Phosphatidylinositol 4,5-Bisphosphate Regulates Adipocyte Actin ...

THE JOURNAL OF BIOLOGICAL CHEMISTRY © 2004 by The American Society for Biochemistry and Molecular Biology, Inc.

Vol. 279, No. 29, Issue of July 16, pp. 30622–30633, 2004 Printed in U.S.A.

Phosphatidylinositol 4,5-Bisphosphate Regulates Adipocyte Actin Dynamics and GLUT4 Vesicle Recycling* Received for publication, February 9, 2004, and in revised form, April 26, 2004 Published, JBC Papers in Press, April 28, 2004, DOI 10.1074/jbc.M401443200

Makoto Kanzaki‡, Megumi Furukawa, William Raab, and Jeffrey E. Pessin From the Department of Pharmacological Sciences, State University of New York at Stony Brook, Stony Brook, New York 11794

To investigate the potential role of phosphatidylinositol 4, 5-bisphosphate (PI(4,5)P2) in the regulation of actin polymerization and GLUT4 translocation, the type I phosphatidylinositol 4-phosphate 5-kinases (PIP5Ks) were expressed in 3T3L1 adipocytes. In preadipocytes (fibroblasts) PIP5K expression promoted actin polymerization on membrane-bound vesicles to form motile actin comets. In contrast, expression of PIP5K in differentiated 3T3L1 adipocytes resulted in the formation of enlarged vacuole-like structures coated with F-actin, cortactin, dynamin, and N-WASP. Treatment with either latrunculin B (an inhibitor for actin polymerization) or Clostridium difficile toxin B (a general Rho family inhibitor) resulted in a relatively slower disappearance of coated F-actin from these vacuoles, but the vacuoles themselves remained unaffected. Functionally, the increased PI(4,5)P2 levels resulted in an inhibition of transferrin receptor and GLUT4 endocytosis and a slow accumulation of these proteins in the PI(4,5)P2-enriched vacuoles along with the non-clathrin-derived endosome marker (caveolin) and the AP-2 adaptor complex. However, these structures were devoid of early endosome markers (EEA1, clathrin) and the biosynthetic membrane secretory machinery markers p115 (Golgi) and syntaxin 6 (trans-Golgi Network). Taken together, these data demonstrate that PI(4,5)P2 has distinct morphologic and functional properties depending upon specific cell context. In adipocytes, altered PI(4,5)P2 metabolism has marked effects on GLUT4 endocytosis and intracellular vesicle trafficking due to the derangement of actin dynamics.

Insulin stimulation of glucose uptake in striated muscle and adipose tissue is induced by translocation of the insulin-responsive glucose transporter isoform (GLUT4) from intracellular storage sites to the plasma membrane (1–3). In the basal state, GLUT4 cycles slowly between the plasma membrane and multiple intracellular compartments, with the most of GLUT4 residing inside the cell. In contrast, insulin binding to its specific receptor evokes a large increase in the rate of GLUT4 vesicle exocytosis, with a small decrease in the rate of its internalization by endo* This work was supported by Research Grants DK33823 and DK59291 from the National Institutes of Health (to J. E. P.) and Grant 1-03-JF-14 from the American Diabetes Association (to M. K.). The costs of publication of this article were defrayed in 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. ‡ To whom correspondence should be addressed: Tohoku University Biomedical Engineering Research Organization (TUBERO), Bldg. 1, Graduate School of Medicine, 2-1 Seiryo-machi, Aoba, Sendai, Miyag 980-8575, Japan. Tel.: 81-22-717-7581; Fax: 81-22-717-7873; E-mail: [email protected].

cytosis (4 – 6). Thus, the enhanced glucose uptake in response to insulin is achieved by a net increase of GLUT4 protein levels on the cell surface resulting from the insulin-induced shift in the cellular dynamics of GLUT4 vesicle trafficking. Recently multiple studies have begun to dissect the molecular machinery involved in the trafficking of GLUT4 vesicles, and it is increasingly apparent that both the microtubule (7– 12) and actin cytoskeleton (13–18) play an indispensable role in this process. In particular, the insulin-induced GLUT4 translocation in adipocytes requires dynamic actin remodeling at the inner surface of the plasma membrane (cortical actin) and in the perinuclear region (13). The dynamic actin rearrangement required for GLUT4 translocation is regulated by at least two distinct insulin receptor-mediated signals, one leading to the activation of phosphatidylinositol (PI)1 3-kinase (19 –22) and the other to the activation of Rho family small GTP-binding protein TC10 (13, 23, 24). Although the precise molecular mechanisms remain uncertain, multiple mechanisms have been proposed for the function of actin in the control of vesicle trafficking events (25–28). For example, several studies have demonstrated that proper actin regulation is required for endocytosis (29), vesicular fusion (30, 31), trans-Golgi network (TGN) exit (32, 33), spatial targeting of secretory proteins, and the maintenance of the Golgi complex structure (34 –36) including the GLUT4 storage compartment (17). Several recent studies have suggested a mechanism linking phosphatidylinositol 4,5-bisphosphate (PI(4,5)P2) with actin polymerization through the function of N-WASP. Interaction of N-WASP with PI(4,5)P2 and activated Rho proteins including Cdc42 (37) and TC10 (24) exposes the VCA domain (verprolin homology, cofilin homology, and acidic region), which in turn activates the Arp2/3 complex, resulting in a burst of de novo actin polymerization in response to extracellular stimuli (38, 39). We have reported recently that insulin stimulates N-WASPdependent formation of actin comet tails on GLUT4-containing endosomes (40) and also that the expression of a dominantinterfering N-WASP lacking the VCA domain in 3T3L1 adipocytes significantly inhibited GLUT4 translocation (14, 40). In fibroblasts, expression of type I phosphatidylinositol 4-phosphate 5-kinase (PIP5K) to generate PI(4,5)P2 results in actin comet tails that can propel vesicles from donor to acceptor compartments through the cytoplasm in an N-WASP-depend1 The abbreviations used are: PI, phosphatidylinositol; AP-2, adaptor protein-2; EGFP, enhanced green fluorescence protein; GM1, Il3NeuAcGgOse4Cer; HA, hemagglutinin; PBS, phosphate-buffered saline; PH domain, pleckstrin homology domain; PIP2, phosphatidylinositol bisphosphate; PI(3,4,5)P3, phosphatidylinositol 3,4,5-trisphosphate; PI(4,5)P2, phosphatidylinositol 4,5-bisphosphate; PIP5K, phosphatidylinositol 4-phosphate 5-kinase; PLC, phospholipase C; TfR, transferrin receptor; TGN, trans-Golgi network; VCA domain, verprolin homology, cofilin homology, and acidic region; VSV-G, vesicular stomatitis virus G protein; WT, wild-type.

30622

This paper is available on line at http://www.jbc.org

30623

PIP2 Regulation of GLUT4 Endocytosis ent manner (41, 42). Furthermore, PIP5K can recruit the large GTP-binding protein dynamin to actin comet tails (43– 45). Time-lapse microscopy in living cells suggested that the formation of PIP5K-induced actin comet tails was reflective of the rapid motility of intracellular membranes, including endocytic vesicles from the plasma membrane (44) and the vesicles from the Golgi complex (41), both locations at which dynamin mediates vesicle fission (26, 44). Interestingly, actin comets are reportedly formed preferentially from vesicles that are enriched in lipid rafts containing high levels of cholesterol and sphingolipids, and disruption of lipid rafts reduces the number of actin comet tails (41). Together, these data raise the possibility of a direct linkage among PI(4,5)P2, actin polymerization, and vesicle transport. On the other hand, PI(4,5)P2 also appears to play an important role in the assembly/disassembly of plasma membrane endocytotic coat proteins. In the process of endocytosis, PIP2 serves as a binding site for multiple proteins crucial for the assembly of clathrin-coated vesicles (46 – 48). For example, mutants of both epsin (responsible for recruiting clathrin) and AP-2 (responsible for binding endocytic cargo) that are unable to bind to PIP2 function in a dominant-interfering manner to inhibit endocytosis (49, 50). Similarly, reagents that sequester PIP2 also block clathrin-dependent endocytosis in vitro (46). Furthermore, several other components of the endocytic machinery also interact directly with PIP2, including Eps15, AP-180, dynamin, and clathrin assembly lymphoid myeloid leukemia protein (CALM) (51–54). Thus, the functional role of PI(4,5)P2 in the control of specific types of actin dynamics and vesicle trafficking events appears to be markedly cell context-dependent. To explore relationship among N-WASP-dependent actin polymerization, PI(4,5)P2 metabolism, GLUT4 endocytosis, and translocation, we examined effect of PIP5K expression on actin organization and membrane transport pathways in 3T3L1 adipocytes. Our data demonstrate that increased PI(4,5)P2 production by PIP5K expression in 3T3L1 adipocytes induces intracellular multiple vacuoles coated with F-actin, N-WASP, dynamin, cortactin, caveolin, and the integrin ␤1 receptor subunit consistent with the convergence of fused endosomes from both clathrin- and non-clathrin-derived endocytic vesicles. These data also demonstrate that derangement of PI(4,5)P2 metabolism severely affects the endocytic recycling pathway that consequently results in a persistent GLUT4 localization at the cell surface. EXPERIMENTAL PROCEDURES

