Article |
Address correspondence to Margaret S. Robinson, University of Cambridge, CIMR, Wellcome Trust/MRC Building, Addenbrooke's Hospital, Hills Road, Cambridge CB2 2XY, UK. Tel.: 44-1223-330163. Fax: 44-1223-762640. email: msr12{at}mole.bio.cam.ac.uk
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Abstract |
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Key Words: coated vesicles; adaptors; RNA interference; receptor-mediated endocytosis; internalization signals
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Introduction |
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Although most of the available data are consistent with this model, there are a few discrepancies. First, the budding yeast Saccharomyces cerevisiae has an AP-2related complex, AP-2R; however, deleting genes encoding AP-2R subunits has no apparent effect on clathrin-mediated endocytosis, or indeed, on any other pathway that has been investigated (Yeung et al., 1999). A caveat here is that AP-2R may not be functionally equivalent to AP-2 in higher cells. Second, some of the accessory proteins that bind to AP-2 have properties suggesting that they may be adaptors in their own right. Epsin, ß-arrestin, AP180/CALM, Hip1, Dab2, and ARH have all been shown to bind not only to AP-2, but also to PIP2 and to clathrin, indicating that they may be able to bind to the plasma membrane and to recruit clathrin in an AP-2independent manner (Gaidarov et al., 1999; Ford et al., 2001, 2002; Mishra et al., 2001, 2002a,b). There is also some evidence that each of these proteins may interact with a specific type of cargo. For instance, epsin has ubiquitin-interacting motifs that may help to facilitate the internalization of plasma membrane proteins like the EGF receptor, which is ubiquitinated in response to ligand binding. Dab2 and ARH are able to bind NPXY motifs, found in members of the LDL receptor family (for review see Bonifacino and Traub, 2003). Thus, an alternative view is that AP-2 may be just one of several endocytic adaptors, and that although it participates in the network of proteinprotein interactions and is required for the uptake of certain types of cargo, it is not essential for clathrin-mediated endocytosis.
We asked the question, what would happen if AP-2 were to drop out of the network? Would clathrin and accessory proteins still be recruited onto the plasma membrane? What would happen to receptor-mediated endocytosis? Using small interfering RNAs (siRNAs), we have been able to knock down the expression of the µ2 subunit of the AP-2 complex to undetectable levels in HeLaM cells. Then, we compared the phenotype of the AP-2depleted cells with that of cells treated with siRNAs directed against clathrin heavy chain.
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Results |
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Both AP-2 and clathrin depletion were found to slow down the growth of the cells, but we did not see any apparent increase in apoptosis (unpublished data), although clathrin loss has been reported to cause apoptosis in a chicken B cell line (Wettey et al., 2002). By phase-contrast microscopy, the µ2-2treated cells look essentially normal (compare Fig. 1 e with the control cells in Fig. 1 d). However, many of the chc-2treated cells were found to be heavily vacuolated, and nearly half of the cells had two or more nuclei (Fig. 1 f), indicating that cytokinesis is blocked in clathrin-depleted cells. Interestingly, a similar phenotype has been reported in clathrin-deficient Dictyostelium (Niswonger and O'Halloran, 1997).
Fig. 1 (gn) shows the appearance of the cells by immunofluorescence. In gk, either control cells (g) or AP-2 knockdown cells (hk) were labeled with an antibody against the AP-2 subunit. After a single transfection, the
-adaptin labeling was found to be very patchy in both
-2treated (h) and µ2-2treated (i) cells; individual spots were of approximately equal intensity to those in control cells, but much fewer in number and usually occurring in clusters. After two transfections (j and k), membrane-associated labeling was essentially undetectable in the µ2-2treated cells, although cytosolic labeling of partial complexes containing the
subunit could be seen (k). The
-2 knockdown (j) was not quite so complete in that some of the cells still had residual spots after the second transfection. Clathrin knockdowns (ln) caused a more uniform loss of signal. After a single transfection (m), the number of clathrin-positive spots per cell was similar to controls (l), but the intensity of the spots was reduced. After two transfections, clathrin labeling was essentially undetectable (n). In all three cases, virtually 100% of the cells were affected by the end of the full course of treatment.
Localization of other proteins in the siRNA-treated cells
To find out what happens to other coat proteins in cells where one of them has been depleted, we performed triple labeling on cells treated with both µ2-2 and chc-2 siRNAs. For these experiments, we plated out equal numbers of control and siRNA-treated cells onto microscope slides so the two types of cells could be viewed together in the same field, and then labeled the cells with antibodies against -adaptin (blue), clathrin heavy chain (red), and epsin 1 (green).
