization) when >80% of the transferrin had been recycled (compare with Fig. 1 G), suggesting that a requirement for Rab4 might not be limited to early stages.
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Rab4, Rab5, and Rab11 Label Distinct Domains on the Same Endosomes
We next asked whether Rab4, Rab5, and Rab11 were differentially localized to domains of the same endosome. In other words, were Rab5 negative tubules that emerged from a transferrin-labeled endosome occupied by Rab4 or Rab11? To address this question, combinations of CFP and YFP fusions of the three Rab proteins were coexpressed in A431 cells. Texas red transferrin was internalized for 30 min to reach steady state labeling. After fixation, confocal serial sections were obtained and processed with a deconvolution software before quantification of overlapping fluorescent signals. Single endosomes were defined as structures with continuous transferrin signal (see Materials and Methods).
Interestingly, although residing on the same structure, as defined by the continuous labeling for transferrin, the fluorescent signals for the individual Rab proteins were mostly segregated (Fig. 3, insets). This suggested that Rab4, Rab5, and Rab11 could indeed occupy distinct domains on continuous membranes. Transferrin traverses through Rab5-positive endosomes early after internalization (Fig. 2 A). Consistently, at steady state the cargo was mainly found in Rab4 and Rab11 containing membranes (Fig. 3 and Table ). Remarkably, the majority (63 ± 5%) of these transferrin structures harbored both Rab4 and Rab11. In contrast, only about a quarter of the transferrin positive endosomes contained Rab4 and Rab5 (23.5 ± 7%, Fig. 3 and Table ). The lowest overlap was seen for Rab5 and Rab11 (19 ± 8%). The time course suggested that after leaving Rab5 endosomes, transferrin rapidly reached Rab4-positive membranes (Fig. 2 A). We therefore speculated that this transit might occur within the same endosomes containing both Rab4 and Rab5. Consistently, when we analyzed cells shortly after internalization, more than twice as many transferrin-filled endosomes contained Rab4 and Rab5, compared with the steady state situation (51 ± 8% vs. 23.5 ± 7%, Table ). This suggests that the cargo could be sorted from a Rab5 into a Rab4 domain, residing on the same membrane.
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Electron Microscopy Confirms that Rab Proteins Localize to Distinct Membrane Domains on the Same Endosome
The data presented so far support our model for a domain organization of endosomal membranes. However, the size of the observed structures is at the limit of resolution for light microscopy techniques. We therefore sought to substantiate these findings by electron microscopy. To have the entire surface area available for immunolabeling, we decided to avoid sectioning of the membranes. Cells expressing GFP-Rab4 (Fig. 4A and Fig. B) or GFP-Rab5 (Fig. 4 D) were incubated with 5-nm gold-conjugated to BSA for 30 min at 37°C. Membranes from postnuclear supernatants of these cells were adsorbed to copper grids by incubation at room temperature in the presence of an ATP-regenerating system and GTP. The material was labeled with antibodies to GFP and EEA1 or Rab11.
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Consistent with this observation, transit of transferrin from Rab4/Rab5-positive endosomes to the Rab4/Rab11 network proceeded when cells were pretreated with BFA to allow tubule formation. Rhodamine transferrin, internalized for 2 min at 37°C, was followed by a chase in the presence of BFA (Fig. 6). The amount of Rab5-positive transferrin structures was slightly higher than in control cells at early time points (Fig. 6 A), however, we found no significant morphological difference to nontreated cells (Fig. 6 B, Rab5 panel, compare with Fig. 2 B). Transfer of transferrin into Rab4-labeled endosomes proceeded faster in the presence of BFA (Fig. 6 A, 2 min), but at early stages the cargo remained restricted to globular Rab4-positive endosomes and was largely excluded from the tubular network (Fig. 6 B). After 5 min, transferrin had left Rab5 positive and reached Rab4-labeled membranes to similar levels as seen in control cells (Fig. 6 A). At this point some cargo had entered the BFA-induced tubular network (Fig. 6 B). This was accompanied by a slight increase for Rab11 labeling compared with nontreated cells, consistent with the observation that Rab4 and Rab11 were not clearly segregated in these tubules. At later time points most of the transferrin was found in a membrane network that did not permit quantitation of individual structures (not shown).