Materials—Clostridium difficile toxin B was obtained from Techlab, Inc. (Blacksburg, VA). Latrunculin B was purchased from Calbiochem. Texas Red-conjugated transferrin and fluorescent secondary antibodies were purchased from Molecular Probes. Rhodamine-conjugated cholera toxin B subunit was purchased from List Biological Laboratories, Inc. (Campbell, CA). Rhodamine-conjugated phalloidin was purchased from Sigma. Rabbit polyclonal anticaveolin 1 antibody, mouse monoclonal anti-p115, antisyntaxin 6, anti-EEA1, and anticlathrin heavy chain antibodies were purchased from BD Transduction Laboratories (Lexington, KY). The HA and Myc epitope tag antibodies and the cortactin antibody were purchased from Upstate Biotechnology (Lake Placid, NY). The vesicular stomatitis virus G protein (VSV-G) antibody was from Accurate Chemicals. The anti-human transferrin receptor (TfR) and the anti-integrin ␤1 antibodies were from Zymed Laboratories Inc.(San Francisco) and BD Pharmingen (San Diego), respectively. cDNA Expression Constructs—The mouse type I PIP5K␤ was cloned from 3T3L1 adipocyte cDNA library by PCR, and the kinase-inactive point mutation of PIP5K␤ (PIP5K␤/K307A) was generated by PCR. These cDNAs were then inserted into the pKH3 mammalian expression vector (pKH3-PIP5K␤) containing a HA epitope tag. It is important to note that the nomenclature used throughout this paper is designated for the mouse isoforms (mouse PIP5K␤ is human PIP5K␣) (55, 56). pEGFP-PIP5K␤87kDa, PIP5K␥87kDa/K138A, PIP5K␥90kDa, and

PIP5K␥90kDa/K138A were generous gifts from Dr. Richard Anderson (University of Wisconsin). VSV-G cDNA was obtained from the University of Iowa DNA Core Facility. Human TfR cDNA was purchased from ATCC and was subcloned into pcDNA3 vector (pcDNA3-TfR). pKH3TC10/T31N was prepared as described previously (23). pKH3-dynamin2 and pKH3-dynamin2/K44A were generated by exchanging a BamHI and EcoRI fragment containing the nucleotide sequences encoding the wild-type lysine 44 with a homologous fragment encoding alanine 44 from the mutant dynamin 2 amino acid-EGFP clone as reported previously (57). pEGFP-N-WASP/WT and -N-WASP/⌬VCA were generated by subcloning EcoRI and ApaI fragments from pcDNA3-N-WASP and pcDNA3-N-WASP⌬VCA, respectively (40). pcDNA3-Myc-GLUT4-EGFP, pEGFP-GRP/PH, and pEGFP-PLC␦/PH were prepared as described previously (58). Cell Culture and Transient Transfection of 3T3L1 Adipocytes—Murine 3T3L1 preadipocytes were maintained, differentiated into adipocytes, and transfected by electroporation as described previously (59). After electroporation, cells were plated on glass coverslips and allowed to recover in complete medium. Immunofluorescence and Image Analysis—Transfected and intact adipocytes were washed in phosphate-buffered saline (PBS) and fixed for 20 min in 2% paraformaldehyde and PBS. The cells were washed briefly in PBS, permeabilized in PBS containing 0.1% Triton X-100 for 10 min, and then blocked in 5% donkey serum (Sigma) for 1 h at room temperature. Primary and secondary antibodies were used at 1:100 dilutions (unless otherwise indicated) in 1% bovine serum albumin and PBS, and samples were mounted on glass slides with Vectashield (Vector Laboratories). Cells were imaged using a Zeiss LSM510 confocal fluorescence microscope. Images were then imported into Adobe Photoshop (Adobe Systems, Inc.) for processing. Transferrin Receptor Endocytosis—3T3L1 adipocytes expressing human TfR with empty vector, PIP5K␥87kDa, or its K138A mutant were chilled on ice and incubated with 5 ␮g/ml Texas Red-conjugated transferrin for 1 h at 4 °C to label the surface TfR. Some cells were returned to 37 °C and incubated for the indicated period to allow the endocytosis of labeled TfR. The cells were washed once with ice-cold PBS and fixed in 2% paraformaldehyde and PBS at room temperature for 10 min, and TfR endocytosis was assessed by confocal laser scanning microscopy. GLUT4 Endocytosis Assay—3T3L1 adipocytes expressing exofacial Myc-tagged GLUT4-EGFP with empty vector, PIP5K␤, or the D307A mutant were insulin stimulated for 30 min. Then the cells were chilled and incubated with c-Myc antibody (9E10) for 1 h at 4 °C, and the cells were washed to remove insulin and excess c-Myc antibody as described previously (11). Some cells were returned to 37 °C and incubated for various times to allow the Myc antibody-bound Myc-GLUT4-EGFP to internalize. Then the cells were washed once with ice-cold PBS and fixed in 2% paraformaldehyde at room temperature for 10 min, permeabilized, and blocked with 5% donkey serum. The cells were incubated with Texas Red-conjugated donkey anti-mouse IgG antibody for 2 h at room temperature, mounted in Vectashield, and examined by confocal laser scanning microscopy. Cholesterol Loading and Extraction—To examine the effect of cholesterol, cholesterol-methyl-␤-cyclodextrin complexes were synthesized as described previously (60). Briefly, 9.5 mg of cholesterol was dissolved in 126 ␮l of isopropyl alcohol:CHCl3 (2:1) solution. Methyl-␤-cyclodextrin (315 mg) was dissolved in 3.46 ml of water and heated to 80 °C with stirring. The cholesterol was added to methyl-␤-cyclodextrin, and the solution was stirred until clear. This solution contained 6.8 mM cholesterol. For use, complexes were diluted into medium to a final concentration of 0.1 mM. For cholesterol removal experiments, 18 h after electroporation PIP5K-expressing cells were treated with 10 mM methyl-␤-cyclodextrin for 30 min. RESULTS

Increased Expression of PIP5K Induces Actin Comet Tails in 3T3L1 Fibroblasts but Results in F-actin-coated Intracellular Vacuoles in 3T3L1 Adipocytes—To examine the role of PI(4,5)P2 in the GLUT4 translocation process, we initially expressed the type I PIP5Ks in both 3T3L1 fibroblasts and adipocytes. In 3T3L1 fibroblasts (preadipocytes), overexpression of PIP5K␥87kDa resulted in a formation of several cytoplasmic actin comet tails (Fig. 1, a– d). The PI(4,5)P2 formation of actin comet tails observed in our 3T3L1 fibroblasts was similar to that observed previously in several other fibroblast cell lines (41, 42, 44, 45). Although it has been reported that about 50% of PIP5K-induced actin comet tails were formed from PIP5K-

30624

PIP2 Regulation of GLUT4 Endocytosis

FIG. 1. Increased expression of type I PIP5K induces actin comet tails in 3T3L1 preadipocytes but induces intracellular aggregated vacuoles in differentiated 3T3L1 adipocytes. 3T3L1 preadipocytes (fibroblasts) (A) or differentiated 3T3L1 adipocytes (B) were electroporated with cDNA encoding EGFP-tagged PIP5K␥87kDa. A, 18 h later, the cells were fixed and subjected to confocal microscopy for EGFP (a) and rhodamine-phalloidin (b). The merged image and its magnified image are shown in c and d, respectively. Arrowheads indicate actin comet tails (d). B, 18 h later, the cells were fixed and subjected to confocal microscopy for EGFP and rhodamine-phalloidin. Side views (x and y axis for a and b, respectively) were reconstituted by 20 z section confocal images. This is a representative image from three independent experiments.