Fig. 2 shows that normally, the three proteins have overlapping distributions at the plasma membrane, with additional intracellular structures labeled with the clathrin antibody. Knocking down µ2 causes a reduction in the number of epsin spots (c). These spots do not contain any detectable -adaptin, which is now diffuse and cytosolic (a). However, many of the epsin-positive spots in these cells are also positive for clathrin (b; insets), suggesting that the two proteins are able to co-assemble on the plasma membrane even in the absence of AP-2.
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Ultrastructure of the cells
To observe the phenotype of AP-2 and clathrin-depleted cells at the ultrastructural level, EM was performed. Fig. 3 (a and b) shows that control cells contain numerous clathrin-coated pits associated with the plasma membrane, indicated with the large arrowheads. A morphometric analysis (Fig. 4) showed that in these cells, 0.6% of the cell surface is occupied by clathrin-coated pits. In the AP-2depleted cells (ce), clathrin-coated pits were found to be 12-fold less abundant, occupying only 0.05% of the cell surface. The morphology of the coat appears to be identical in control and AP-2depleted cells; however, the coated pits tend to be smaller in the AP-2depleted cells. In the clathrin-depleted cells, clathrin-coated pits were undetectable, and nearly all of the budding profiles that could be observed at the plasma membrane had the characteristic appearance of caveolae.
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Effects on clathrin-mediated endocytosis: transferrin receptor
It has been well documented that AP-2 is required for the uptake of cargo proteins with YXX-type sorting signals, such as the transferrin receptor. This motif has been shown to bind to hydrophobic pockets in the µ2 subunit (Ohno et al., 1995; Owen and Evans, 1998). Mutating the signal in the transferrin receptor prevents the receptor from entering the cell by clathrin-mediated endocytosis, causing it to be internalized 510 times more slowly (Jing et al., 1990). Similarly, mutating the µ2 subunit so that it can no longer bind the motif, and then overexpressing it in cells so that most of the AP-2 complexes contain mutant rather than wild-type µ2, also strongly inhibits uptake of the transferrin receptor (Nesterov et al., 1999).
Fig. 5 a shows the effects of AP-2 and clathrin knockdown on transferrin receptor endocytosis. For these experiments, the cells were incubated at 4°C with 125I-labeled transferrin to allow binding (but not internalization) to occur, and then were warmed to 37°C for various lengths of time. At the end of the incubation, the medium was harvested, surface-bound ligand was stripped off at low pH, the cells were solubilized in 1 M NaOH, and the label in all three fractions was quantified. The percentage of counts in the NaOH extract (i.e., intracellular counts) is shown in the graph. In control cells, >50% of the prebound transferrin is internalized within 5 min after warm-up, but after 10 min, the amount of intracellular transferrin starts to go down, as it gets recycled back into the medium. Knocking down both AP-2 and clathrin causes a profound inhibition of transferrin uptake. In both cases, the transferrin never accumulates inside the cells, but remains primarily on the cell surface, where it slowly dissociates into the medium. Even after 30 min, <10% of the transferrin is recovered in the intracellular fraction. Thus, as expected, both AP-2 and clathrin are required for efficient internalization of the transferrin receptor.
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Fig. 5 b shows a similar experiment to the one shown in Fig. 5 a, but using 125I-labeled EGF as the ligand. In control cells, EGF is internalized slightly more slowly than transferrin, but by 10 min, >50% of the label is intracellular. Knocking down clathrin strongly inhibits the receptor-mediated endocytosis of EGF. Strikingly, however, knocking down AP-2 has no significant effect on EGF uptake. The rate of accumulation of intracellular EGF is virtually identical in control and AP-2depleted cells.
LDL receptor
There is currently some controversy as to how the LDL receptor's internalization signal actually works. This signal has been reported to bind to several coat components, including the µ2 subunit of AP-2 (Boll et al., 2002), clathrin (Kibbey et al., 1998), and proteins with phosphotyrosine-binding domains like Dab2 and ARH (Mishra et al., 2002a,b). Initially, we attempted to monitor receptor internalization using 125I-labeled LDL, but these experiments were complicated by the very rapid dissociation of the ligand from its receptor in HeLa cells (see Materials and methods). Therefore, we stably transfected the cells with a construct consisting of the extracellular and transmembrane domains of CD8 fused to the LDL receptor tail, then assayed for internalization following a similar protocol to the one just mentioned, but with anti-CD8 followed by 125I-labeled protein A as the ligand. Fig. 5 c shows that in control cells, the LDL receptor chimera is efficiently internalized, and that knocking down clathrin strongly inhibits uptake. However, knocking down AP-2 again has no effect on the rate of endocytosis of the protein. Thus, out of the three transmembrane proteins that we have analyzed in siRNA-treated HeLa cells, two are internalized in an AP-2independent manner.