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Wortmannin Treatment Delays Exit of Transferrin from Rab4/Rab5-Positive Endosomes
Cells treated with the phosphatidylinositol-kinase inhibitor wortmannin show a delay in transferrin recycling (Spiro et al. 1996). The activity of the PI-(3)-kinase hVPS34 is necessary for the membrane recruitment of the Rab5 effector EEA1 (Mills et al. 1998; Simonsen et al. 1998; Christoforidis, et al. 1999b), suggesting that a Rab5 domain requires a microenvironment of PI(3)P and active Rab5 with its effectors on the membrane. Therefore, depletion of PI(3)P by wortmannin treatment should perturb the Rab5 domain. To test whether this might interfere with the transfer of transferrin to Rab4-positive domains, we quantitated the codistribution of Rab4, Rab5, and Rab11 with transferrin after 30 min internalization in the presence of 50 nm wortmannin (Table ). Consistent with our hypothesis, we found that compared with control cells twice as many transferrin-labeled endosomes contained Rab4 and Rab5 (47 ± 14% vs. 23 ± 7%, Table ). Likewise, the number of cargo-filled structures containing Rab4 and Rab11 was reduced to almost half compared with control cells (35 ± 6% vs. 63.5 ± 5%, Table ), indicating that transferrin was retained in Rab4/Rab5 endosomes upon wortmannin treatment. Moreover, also the domain arrangement was changed as judged by the number of Rab4-positive endosomes containing either Rab5 or Rab11. More Rab4 endosomes were positive for Rab5 (65.5 ± 4% vs. 52 ± 6%) and fewer labeled for Rab11 (34.5 ± 9% vs. 50 ± 1%, Table ). Sorting of membrane associated cargo has been suggested to occur by segregation from globular into tubular domains of the endosome (Mayor et al. 1993). The observed delay of transferrin recycling in wortmannin-treated cells might be explained by the perturbation of this domain structure. Consistently, wortmannin treatment frequently resulted in structural changes of Rab5 endosomes (video 10), which were not observed on Rab11 membranes.
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Discussion |
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How then is efficient receptor recycling achieved? By mathematical modeling the transferrin route is best described assuming two recycling stations with different kinetic properties (Sheff et al. 1999). The first is filled maximally within 5 min, subsequently more than two-thirds of the remaining transferrin will accumulate in the second station after 25 min. Our data would imply a role for Rab4 on both of these sorting stations. 90% of the transferrin was associated with Rab4-positive endosomes at early time points, and 64% of the transferrin was found in Rab4 and Rab11-positive endosomes after 30 min of internalization. Fast recycling seems to require a rapid sorting of transferrin from a Rab5 into a Rab4 domain on the same endosome. This process is delayed when the Rab5 domain is perturbed by wortmannin treatment. One could envisage that neighboring domains would communicate via common effector proteins. One such candidate protein would be rabaptin5 (Vitale et al. 1998), which binds both Rab4 and Rab5. Once transferrin has traversed Rab5/Rab4 endosomes, it would accumulate in membranes enriched for Rab4 and Rab11 domains. These membranes would exert slower kinetics of recycling and exhibit different sorting capacity. Whereas the role for Rab5 in endosome fusion is well established, the exact functions of Rab4 and Rab11 are unknown. These two GTPases might be directly involved in the formation of recycling vesicles, possibly targeted to different sites of the plasma membrane. For example, Rab11 might have a role in polarized transport in epithelial or motile cells (e.g., facilitate transcytosis of cargo like the polymeric IgA receptor; Sheff et al. 1999), whereas Rab4 would regulate a more general route of membrane recycling. The nature of the recycling vesicles is unclear. Coated buds have been observed on tubulovesicular endosomes (Stoorvogel et al. 1996), and the tubulation of Rab4/Rab11 membranes upon BFA treatment is most likely due to the inhibition of ARF-dependent processes (Donaldson et al. 1992; Helms and Rothman 1992). However, whether coated vesicles actively participate in recycling remains to be established. If sorting is completed once transferrin has reached a Rab4-positive domain, coat assembly might not be absolutely required. Alternatively, a BFA-insensitive exchange factor for ARF might exist on Rab4/Rab5 endosomes. Either mechanism would explain why recycling is only marginally effected by BFA.
In summary, envisaging endosomes as dynamic arrangements of membrane domains can give an integrative view on the complexity and robustness of the recycling pathway. This model not only agrees with the observed kinetics (Sheff et al. 1999), but also explains how in some cell types a seemingly continuous membrane system can achieve compartmentalization (Hopkins et al. 1990). However, at the molecular level it is mainly based on the interactions of a single Rab protein, Rab5, with its effectors. Besides the morphological evidence reported here, to show that these domains generally serve as a platform for membrane organization, it will be necessary to characterize Rab4 and Rab11 effector proteins. This is now the focus of our attention.
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Acknowledgments |
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B. Sönnichsen and E. Nielsen were supported by long term European Molecular Biology Organization fellowships. This work was supported by the Max Planck Society and grants from the Human Frontier Science Program (RG-432/96), European Union Training and Mobility of Researchers (ERB-CT96-0020), and Biomed (BMH4-97-2410; M. Zerial).
Submitted: 17 February 2000
Revised: 28 March 2000
Accepted: 10 April 2000
The online version of this paper contains supplemental material.
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References |
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