positive vesicles (41), we seldom observed PIP5K accumulation at the head of actin comet tails (Fig. 1d). This suggests that an increased level of PI(4,5)P2 concentration on the vesicles themselves, rather than PIP5K protein itself, is sufficient for the formation of actin comet tails. In contrast to the preadipocytes, expression of PIP5K (90-kDa and 87-kDa isoforms) in 3T3L1 adipocytes resulted in a formation of multiple aggregated intracellular vacuoles that were coated with F-actin (Fig. 2A, a– d and i–l). Three-dimensional reconstruction of confocal images demonstrated that these PIP5K-induced vacuoles were present in the interior of the 3T3L1 adipocytes and were not open to the extracellular space (Fig. 1B). Consistent with these data, the

vacuolar structures were not accessible to the transferrin ligand at 4 °C (see Fig. 6). The formation of PIP5K-induced vacuoles was totally dependent on lipid kinase activity because kinase-inactive mutant forms of PIP5K␥ (90kDa/K138A) and (87kDa/K138A) had no effect (Fig. 2A, e– h and m–p). It should be noted that all of the type I PIP5K isoforms (PIP5K␥90 kDa, 87kDa, and PIP5K␤) were localized in the PIP5K-induced vacuoles but were also distributed in the plasma membrane in adipocytes (Fig. 2, A and B). To confirm that the PIP5K-induced vacuoles contain PI(4,5)P2, we coexpressed the PLC␦/PH domain as an EGFP fusion protein (PLC␦/PH-EGFP) that has a high affinity and specificity for PI(4,5)P2 (61, 62). Because PI(4,5)P2 is localized primarily to the plasma membrane, the PLC␦/PH-EGFP reporter was also confined to the plasma membrane in control cells (Fig. 2B, a). As expected, PIP5K-induced vacuoles displayed high levels of PI(4,5)P2 accumulation as assessed by the recruitment of the PLC␦/PH-EGFP fusion protein (Fig. 2B, b– d). As a control, we also coexpressed the PI(3,4,5)P3-selective PH domain from GRP1 as an EGFP fusion protein (61). Although the vacuole structures did recruit a small amount of the GRP1/PH-EGFP fusion protein, the extent of plasma membrane labeling was not significantly different from that of control cells (Fig. 2B, e– h). Furthermore, the phosphorylation of Akt/protein kinase B, a downstream target for the PI(3,4,5)P3 signaling pathway, was not detected in PIP5K-expressing 3T3L1 adipocytes (data not shown). Thus, the small amount of vacuole labeling by the GRP1/PH-EGFP fusion protein probably reflects a small degree of cross-reactivity with PI(4,5)P2. In any case, these data demonstrate that the PIP5K-induced vacuoles resulted primarily from the accumulation of PI(4,5)P2 in these structures. PIP5K-induced Vacuoles Are Coated with Stabilized Filamentous Actin—To examine the connection of F-actin with these PIP5K-induced vacuole structures, we assessed the colocalization of several proteins associated with F-actin polymerization. For example, both cortactin and dynamin have been implicated as important regulators of actin comet tails and actin-dependent plasma membrane endocytosis (43– 45, 63– 65). In the absence of PIP5K expression, cortactin was found dispersed throughout the cell periphery and was concentrated in several plasma membrane regions reminiscent of ruffling (Fig. 3A, a). However, expression of PIP5K induced the strong recruitment of cortactin to the vacuole structures (Fig. 3A, b– d). Similarly, dynamin was distributed primarily at the plasma membrane in control cells but also redistributed into these vacuole structures after PIP5K expression (Fig. 3A, e– h). In addition, the ␣-subunit of clathrin adaptor protein (AP-2) was also distributed in a clustered pattern along the plasma membrane but after PIP5K expression was recruited dramatically to the PIP5K-induced vacuoles (Fig. 3A, j–l). Several studies have also reported that although N-WASP was generally thought to be a cytosolic protein, it is localized predominantly in the nucleus in intact cells (66, 67). Consistent with these reports, expression of N-WASP as well as a dominant-interfering N-WASP mutant (N-WASP/⌬VCA) resulted in a strong nuclear localization in adipocytes (Fig. 3B, a and f). However, both N-WASP/WT and N-WASP/⌬VCA were strongly recruited into the PIP5K-induced vacuoles (Fig. 3B, b– e and g–j, arrowheads). Despite strong accumulation of a dominantinterfering N-WASP/⌬VCA on these vacuoles, there were essentially no effects on either formation of the PIP5K-induced vacuoles or their coated F-actin structure. The fact that the dominant-interfering N-WASP/⌬VCA mutant did not prevent the formation of these PIP5K-induced structures suggests that actin reorganization was not involved


30625

FIG. 2. Overexpression of type I PIP5K induces PI(4,5)P2-containing multiple vacuoles coated with Factin in 3T3L1 adipocytes. A, 3T3L1 adipocytes were electroporated with the cDNAs encoding EGFP-tagged PIP5K␥90kDa (a– d), its K138A mutant (e– h), PIP5K␥87kDa (i–l), or its K138A mutant (m–p). 18 h later, the cells were fixed and subjected to confocal microscopy for EGFP (a, e, i, and m), rhodamine-phalloidin (b, f, j, and n), and DAPI (c, g, k, and o). The merged images are shown in d, h, l, and p. These are a representative field of cells from three or four independent experiments. B, 3T3L1 adipocytes were electroporated with the empty vector (a and e) or cDNAs encoding HA-tagged PIP5K␤ plus cDNA encoding either EGFP-tagged PLC␦/PH domain (a– d) or EGFP-tagged GRP/PH domain (e– h). 18 h later, the cells were serum starved, fixed, and subjected to confocal fluorescent microscopy for the expressed HA-tagged PIP5K␤ (b and f) and expressed EGFPtagged PLC␦/PH (c) or EGFP-tagged GRP/PH (g). The merged images are presented in d and h and are representative fields of cells from three or four independent experiments.

in this process. To address this issue more specifically, we next treated adipocytes with latrunculin B, a specific actin monomer-sequestering reagent (68). Although latrunculin B completely disrupted cortical actin, the F-actin associated with the PIP5K-induced vacuole remained largely intact (Fig. 4A, a– c, arrow). Similarly, C. difficile toxin B, a specific inhibitor of Rho function (69), also resulted in a similar F-actin phenotype displaying a total disruption of the cortical actin with a partial loss of F-actin on the PIP5K-induced vacuoles (Fig. 4, d–f). Moreover, even though PIP5K-induced vacuoles strongly recruited the dominant-interfering mutant form of TC10 (TC10/ T31N) and reduced the extent of vacuole-coated F-actin, the vacuoles themselves remained essentially unaffected (Fig. 4B, a– d). Taken together, these data demonstrate that the formation of PIP5K-induced vacuoles is not dependent on the assembly of F-actin but that stabilized actin reorganizes around the PIP(4,5)P2 preformed vacuole structures.

PIP5K-induced Vacuoles Do Not Contain Secretory Membrane Components—It has been reported that in both PIP5K and constitutively active Arf6 (Arf6/Q67L)-transfected HeLa cells the formation of these enlarged F-actin-coated vacuoles appeared to result from a convergent of endosomes derived from both clathrin- and non-clathrin-dependent endocytic vesicles (70, 71). To characterize PIP5K-induced vacuoles in 3T3L1 adipocytes, we next examined the colocalization of these vacuoles with antibodies directed against several endomembrane markers including early endosome antigen (EEA1), p115, syntaxin 6, the cation-independent mannose 6-phosphate receptor (M6PR) and the clathrin heavy chain (Fig. 5). The PIP5K-induced vacuoles were devoid of early endosome markers such as EEA1 (Fig. 5A, a, e, and i), the Golgi marker p115 (Fig. 5A, b, f, and j), the TGN marker syntaxin 6 (Fig. 5A, c, g, and k), the TGN-lysosome marker M6PR (Fig. 5A, d, h, and l) and the early endosomal vesicle protein clathrin (Fig. 5B, a, e,

30626


FIG. 3. PIP5K-induced vacuoles recruit N-WASP, cortactin, and dynamin. A, 3T3L1 adipocytes were electroporated with the empty vector (a, e, and i) or cDNA encoding EGFP-PIP5K␥87kDa alone (b– d and j–l) or plus cDNA encoding HA-tagged dynamin 2 (f– h). 18 h later, the cells were fixed and stained with a cortactin antibody (a and b), a HA antibody (e and f), or a ␣-adaptin (AP-2) antibody (i and j), or with the anti-HA antibody (e and d) and Alexa 594-conjugated secondary IgG. The subcellular localization was observed under confocal fluorescent microscopy. These are representative fields of three independent experiments. B, 3T3L1 adipocytes were electroporated with the empty vector (a and f) or cDNAs encoding HA-tagged PIP5K␤ plus either EGFPtagged N-WASP (a– e) or N-WASP/⌬VCA (f–j). 18 h later, the cells were fixed and subjected to confocal fluorescent microscopy for HA-PIP5K␤ stained with the anti-HA antibody and Alexa 647-conjugated secondary IgG (b and g), EGFP-NWASP/WT (c), or its ⌬VCA mutant (h), and F-actin labeled with rhodamine-phalloidin (d and i). The merged images are shown in e and j. These are representative fields of three independent experiments.