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Discussion |
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Over the last several years, a number of reports have been published that challenge the view that all cargo proteins are endocytosed by the same molecular mechanism. Marks et al. (1996) showed that overexpression of constructs with either YXX or dileucine internalization signals causes endogenous proteins with the same signal to accumulate on the cell surface, but endogenous proteins with the other signal are still endocytosed normally. They concluded that the two signals were competing for two distinct components of the endocytic machinery. However, they could not rule out the possibility that both of these components were part of the AP-2 complex, and indeed, subsequent reports showed that dileucine signals can also bind to AP complexes, most likely through their ß subunits (for review see Bonifacino and Traub, 2003). Warren et al. (1998) used a similar approach to show that the transferrin, EGF, and LDL receptors do not compete with each other for internalization. They suggested that although AP-2 interactions might be necessary for the internalization of some cargo proteins, other cargo proteins could be interacting with other components of the coat. However, until now, it has not been possible to test these ideas by specifically depleting different coat components from the cell.
The approach that we have used in the present paper should be widely applicable for looking at the internalization of other transmembrane proteins, to determine whether or not their endocytosis is dependent on AP-2. So far, most functional analyses of the roles of various proteins in clathrin-mediated endocytosis have made use of dominant-negative mutants (for review see Conner and Schmid, 2003a), but these may lead to indirect effects (e.g., overexpressing a truncated version of a particular protein may impede interactions involving other components of the coated pit). RNA interference is a much cleaner way of testing the function of a protein because it removes the protein instead of altering it and adding it back in excess amounts. The CD8 chimera system may be especially useful, because theoretically, one could transplant the cytoplasmic tail from any type I membrane protein, or an artificial tail designed to test a potential internalization signal, onto the CD8 reporter, and then assay for uptake using reagents that are commercially available.
Fig. 6 is a schematic diagram summarizing our results and how we interpret them. Normally, clathrin, AP-2, and alternative adaptors all co-assemble at the plasma membrane, bringing cargo with different types of internalization signals into the coated pit. In the absence of AP-2, alternative adaptors are still recruited onto the plasma membrane, where they interact with a subset of the cargo proteins, including the EGF and LDL receptors, and co-assemble with clathrin. These cargo proteins are still efficiently internalized, but cargo proteins like the transferrin receptor, which can only interact with AP-2, stay on the cell surface. In addition, because AP-2 is the major clathrin adaptor at the plasma membrane, there is less clathrin recruited onto the membrane and fewer (and perhaps smaller) coated pits per cell. In the absence of clathrin, both AP-2 and alternative adaptors are still recruited onto the plasma membrane, where they interact with each other and most likely with potential cargo proteins as well, but there are no morphologically recognizable coated pits, and all of the cargo proteins are internalized much more slowly.
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Materials and methods |
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Two independent siRNAs were used to investigate the effects of knockdown of each target. For AP-2, the targets were the and µ2 subunits of the complex. The
-2 siRNA target sequence was AAGAGCAUGUGCACGCUGGCCA and the µ2-2 target sequence was AAGUGGAUGCCUUUCGGGUCA (other sequences, designated
-1 and µ21, were ineffective). The clathrin heavy chain target sequences were AAG-CUGGGAAAACUCUUCAGA (chc-1) and UAAUCCAAUUCGAAGACCAAU (chc-2). The control siRNA was a nonfunctional oligo, µ21, originally designed to knock down the µ2 subunit, target sequence AACACAGCAACCUCUACUUGG. All siRNAs were designed according to the manufacturer's instructions and were synthesized as Option C siRNAs by Dharmacon, Inc.
Immunofluorescence and Western blotting
Antibodies used for immunofluorescence included an affinity-purified rabbit polyclonal antiserum directed against clathrin heavy chain (Simpson et al., 1996); a mouse monoclonal anti--adaptin, AP.6, provided by Frances Brodsky (University of California, San Francisco, San Francisco, CA); and a commercially available goat anti-epsin 1 (Santa Cruz Biotechnology, Inc.). Secondary antibodies were all purchased from Molecular Probes, Inc. For Western blotting,
and µ2 subunits of AP-2 were detected using mouse monoclonal antibodies from Transduction Laboratories, clathrin heavy chain was detected using the same rabbit antibody that was used for immunofluorescence, and actin was detected using a rabbit antibody from Sigma-Aldrich. Incubations with the mouse antibodies were followed by an incubation with rabbit antimouse immunoglobulin (DakoCytomation). The blots were then probed with 125I-labeled protein A (Amersham Biosciences).