and i). Furthermore, exogenously expressed VSV-G, a well characterized virus glycoprotein that traffics through constitutive secretory membrane transport vesicles, was also absent in the PIP5K-induced vacuoles (Fig. 5B, d, h, and l). In contrast, the integrin ␤1 receptor subunit (Fig. 5B, b, f, and j) was observed in the PIP5K-induced vacuoles as reported previously for Arf6-expressing HeLa cells (72). The integrin ␤1 receptor subunit has been reported to traffic through the Arf6-regulated endosomal recycling system (72–74). In addition to the integrin ␤1 receptor subunit, the PIP5K-induced vacuoles also contained caveolin (Fig. 5B, c, g, and k), which is also known to undergo endocytosis through a clathrin-independent pathway (75–77). Similarly, TC10 has been well established to localize to caveolin-enriched plasma membrane lipid raft microdomain (78, 79), and was also found to concentrate in the PIP5Kinduced vacuoles (Fig. 4B). Consistent with these results, Texas Red-conjugated cholera toxin B subunit, which binds to lipid raft resident ganglioside GM1, was accumulated in the PIP5K-induced vacuoles (data not shown). Thus, the accumulation of several adipocyte endocytotic and lipid raft microdomain markers in the PIP5K-induce vacuoles is consistent with

these structures being derived from non-clathrin-dependent endocytic vesicles. Transferrin Receptor Accumulates in the PIP5K-induced Vacuoles after Endocytosis—Because these data suggest the presence of a non-clathrin endocytotic trafficking route to the PIP5K-induced vacuoles, we next assessed the possible convergence with the classic clathrin-dependent endocytosis pathway. To accomplish this, we coexpressed the human TfR and examined its trafficking in 3T3L1 adipocytes. 18 h after transfection, cells were cooled to 4 °C and then labeled with the Texas Red-conjugated transferrin (Texas Red-transferrin) for 1 h. At this time point, a substantial amount of TfR was found at the plasma as well as in the PIP5K-induced vacuoles (Fig. 6A, e) and displayed both plasma membrane and vacuolar distribution after warming to 37 °C (Fig. 6A, f– h). In contrast, prior to warming the transferrin ligand was confined primarily to the cell surface with no interior TfR labeling (Fig. 6A, i). However, there was a time-dependent accumulation of Texas Red-transferrin in the PIP5K-induced vacuoles (Fig. 6A, j–l), and most of PIP5K-induced vacuoles were labeled with Texas Red-transferrin after reincubating at 37 °C for 120 min (Fig. 6A, l). These


30627

FIG. 4. Latrunculin B, C. difficile toxin B, and TC10/T31N do not prevent the formation of the PIP5K-induced vacuoles. A, 3T3L1 adipocytes were electroporated with PIP5K␥87kDa cDNA. 18 h later, the cells were incubated with 20 ␮M latrunculin B (a– c) or 0.5 mg/ml toxin B (d–f) for 2 h at 37 °C and then fixed. The cells were subjected to confocal fluorescent microscopy for EGFPPIP5K (a and d) and F-actin labeled with rhodamine-phalloidin (b and e). The merged images are shown in c and f. These are representative fields of three independent experiments. B, 3T3L1 adipocytes were electroporated with cDNAs encoding EGFP-PIP5K␥87kDa and Myc-tagged TC10/T31N. 18 h later, cells were fixed and subjected to confocal fluorescent microscopy for EGFP-PIP5K (a), F-actin labeled with rhodamine-phalloidin (b), and Myctagged TC10/T31N labeled with the antiMyc antibody and Alexa 647-conjugated secondary IgG (c). The merged image is shown in d. This is a representative field of three independent experiments.

data coupled with the steady-state colocalization of caveolin and integrin ␤1 receptor subunit (Fig. 4B) demonstrate that the PIP5K-induced vacuoles result from the convergence of endosomes derived from clathrin- and non-clathrin endocytosis. We also quantified the rate of TfR endocytosis by counting the number of cells displaying internalized Texas Red-transferrin at 5, 15, 30, and 60 min after reincubation at 37 °C (Fig. 6B). Expression of the kinase-deficient PIP5K mutant (PIP5K␤/D307A) had no significant effect on the rate of TfR endocytosis compared with control transfected cells. In contrast, adipocytes expressing wild-type PIP5K␤ (PIP5K␤/WT) displayed a reduced rate of TfR endocytosis. Overexpression of PIP5K Results in a Persistent GLUT4 Localization at the Plasma Membrane Because of an Inhibition of GLUT4 Endocytosis—It is generally accepted that the TfR and GLUT4 share the same initial steps of plasma membrane endocytosis but diverge at some point during endocytotic recycling (80 – 82). We therefore next examined the effect of PIP5K overexpression on insulin-induced GLUT4 translocation. Coexpression of GLUT4-EGFP with the empty vector resulted in the characteristic localization of GLUT4-EGFP to the perinuclear regions and various small vesicular compartments scattered

throughout the cytoplasm (Fig. 7A, a). Insulin stimulation resulted in a redistribution of the intracellular compartmentalized GLUT4-EGFP to the plasma membrane (Fig. 7A, b). Expression of wild-type PIP5K␤ resulted in the formation of the multiple aggregated vacuoles and the accumulation of GLUT4EGFP in these structures (Fig. 7A, c and d). Furthermore, GLUT4-EGFP was plasma membrane-localized in both the basal and insulin-stimulated states. In contrast, expression of the kinase-deficient PIP5K␤/D307A mutant had no significant effect on the distribution of GLUT4-EGFP (Fig. 7A, e and f). As summarized in Fig. 7B, under basal conditions only 11.7 ⫾ 6.8% of cells displayed plasma membrane localization of GLUT4-EGFP, whereas insulin stimulation resulted in 81.7 ⫾ 2.3% of the cells with a cell surface distribution. Essentially identical results were obtained in cells expressing the PIP5K␤/ D307A mutant. Although expression of PIP5K␤/WT did not affect the extent of insulin-stimulated GLUT4 translocation, in this case 56.2 ⫾ 6.5% of cells displayed plasma membrane localization in the basal state. The persistent localization of GLUT4 at the plasma membrane caused by the PIP5K␤/WT could result from either a stimulation of GLUT4 exocytosis and/or an inhibition of its

30628


FIG. 5. Characterization of PIP5Kinduced vacuoles in 3T3L1 adipocytes. A, 3T3L1 adipocytes were electroporated with cDNA encoding EGFPtagged PIP5K␥87kDa. 18 h later, the cells were fixed and subjected to confocal fluorescent microscopy for the expressed EGFP-tagged PIP5K␥87kDa and following endomembrane marker proteins. The cells were stained for early endosome marker (EEA1, a), cis-Golgi marker (p115, b), TGN marker (syntaxin 6, c), a marker protein that shuttles between endosome and TGN (mannose 6-phosphate receptor, M6P-R, d). B, the cells were stained for clathrin heavy chain (Clathrin-HC, a), integrin ␤1 (b), and caveolin 1 (c). For the VSV-G experiments, 3T3L1 adipocytes were electroporated with cDNA encoding EGFP-tagged PIP5K ␥ 87kDa plus cDNA encoding VSV-G (d, h, and l). After fixing the cells, VSV-G was stained with the VSV-G antibody (d). The merged images are shown in i–l. These are representative fields of the cells from three independent experiments.

endocytosis. The accumulation of endocytotic TfR (Fig. 6A) and its significant decreased rate of endocytosis (Fig. 6B) strongly suggest that the accumulation of GLUT4 in these structures also results from an inhibition of plasma membrane endocytosis. Therefore, we examined the localization of GLUT4-EGFP at an earlier time point (9 h after electroporation) in PIP5K␤expressing 3T3L1 adipocytes (Fig. 7C). At this time, the newly synthesized GLUT4 is just transported into the GLUT4 storage compartment and has yet recycled through the plasma membrane (83). 9 h after transfection, the F-actin-coated vacuoles were already formed in 3T3L1 adipocytes expressing PIP5K␤/WT (Fig. 7C, a– d), whereas the coexpressed GLUT4EGFP was not localized in these PIP5K-induced vacuoles (Fig. 7C, c). Furthermore, GLUT4-EGFP was not detected at the plasma membrane at this time point (Fig. 7C, a– c, arrowheads). However, an accumulation of GLUT4-EGFP at both plasma membrane (arrowheads) and PIP5K-induced vacuoles (arrow) was observed 18 h after transfection even in the absence of insulin (Fig. 7C, e– h). Because the relative long time required for GLUT4 accumulation in the PIP5K-induced vacuoles (18 h) is consistent with

a slow recycling process from the plasma membrane, we took advantage of an exofacial tagged Myc-GLUT4-EGFP construct. As observed previously, expression of Myc-GLUT4-EGFP with PIP5K␤/WT for 18 h resulted in the colocalization of GLUT4 with the PIP5K-induced vacuoles (Fig. 8A, a and b). Similarly, incubating the Myc-GLUT4-EGFP-expressing cells with the Myc antibody continually for 3 h also resulted in the endocytosis of the Myc epitope into the PIP5K-induced vacuoles (Fig. 8A, c). This result clearly demonstrates that the accumulation of the labeled Myc-GLUT4-EGFP in PIP5K-induced vacuoles was caused by plasma membrane endocytosis and not a direct trafficking from other intracellular compartments. To quantify the rate of GLUT4 endocytosis, the cells were first stimulated with insulin for 30 min, then labeled with the Myc antibody at 4 °C followed by acid washing to remove both insulin and unbound Myc antibody, and then warmed to 37 °C for various times. The Myc-GLUT4-EGFP was strongly labeled by the Texas Red-conjugated secondary antibody at the cell surface with no interior labeling at 4 °C (data not shown). In control cells, there was a rapid time-dependent internalization of the labeled GLUT4, which concentrated in small compartments

30629


FIG. 6. PIP5K inhibits TfR endocytosis in 3T3L1 adipocytes. A, 3T3L1 adipocytes were electroporated with cDNA encoding EGFP-tagged PIP5K ␥ 87kDa plus cDNA encoding human TfR. 18 h later, the cells were chilled, incubated with 5 ␮g/ml of Texas Red-conjugated transferrin at 4 °C for 1 h, and warmed to 37 °C for the indicated periods. The cells were then fixed and subjected to confocal fluorescent microscopy for EGFP-PIP5K (a--d), TfR (e– h), and transferrin ligand (i–l). The merged images are shown in m– q. These are representative fields of three independent experiments. B, 3T3L1 adipocytes were electroporated with cDNA encoding human TfR plus cDNA encoding empty vector, EGFP-tagged PIP5K␥87kDa, or PIP5K␥87kDa/K138A. 18 h later, the cells were labeled with Texas Red-transferrin at 4 °C and then warmed to 37 °C for the indicated period. After fixing the cells, the amount of internalized transferrin was quantified by counting the number of cells displaying intracellular labeling relative to control vector-transfected cells. These data were obtained from the counting a total 50 – 60 cells from three independent experiments.

beneath the plasma membrane and in the perinuclear region as described previously (11). As observed for the TfR, overexpression of PIP5K1␤/WT, but not the kinase-defective mutant PIP5K1␤/D307A, resulted in a marked inhibition of GLUT4 endocytosis (Fig. 8B). These data demonstrate that overexpression of PIP5K inhibits GLUT4 endocytosis, which consequently results in its accumulation at the plasma membrane. PI(4,5)P2-induced Phenotype Is Dependent upon Cholesterol Levels—Recently it was reported that cholesterol levels directly affect the plasma membrane organization of PI(4,5)P2, which in turn modulates actin cytoskeleton (84). Furthermore, experimental modulation of cellular cholesterol levels affects multiple membrane trafficking events (85, 86) including GLUT4 endocytosis (87, 123). Because adipocytes express high levels of caveolin and contain large amounts of caveolae/lipid rafts enriched in cholesterol and sphingolipids (88, 89), we hypothesized that the phenotype induced by PIP5K expression was dependent upon cellular cholesterol levels. To address this possibility, we examined the effect of cholesterol loading on PIP5K-induced vacuole formation (Fig. 9). As observed previously, expression of PIP5K resulted in the formation of actin comet tails in 3T3L1 fibroblasts (Fig. 9, a– c). In contrast, cholesterol loading by itself had no obvious effect (data not shown), but in the presence of PIP5K overexpression now resulted in aggregated vacuole formation (Fig. 9, d–f). Interestingly, although some of the cholesterol-loaded PIP5K-induced

vacuoles in fibroblast displayed F-actin coating, many did not (Fig. 9, d–f, arrows and arrowheads). This suggests that in fibroblasts the amount of F-actin and/or factors responsible for its assembly may be limiting but supports the observation that actin assembly was not primarily required for PIP5K-induced vacuole formation (Fig. 4). Together, these data indicate that amount of cholesterol is involved in the formation of vacuoles induced by PIP5K expression, suggesting that composition of membrane lipids including cholesterol and PI(4,5)P2 can directly affect multiple membrane trafficking events and actin regulation. DISCUSSION

In the context of insulin action, PI(4,5)P2 is well established as the precursor for synthesizing PI(3,4,5)P3 which is a critical second messenger directly involved in GLUT4 translocation, and intense research has focused on generation/degradation of 3⬘-phosphoinositides and its downstream effectors (20, 90). It is generally well accepted that PI(3,4,5)P3 serves as a second messenger to activate both 3⬘-phosphoinositide protein kinase and Akt/protein kinase B leading to GLUT4 vesicle exocytosis (91). However, in fibroblasts ectopically expressing both insulin receptor and GLUT4, insulin fails to induce GLUT4 translocation (92). One possibility is that proper distribution of and trafficking of GLUT4 protein in response to insulin requires specialized and/or compartmentalized adipocyte membrane lip-

30630


FIG. 7. Expression of PIP5K results in increased plasma membrane localized GLUT4 protein. A, 3T3L1 adipocytes were electroporated with 50 ␮g of cDNA encoding GLUT4-EGFP plus 200 ␮g of cDNA encoding empty vector (a and b), PIP5K␤/WT (c and d), or PIP5K␤/ D307A (e and f). 18 h later, the cells were serum starved and incubated in the absence (a, c, and e) or presence (b, d, and f) of 100 nM insulin for 30 min. The cells were fixed, and the subcellular localization of GLUT4-EGFP was determined by confocal fluorescent microscopy. These are representative fields of three independent experiments. B, quantification of the number of cells displaying GLUT4EGFP plasma membrane fluorescence was determined from the counting of 50 – 60 cells in three independent experiments. C, 3T3L1 adipocytes were electroporated with cDNAs encoding GLUT4EGFP and HA-tagged PIP5K␤/WT. The cells were serum starved and fixed at 9 h (a– d) or 18 h (e– h) after electroporation. The cells were subjected to confocal fluorescent microscopy for HA-PIP5K␤/WT using the HA antibody (a and e), F-actin labeled with rhodamine-phalloidin (b and f), and GLUT4-EGFP (c and g). The merged images are shown in d and h. These are representative fields of three independent experiments.

FIG. 8. Expression of PIP5K inhibits GLUT4 endocytosis. A, 3T3L1 adipocytes were electroporated with cDNAs encoding GLUT4-EGFP and PIP5K␤/WT. 18 h later, the cells were incubated with the Myc antibody for 3 h, fixed, and the distribution of the Myc antibody was examined by confocal fluorescent microscopy. This is a representative image from three independent experiments. B, 3T3L1 adipocytes were electroporated with 50 ␮g of cDNA encoding GLUT4-EGFP plus 200 ␮g of cDNA encoding empty vector, PIP5K␤/WT, or PIP5K␤/D307A. 18 h later, the cells were serum starved, incubated with 100 nM insulin for 30 min at 37 °C, and then cooled to 4 °C. The cells were incubated with the Myc antibody for 1 h, washed extensively to remove the insulin and unbound antibody, and returned to 37 °C for the indicated times. The Myc antibody-labeled Myc-GLUT4-EGFP was detected by using Alexa 594-conjugated secondary IgG, and the presence of intracellular localization of Myc epitope was examined. These data were obtained from counting of 50 – 60 cells/experiment from three independent determinations.

ids (78). In this regard, a recent study has suggested that the role of insulin-stimulated PI 3-kinase activation in GLUT4 translocation is the loss or masking of PI(4,5)P2 rather than the generation of PI(3,4,5)P3 itself (93). Consistent with this hy-

pothesis, endothelin-1 can induce GLUT4 translocation (94, 95) and results in decreased in PI(4,5)P2 levels through Gq-mediated PLC activation (96, 97). Similarly, persistent PLC activation and subsequent decrease in PIP2 levels by expression of a


30631

FIG. 9. Cholesterol loading induces aggregated vacuole formation in 3T3L1 fibroblasts expressing PIP5K. 3T3L1 preadipocytes (fibroblasts) were electroporated with cDNA encoding EGFP-tagged PIP5K␥87kDa. The cells were cultured in the absence (a– c) or presence (d–f) of 0.1 mM cholesterol-methyl-␤-cyclodextrin. 18 h later, the cells were fixed and subjected to confocal microscopy for EGFP (a and d) and rhodamine-phalloidin (b and e). The merged images are shown in c and f. Arrowheads and arrows indicate vacuoles with and without F-actin, respectively. This is a representative image from three independent experiments.