EM
Cells to be used for EM were first fed with mouse anti-transferrin receptor B3/25 conjugated to 8-nm gold, a gift from Colin Hopkins and Liz Alichin (Imperial College London, London, UK). The cells were incubated with the antibodygold conjugate for 30 min at 4°C, followed by a 10-min chase at 37°C. They were then fixed in tissue culture dishes by adding double-strength fixative (5% glutaraldehyde and 4% PFA in 0.2 M sodium cacodylate, pH 7.2, containing 6 mM CaCl2) to an equal volume of tissue culture medium for 2 min at 37°C. This solution was aspirated off and replaced by 2.5% glutaraldehyde and 2% PFA in 0.1 M sodium cacodylate buffer, pH 7.2, containing 3 mM CaCl2. The cells were fixed for a further 3 h at RT, washed with 0.1 M sodium cacodylate buffer, pH 7.2, and post-fixed with 1% osmium tetroxide in 0.1 M sodium cacodylate buffer, pH 7.2, for 1 h. The cells were scraped from the dish and pelleted, and the pellet was washed with 0.05 M sodium maleate buffer, pH 5.2, and en bloc stained with 0.5% uranyl acetate in 0.05 M sodium maleate buffer. The cell pellets were dehydrated in ethanol, exchanged into 1,2-epoxy propane, and embedded in Araldite CY212 epoxy resin (Agar Scientific).
50-nm ultrathin sections were cut using a diamond knife mounted on an ultramicrotome (Reichert Ultracut S; Leica), collected onto formvar/carbon-coated EM grids, and stained with uranyl acetate and Reynolds lead citrate (Reynolds, 1963). The sections were observed in a transmission electron microscope (model CM 100; Philips) at an operating voltage of 80 kV.
For quantification of the percentage of plasma membrane occupied by coated pits, cell pellets were randomly sectioned and oriented in the electron microscope. Grids were systematically scanned at a magnification of 13,500, and 128 images of each condition were captured using a camera (Megaview II TEM; Soft Imaging System). A 500-nm lattice overlay was used to score intersections with the plasma membrane and coated pits (Weibel, 1979; Griffiths, 1993) using analySIS® 3.0 image analytical software.
Internalization assays
Ligand uptake assays were performed on cells seeded onto 35-mm dishes on the day preceding the experiment. On the day of the experiment, the dishes were placed on ice and were washed briefly with prechilled serum-free medium (SFM; DME and 20 mM Hepes containing 1% BSA). The cells were then incubated on a rocker for 30 min at 4°C in 0.6 ml SFM containing either 125I-labeled EGF or 125I-labeled transferrin (both obtained from PerkinElmer) at a concentration of 500 nCi/ml. The dishes were washed six times with ice-cold SFM, and the surface counts were collected for the zero time point by incubating the cells for 5 min at 4°C in 0.8 ml ice-cold acid wash (0.2 M acetic acid and 0.5 M NaCl), followed by a brief rinse with another 0.8-ml acid wash. The other dishes were incubated with 1.5 ml prewarmed medium (DME containing 10% FCS) at 37°C for various times. Endocytosis was stopped by placing the cells on ice, collecting the medium, and rinsing the cells with ice-cold SFM. Label associated with the cell surface was then collected by acid wash, as above. Finally, the intracellular fraction was collected by extracting the cells twice with 0.8 ml 1 M NaOH. The radioactivity in the medium, acid wash, and NaOH extract was quantified using a gamma counter (Nuclear Enterprises).
We also attempted to follow the fate of 125I-labeled LDL, but found that >50% of the prebound ligand dissociated within the first few minutes of warm-up. Therefore, we made use of HeLaM cells expressing a chimera of the extracellular and transmembrane domain of CD8 fused to the tail domain of the LDL receptor. Human CD8 cDNA in pBlueScript® was a gift from Gudrun Ihrke (University of Cambridge, Cambridge, UK; Ihrke et al., 2001). The cytoplasmic tail of the mouse LDL receptor (from the arginine residue at position 813) was amplified by PCR from an EST (Clone ID 2581960; GenBank/EMBL/DDBJ accession no. gi6519196) obtained from the I.M.A.G.E. Consortium, incorporating an AflII site into the 5' end. The resulting fragment was ligated to the AflII site at the end of the transmembrane domain coding sequence of CD8, and the chimera was cloned into pIRES2Neo (CLONTECH Laboratories, Inc.). The construct was sequenced to confirm that a correct in-frame fusion had been achieved, and was then transfected into HeLaM cells. Stably transfected cells were selected and maintained in the presence of 500 µg/ml G418 (GIBCO BRL).
To monitor uptake of the chimera, the cells were treated as above, but incubated with SFM containing 1:100 diluted anti-CD8 (153020; Ancell Corp.) instead of with a radiolabeled ligand, and the incubation was for 45 min at 4°C instead of for 30 min. The cells were then washed and incubated for a further 45 min at 4°C with SFM containing 125I-labeled protein A (1:1,000 diluted; Amersham Biosciences). The rest of the experiment was performed exactly as above.
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Acknowledgments |
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This work was supported by grants from the Wellcome Trust (grant 053316/Z/98 to M.S. Robinson) and the Medical Research Council (grant G9310915 to M.S. Robinson and J.P. Luzio).
Submitted: 30 May 2003
Accepted: 24 July 2003
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