constitutively active mutant form ␣-subunit of Gq/11 stimulates GLUT4 translocation (58, 98), most likely through an Arf6-dependent mechanism (99, 100). These data are also analogous to the requirement for the Arf6-dependent dynamic turnover of PI(4,5)P2 for growth hormone release in PC12 cells and insulin secretion in MIN6 cells (101, 102). The cellular content of PI(4,5)P2 is dynamically regulated by lipid kinases, phosphatases, and the availability of PI precursors. The major pathway for PI(4,5)P2 synthesis is by sequential phosphorylation of PI by a PI 4-kinase to generate PI4P followed by phosphorylation with the type I PIP5K (103). Alternatively, PI(4,5)P2 can also be synthesized by the type II kinases via the recently discovered lipid PI5P (104). Currently, three PIP5K isoforms (␣, ␤, and ␥) have been identified which display distinct tissue and developmental regulation (55, 56, 105), and recent studies have demonstrated that all of these enzymes are activated by Rho family small GTPases such as RhoA, Cdc42, and Rac1 (106). Previous studies have also reported that exogenous expression of PIP5K induced dramatic changes in actin cytoskeleton with different phenotypes of actin filaments depending on cell type. For example, Swiss 3T3 cells, REF52 cells, and mouse embryonic fibroblast cells (41, 42) expressing PIP5K form N-WASP-dependent actin comet tails, whereas CV1 cells expressing PIP5K stimulate stress fiber formation, inhibit membrane ruffling, and block the generation of actin nuclei (107). In 293T cells (70), and HeLa cells (72), overexpression of PIP5K resulted in the accumulation of large vacuoles coated with F-actin. The data presented in this study demonstrated that the phenotype induced by increased levels of PI(4,5)P2 production by PIP5Ks expression were different between 3T3L1 preadipocytes (fibroblasts) and adipocytes. In preadipocytes PIP5K expression resulted in actin comet tails but induced the formation of F-actin-coated multiple aggregated vacuoles in differentiated adipocytes. Moreover, we found in 3T3L1 fibroblasts that the amount of cholesterol is one of the key factors involved in this phenotype conversion because cholesterol loading resulted in the formation of intracellular vacuoles in fibroblasts expressing PIP5K. Although the precise mechanism underlying this cholesterol-induced phenotype conversion remains unclear, a possibility that cholesterol can directly affect membrane thickness and curvature may be involved in this phenomenon (85, 108, 109). In addition, several lines of evidence have indicated that the amount of cellular cholesterol directly influences PI(4,5)P2 levels and distribution, which subsequently affect

actin cytoskeletal organization and multiple membrane trafficking processes including GLUT4 endocytosis (84, 86, 87). We also reported that the organization of the actin cytoskeleton is changed dramatically during the differentiation of adipocytes from a typical stress fiber actin structure in fibroblasts to a relatively thick cortical actin lining the inner surface of the plasma membrane in adipocytes (13, 110). In differentiated 3T3L1 adipocytes, the relatively thick cortical actin structure appeared to be composed of F-actin spikes emanating from organized rosette-like cluster of caveolae/lipid raft microdomains (110). The data presented here further document that actin regulatory mechanisms are altered dramatically during adipogenesis and underscore the specificity of actin organization and function, which are dependent upon cellular context including composition and amount of membrane lipids. Recently, the PI(4,5)P2 regulation of actin dynamics has also been implicated in the control of vesicle trafficking through the recruitment of N-WASP (41, 42, 111). In the present study, we also found that the PIP5K-induced vacuoles recruited N-WASP, dynamin, and cortactin. However, expression of the dominant-interfering N-WASP mutant failed to block actin accumulation. Furthermore, treatment with latrunculin B (an inhibitor of actin polymerization) and toxin B (a broad Rho family GTPase inhibitor) also had minimal effects on both the structure of PIP5K-induced vacuoles and the distribution of vacuole-coated F-actin. Despite the apparent link between PI(4,5)P2 formation and the stimulation of actin-based motility in other systems, our data suggest that actin polymerization is not primarily required for forming and/or maintaining PIP5Kinduced vacuoles in 3T3L1 adipocytes. The type I PIP5Ks have been implicated in various aspects of membrane transport including endocytosis, exocytosis, and vesicle budding/formation all through regulating clathrin coat assembly (103, 112, 113). For example, immunoabsorption of the type I PIP5K and depletion of PI(4,5)P2 from the plasma membrane resulted in a decreased vesicle priming required for exocytosis and inhibition of dense core vesicle exocytosis (102). PI(4,5)P2 has also been implicated in plasma membrane endocytosis because several proteins important for the assembly of clathrin-coated vesicles bind to PI(4,5)P2 (113, 114). These proteins include the AP-2 adaptor responsible for binding endocytic vesicles (47, 50); epsin, which recruits clathrin and curves membranes (115); and dynamin, a large GTP-binding protein required for the fission of clathrin-coated pits (53, 116). Thus, it is generally accepted that type I PIP5K recruitment and sub-

30632


sequent PI(4,5)P2 accumulation in the plasma membrane facilitate endocytosis in various cell types (117–119). In fact, a moderate increase in PI(4,5)P2 levels by exogenous PIP5K1␤ expression in CV-1 cells facilitates endocytosis of TfR concomitant with an increase in the number of clathrin-coated pits associated with the AP-2 complex (119). Moreover, Arf6 was found to stimulate clathrin/AP-2 recruitment to synaptic membranes, thereby facilitating clathrin-coated pit assembly by activating PIP5K␥ (120). In adipocytes, these PIP5K-induced vacuoles were devoid of any secretory membrane markers, indicating that these structures are distinct from the biosynthetic membrane trafficking pathway. Although clathrin was also excluded from these compartments, several lipid raft markers including caveolin and TC10 were found to accumulate. Despite the apparent absence of clathrin, TfR endocytosis was reduced significantly and was observed to recycle through these vacuoles. These data are consistent with the formation of PIP5K-induced vacuoles from a convergence of both clathrin-dependent and non-clathrin-dependent endocytosis. The absence of clathrin from this compartment suggests that the uncoating of newly clathrin-derived endocytotic vesicles was not inhibited by expression of PIP5K. Thus, these transport vesicles would be able to fuse and form these vacuolar compartments along with non-clathrin-mediated endocytotic vesicles. This conclusion is supported further by the accumulation of both the TfR and GLUT4 in the PIP5Kinduced vacuoles. In addition to the slow accumulation of lipid raft and nonlipid raft markers into these PIP5K-induced structures, the rate of TfR and GLUT4 endocytosis was also significantly inhibited. Although clathrin was not found within the PIP5Kinduced vacuoles, there was a clear accumulation of the AP-2 complex. Similarly, dynamin was also found to accumulate in the PI,(4,5)P2-enriched vacuoles. Thus, in 3T3L1 adipocytes the formation of the PI(4,5)P2-enriched vacuoles leads to the relative loss of plasma membrane AP-2 and/or dynamin, which probably accounts for the reduced rate of TfR and GLUT4 endocytosis under these conditions. Because PI(4,5)P2 binds with high affinity to the ␣-subunit of the AP-2 complex (47, 121), we speculate that this results in an inhibition of AP-2 uncoating. The persistent and stable formation of AP-2 membrane transport coats would, in turn, result in a reduced rate of endocytosis and a slow accumulation of recycling proteins in the PI(4,5)P2-enriched compartments. This also implies that AP-2 uncoating requires degradation of PI(4,5)P2 in the endocytotic vesicles, whereas clathrin-uncoating is independent of both PI(4,5)P2 turnover and AP-2 vesicle assembly. Although we took advantage of overexpression of PIP5K to explore the potential functional role of PI(4,5)P2 in membrane trafficking and actin polymerization, the accumulation of similar endosomal structures has been also observed in nerve terminals of mice lacking synaptojanin (PIP2 5-phosphatase) and in Sertoli cells of mice lacking InPP5, an inositol polyphosphate 5-phosphatase (48, 122). Together these data provide strong evidence that PI(4,5)P2 and its dynamic turnover through regulation of these lipid enzymes are physiologically critical for maintaining proper membrane trafficking in intact cells. Acknowledgments—We thank Dr. Richard Anderson for the kind gifts of pEGFP-PIP5K␥ cDNAs. REFERENCES 1. Pessin, J. E., Thurmond, D. C., Elmendorf, J. S., Coker, K. J., and Okada, S. (1999) J. Biol. Chem. 274, 2593–2596 2. Bryant, N. J., Govers, R., and James, D. E. (2002) Nat. Rev. Mol. Cell. Biol. 3, 267–277 3. Rudich, A., and Klip, A. (2003) Acta Physiol. Scand. 178, 297–308 4. Jhun, B. H., Rampal, A. L., Liu, H., Lachaal, M., and Jung, C. Y. (1992) J. Biol. Chem. 267, 17710 –17715 5. Czech, M. P., and Buxton, J. M. (1993) J. Biol. Chem. 268, 9187–9190

6. Yang, J., and Holman, G. D. (1993) J. Biol. Chem. 268, 4600 – 4603 7. Fletcher, L. M., Welsh, G. I., Oatey, P. B., and Tavare, J. M. (2000) Biochem. J. 352, 267–276 8. Emoto, M., Langille, S. E., and Czech, M. P. (2001) J. Biol. Chem. 276, 10677–10682 9. Guilherme, A., Emoto, M., Buxton, J. M., Bose, S., Sabini, R., Theurkauf, W. E., Leszyk, J., and Czech, M. P. (2000) J. Biol. Chem. 275, 38151–38159 10. Olson, A. L., Trumbly, A. R., and Gibson, G. V. (2001) J. Biol. Chem. 276, 10706 –10714 11. Shigematsu, S., Khan, A. H., Kanzaki, M., and Pessin, J. E. (2002) Mol. Endocrinol. 16, 1060 –1068 12. Liu, L. B., Omata, W., Kojima, I., and Shibata, H. (2003) J. Biol. Chem. 278, 30157–30169 13. Kanzaki, M., and Pessin, J. E. (2001) J. Biol. Chem. 276, 42436 – 42444 14. Jiang, Z. Y., Chawla, A., Bose, A., Way, M., and Czech, M. P. (2002) J. Biol. Chem. 277, 509 –515 15. Wang, Q., Bilan, P. J., Tsakiridis, T., Hinek, A., and Klip, A. (1998) Biochem. J. 331, 917–928 16. Omata, W., Shibata, H., Li, L., Takata, K., and Kojima, I. (2000) Biochem. J. 346, 321–328 17. Patki, V., Buxton, J., Chawla, A., Lifshitz, L., Fogarty, K., Carrington, W., Tuft, R., and Corvera, S. (2001) Mol. Biol. Cell 12, 129 –141 18. Tong, P., Khayat, Z. A., Huang, C., Patel, N., Ueyama, A., and Klip, A. (2001) J. Clin. Invest. 108, 371–381 19. Okada, T., Kawano, Y., Sakakibara, R., Hazeki, O., and Ui, M. (1994) J. Biol. Chem. 269, 3568 –3573 20. Corvera, S., and Czech, M. P. (1998) Trends Cell Biol. 8, 442– 446 21. Khayat, Z. A., Tong, P., Yaworsky, K., Bloch, R. J., and Klip, A. (2000) J. Cell Sci. 113, 279 –290 22. Martin, S. S., Haruta, T., Morris, A. J., Klippel, A., Williams, L. T., and Olefsky, J. M. (1996) J. Biol. Chem. 271, 17605–17608 23. Chiang, S. H., Baumann, C. A., Kanzaki, M., Thurmond, D. C., Watson, R. T., Neudauer, C. L., Macara, I. G., Pessin, J. E., and Saltiel, A. R. (2001) Nature 410, 944 –948 24. Kanzaki, M., Watson, R. T., Hou, J. C., Stamnes, M., Saltiel, A. R., and Pessin, J. E. (2002) Mol. Biol. Cell 13, 2334 –2346 25. Stamnes, M. (2002) Curr. Opin. Cell Biol. 14, 428 – 433 26. Orth, J. D., and McNiven, M. A. (2003) Curr. Opin. Cell Biol. 15, 31–39 27. Schafer, D. A. (2002) Curr. Opin. Cell Biol. 14, 76 – 81 28. Qualmann, B., and Kessels, M. M. (2002) Int. Rev. Cytol. 220, 93–144 29. Hussain, N. K., Jenna, S., Glogauer, M., Quinn, C. C., Wasiak, S., Guipponi, M., Antonarakis, S. E., Kay, B. K., Stossel, T. P., Lamarche-Vane, N., and McPherson, P. S. (2001) Nat. Cell Biol. 3, 927–932 30. Jahraus, A., Egeberg, M., Hinner, B., Habermann, A., Sackman, E., Pralle, A., Faulstich, H., Rybin, V., Defacque, H., and Griffiths, G. (2001) Mol. Biol. Cell 12, 155–170 31. Eitzen, G., Wang, L., Thorngren, N., and Wickner, W. (2002) J. Cell Biol. 158, 669 – 679 32. Musch, A., Cohen, D., Kreitzer, G., and Rodriguez-Boulan, E. (2001) EMBO J. 20, 2171–2179 33. Luna, A., Matas, O. B., Martinez-Menarguez, J. A., Mato, E., Duran, J. M., Ballesta, J., Way, M., and Egea, G. (2002) Mol. Biol. Cell 13, 866 – 879 34. di Campli, A., Valderrama, F., Babia, T., De Matteis, M. A., Luini, A., and Egea, G. (1999) Cell Motil. Cytoskeleton 43, 334 –348 35. Valderrama, F., Babia, T., Ayala, I., Kok, J. W., Renau-Piqueras, J., and Egea, G. (1998) Eur. J. Cell Biol. 76, 9 –17 36. Valderrama, F., Duran, J. M., Babia, T., Barth, H., Renau-Piqueras, J., and Egea, G. (2001) Traffic 2, 717–726 37. Rohatgi, R., Ma, L., Miki, H., Lopez, M., Kirchhausen, T., Takenawa, T., and Kirschner, M. W. (1999) Cell 97, 221–231 38. Prehoda, K. E., Scott, J. A., Mullins, R. D., and Lim, W. A. (2000) Science 290, 801– 806 39. Welch, M. D., and Mullins, R. D. (2002) Annu. Rev. Cell Dev. Biol. 18, 247–288 40. Kanzaki, M., Watson, R. T., Khan, A. H., and Pessin, J. E. (2001) J. Biol. Chem. 276, 49331– 49336 41. Rozelle, A. L., Machesky, L. M., Yamamoto, M., Driessens, M. H., Insall, R. H., Roth, M. G., Luby-Phelps, K., Marriott, G., Hall, A., and Yin, H. L. (2000) Curr. Biol. 10, 311–320 42. Benesch, S., Lommel, S., Steffen, A., Stradal, T. E., Scaplehorn, N., Way, M., Wehland, J., and Rottner, K. (2002) J. Biol. Chem. 277, 37771–37776 43. Schafer, D. A., Weed, S. A., Binns, D., Karginov, A. V., Parsons, J. T., and Cooper, J. A. (2002) Curr. Biol. 12, 1852–1857 44. Lee, E., and De Camilli, P. (2002) Proc. Natl. Acad. Sci. U. S. A. 99, 161–166 45. Orth, J. D., Krueger, E. W., Cao, H., and McNiven, M. A. (2002) Proc. Natl. Acad. Sci. U. S. A. 99, 167–172 46. Jost, M., Simpson, F., Kavran, J. M., Lemmon, M. A., and Schmid, S. L. (1998) Curr. Biol. 8, 1399 –1402 47. Gaidarov, I., and Keen, J. H. (1999) J. Cell Biol. 146, 755–764 48. Cremona, O., Di Paolo, G., Wenk, M. R., Luthi, A., Kim, W. T., Takei, K., Daniell, L., Nemoto, Y., Shears, S. B., Flavell, R. A., McCormick, D. A., and De Camilli, P. (1999) Cell 99, 179 –188 49. Itoh, T., Koshiba, S., Kigawa, T., Kikuchi, A., Yokoyama, S., and Takenawa, T. (2001) Science 291, 1047–1051 50. Rohde, G., Wenzel, D., and Haucke, V. (2002) J. Cell Biol. 158, 209 –214 51. Ford, M. G., Pearse, B. M., Higgins, M. K., Vallis, Y., Owen, D. J., Gibson, A., Hopkins, C. R., Evans, P. R., and McMahon, H. T. (2001) Science 291, 1051–1055 52. Mao, Y., Chen, J., Maynard, J. A., Zhang, B., and Quiocho, F. A. (2001) Cell 104, 433– 440 53. Lin, H. C., Barylko, B., Achiriloaie, M., and Albanesi, J. P. (1997) J. Biol. Chem. 272, 25999 –26004 54. Tebar, F., Bohlander, S. K., and Sorkin, A. (1999) Mol. Biol. Cell 10, 2687–2702 55. Ishihara, H., Shibasaki, Y., Kizuki, N., Katagiri, H., Yazaki, Y., Asano, T., and

PIP2 Regulation of GLUT4 Endocytosis Oka, Y. (1996) J. Biol. Chem. 271, 23611–23614 56. Loijens, J. C., and Anderson, R. A. (1996) J. Biol. Chem. 271, 32937–32943 57. Kao, A. W., Ceresa, B. P., Santeler, S. R., and Pessin, J. E. (1998) J. Biol. Chem. 273, 25450 –25457 58. Kanzaki, M., Watson, R. T., Artemyev, N. O., and Pessin, J. E. (2000) J. Biol. Chem. 275, 7167–7175 59. Min, J., Okada, S., Kanzaki, M., Elmendorf, J. S., Coker, K. J., Ceresa, B. P., Syu, L. J., Noda, Y., Saltiel, A. R., and Pessin, J. E. (1999) Mol Cell 3, 751–760 60. Pike, L. J., and Miller, J. M. (1998) J. Biol. Chem. 273, 22298 –22304 61. Kavran, J. M., Klein, D. E., Lee, A., Falasca, M., Isakoff, S. J., Skolnik, E. Y., and Lemmon, M. A. (1998) J. Biol. Chem. 273, 30497–30508 62. Varnai, P., and Balla, T. (1998) J. Cell Biol. 143, 501–510 63. Cao, H., Orth, J. D., Chen, J., Weller, S. G., Heuser, J. E., and McNiven, M. A. (2003) Mol. Cell. Biol. 23, 2162–2170 64. Weaver, A. M., Heuser, J. E., Karginov, A. V., Lee, W. L., Parsons, J. T., and Cooper, J. A. (2002) Curr. Biol. 12, 1270 –1278 65. Krueger, E. W., Orth, J. D., Cao, H., and McNiven, M. A. (2003) Mol. Biol. Cell 14, 1085–1096 66. Suetsugu, S., and Takenawa, T. (2003) J. Biol. Chem. 278, 42515– 42523 67. Mizutani, K., Suetsugu, S., and Takenawa, T. (2004) Biochem. Biophys. Res. Commun. 313, 468 – 474 68. Morton, W. M., Ayscough, K. R., and McLaughlin, P. J. (2000) Nat. Cell Biol. 2, 376 –378 69. Just, I., Selzer, J., Wilm, M., von Eichel-Streiber, C., Mann, M., and Aktories, K. (1995) Nature 375, 500 –503 70. Galiano, F. J., Ulug, E. T., and Davis, J. N. (2002) J. Cell. Biochem. 85, 131–145 71. Naslavsky, N., Weigert, R., and Donaldson, J. G. (2003) Mol. Biol. Cell 14, 417– 431 72. Brown, F. D., Rozelle, A. L., Yin, H. L., Balla, T., and Donaldson, J. G. (2001) J. Cell Biol. 154, 1007–1018 73. Powelka, A. M., Sun, J., Li, J., Gao, M., Shaw, L. M., Sonnenberg, A., and Hsu, V. W. (2004) Traffic 5, 20 –36 74. Wong, K. W., and Isberg, R. R. (2003) J. Exp. Med. 198, 603– 614 75. Puri, V., Watanabe, R., Singh, R. D., Dominguez, M., Brown, J. C., Wheatley, C. L., Marks, D. L., and Pagano, R. E. (2001) J. Cell Biol. 154, 535–547 76. Nichols, B. J. (2002) Nat. Cell Biol. 4, 374 –378 77. Singh, R. D., Puri, V., Valiyaveettil, J. T., Marks, D. L., Bittman, R., and Pagano, R. E. (2003) Mol. Biol. Cell 14, 3254 –3265 78. Watson, R. T., Shigematsu, S., Chiang, S. H., Mora, S., Kanzaki, M., Macara, I. G., Saltiel, A. R., and Pessin, J. E. (2001) J. Cell Biol. 154, 829 – 840 79. Watson, R. T., Furukawa, M., Chiang, S. H., Boeglin, D., Kanzaki, M., Saltiel, A. R., and Pessin, J. E. (2003) Mol. Cell. Biol. 23, 961–974 80. Bogan, J. S., and Lodish, H. F. (1999) J. Cell Biol. 146, 609 – 620 81. Lampson, M. A., Schmoranzer, J., Zeigerer, A., Simon, S. M., and McGraw, T. E. (2001) Mol. Biol. Cell 12, 3489 –3501 82. Robinson, L. J., Pang, S., Harris, D. S., Heuser, J., and James, D. E. (1992) J. Cell Biol. 117, 1181–1196 83. Watson, R. T., Khan A. H., Furukawa, M., Hou, J. C., Li, L., Kanzaki, M., Okada, S., Kandror, K. V., and Pessin, J. E. (2004) EMBO J. 23, 2059 –2070 84. Kwik, J., Boyle, S., Fooksman, D., Margolis, L., Sheetz, M. P., and Edidin, M. (2003) Proc. Natl. Acad. Sci. U. S. A. 100, 13964 –13969 85. Keller, P., and Simons, K. (1998) J. Cell Biol. 140, 1357–1367 86. Ying, M., Grimmer, S., Iversen, T. G., Van Deurs, B., and Sandvig, K. (2003) Traffic 4, 772–784 87. Shigematsu, S., Watson, R. T., Khan, A. H., and Pessin, J. E. (2003) J. Biol. Chem. 278, 10683–10690 88. Rothberg, K. G., Heuser, J. E., Donzell, W. C., Ying, Y. S., Glenney, J. R., and Anderson, R. G. (1992) Cell 68, 673– 682

30633

89. Scherer, P. E., Lisanti, M. P., Baldini, G., Sargiacomo, M., Mastick, C. C., and Lodish, H. F. (1994) J. Cell Biol. 127, 1233–1243 90. Cantley, L. C. (2002) Science 296, 1655–1657 91. Czech, M. P., and Corvera, S. (1999) J. Biol. Chem. 274, 1865–1868 92. Hudson, A. W., Ruiz, M., and Birnbaum, M. J. (1992) J. Cell Biol. 116, 785–797 93. James, D. J., Salau¨ n, C., Brandie, F. M., Connell, J. M., and Chamberlain, L. H. (2004) J. Biol. Chem. 279, 20567–20570 94. Imamura, T., Ishibashi, K., Dalle, S., Ugi, S., and Olefsky, J. M. (1999) J. Biol. Chem. 274, 33691–33695 95. Wu-Wong, J. R., Berg, C. E., Wang, J., Chiou, W. J., and Fissel, B. (1999) J. Biol. Chem. 274, 8103– 8110 96. Yorek, M., Jaipaul, N., Dunlap, J., and Bielefeldt, K. (1999) Arch. Biochem. Biophys. 361, 241–251 97. Jouneaux, C., Mallat, A., Serradeil-Le Gal, C., Goldsmith, P., Hanoune, J., and Lotersztajn, S. (1994) J. Biol. Chem. 269, 1845–1851 98. Imamura, T., Vollenweider, P., Egawa, K., Clodi, M., Ishibashi, K., Nakashima, N., Ugi, S., Adams, J. W., Brown, J. H., and Olefsky, J. M. (1999) Mol. Cell. Biol. 19, 6765– 6774 99. Bose, A., Cherniack, A. D., Langille, S. E., Nicoloro, S. M., Buxton, J. M., Park, J. G., Chawla, A., and Czech, M. P. (2001) Mol. Cell. Biol. 21, 5262–5275 100. Lawrence, J. T., and Birnbaum, M. J. (2001) Mol. Cell. Biol. 21, 5276 –5285 101. Lawrence, J. T., and Birnbaum, M. J. (2003) Proc. Natl. Acad. Sci. U. S. A. 100, 13320 –13325 102. Aikawa, Y., and Martin, T. F. (2003) J. Cell Biol. 162, 647– 659 103. Doughman, R. L., Firestone, A. J., and Anderson, R. A. (2003) J. Membr. Biol. 194, 77– 89 104. Rameh, L. E., Tolias, K. F., Duckworth, B. C., and Cantley, L. C. (1997) Nature 390, 192–196 105. Ishihara, H., Shibasaki, Y., Kizuki, N., Wada, T., Yazaki, Y., Asano, T., and Oka, Y. (1998) J. Biol. Chem. 273, 8741– 8748 106. Weernink, P. A., Meletiadis, K., Hommeltenberg, S., Hinz, M., Ishihara, H., Schmidt, M., and Jakobs, K. H. (2004) J. Biol. Chem. 279, 7840 –7849 107. Yamamoto, M., Hilgemann, D. H., Feng, S., Bito, H., Ishihara, H., Shibasaki, Y., and Yin, H. L. (2001) J. Cell Biol. 152, 867– 876 108. Ikonen, E. (2001) Curr. Opin. Cell Biol. 13, 470 – 477 109. Bretscher, M. S., and Munro, S. (1993) Science 261, 1280 –1281 110. Kanzaki, M., and Pessin, J. E. (2002) J. Biol. Chem. 277, 25867–25869 111. Kessels, M. M., and Qualmann, B. (2002) EMBO J. 21, 6083– 6094 112. Takenawa, T., and Itoh, T. (2001) Biochim. Biophys. Acta 1533, 190 –206 113. Cremona, O., and De Camilli, P. (2001) J. Cell Sci. 114, 1041–1052 114. Martin, T. F. (2001) Curr. Opin. Cell Biol. 13, 493– 499 115. Ford, M. G., Mills, I. G., Peter, B. J., Vallis, Y., Praefcke, G. J., Evans, P. R., and McMahon, H. T. (2002) Nature 419, 361–366 116. Vallis, Y., Wigge, P., Marks, B., Evans, P. R., and McMahon, H. T. (1999) Curr. Biol. 9, 257–260 117. Barbieri, M. A., Heath, C. M., Peters, E. M., Wells, A., Davis, J. N., and Stahl, P. D. (2001) J. Biol. Chem. 276, 47212– 47216 118. Wenk, M. R., Pellegrini, L., Klenchin, V. A., Di Paolo, G., Chang, S., Daniell, L., Arioka, M., Martin, T. F., and De Camilli, P. (2001) Neuron 32, 79 – 88 119. Padron, D., Wang, Y. J., Yamamoto, M., Yin, H., and Roth, M. G. (2003) J. Cell Biol. 162, 693–701 120. Krauss, M., Kinuta, M., Wenk, M. R., De Camilli, P., Takei, K., and Haucke, V. (2003) J. Cell Biol. 162, 113–124 121. Collins, B. M., McCoy, A. J., Kent, H. M., Evans, P. R., and Owen, D. J. (2002) Cell 109, 523–535 122. Hellsten, E., Bernard, D. J., Owens, J. W., Eckhaus, M., Suchy, S. F., and Nussbaum, R. L. (2002) Biol. Reprod. 66, 1522–1530 123. Ros-Baro, A., Lopez-Iglesias, C., Piero, S., Bellido, D., Palecin, M., Zorzano, A., and Camps, M. (2001) Proc. Natl. Acad. Sci. U.S.A. 98, 12050 –12055