Phospholipase D and Its Product, Phosphatidic Acid, Mediate Agonist-dependent Raf-1 Translocation to the Plasma Membrane and the Activation of the Mitogen-activated Protein Kinase Pathway*

Mark A. RizzoDagger , Kuntala ShomeDagger , Chandrasekaran VasudevanDagger , Donna B. Stolz§, Tsung-Chang Sungparallel , Michael A. Frohmanparallel , Simon C. Watkins§, and Guillermo RomeroDagger **

From the Dagger  Department of Pharmacology, § Department of Cell Biology and Physiology, the  Center for Biological Imaging of the University of Pittsburgh, Pittsburgh, Pennsylvania 15261 and from the parallel  Department of Pharmacological Sciences and the Institute for Cell and Developmental Biology, State University of New York, Stony Brook, New York 11794

    ABSTRACT
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Abstract
Introduction
Procedures
Results
Discussion
References

The primary known function of phospholipase D (PLD) is to generate phosphatidic acid (PA) via the hydrolysis of phosphatidylcholine. However, the functional role of PA is not well understood. We report here evidence that links the activation of PLD by insulin and the subsequent generation of PA to the activation of the Raf-1-mitogen-activated protein kinase (MAPK) cascade. Brefeldin A (BFA), an inhibitor of the activation of ADP-ribosylation factor proteins, inhibited insulin-dependent production of PA and MAPK phosphorylation. The addition of PA reversed the inhibition of MAPK activation by BFA. Overexpression of a catalytically inactive variant of PLD2, but not PLD1, blocked insulin-dependent activation of PLD and phosphorylation of MAPK. Real time imaging analysis showed that insulin induced Raf-1 translocation to cell membranes by a process that was inhibited by BFA. PA addition reversed the effects of BFA on Raf-1 translocation. However, PA did not activate Raf-1 in vitro or in vivo, suggesting that the primary function of PA is to enhance the recruitment of Raf-1 to the plasma membrane where other factors may activate it. Finally, we found that the recruitment of Raf-1 to the plasma membrane was transient, but Raf-1 remained bound to endocytic vesicles.

    INTRODUCTION
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Abstract
Introduction
Procedures
Results
Discussion
References

Growth factor-mediated activation of PLD1 has been well documented and occurs in response to a broad class of mitogens, including insulin, platelet-derived growth factor, epidermal growth factor, vasopressin, and phorbol esters (1-4). Activation of PLD occurs through interaction with the small G-proteins of the ADP-ribosylation factor (ARF) (5, 6) and Rac/Rho families (7) as well as with protein kinase C (PKC) (8, 9). The relative contribution of these factors to the activation of PLD is highly dependent on the cell type and signaling model examined. For example, stimulation of Rat-1 fibroblasts overexpressing the human insulin receptor (HIRcB cells) with insulin activates PLD exclusively through the ARF pathway (10), whereas the activation of PLD by insulin in adipocytes appears to be primarily Rho-mediated (11). Activation of PLD has been implicated in a wide variety of intracellular and extracellular processes, including actin polymerization, coatomer assembly, vesicle transport, neutrophil activation, and platelet aggregation (12-16).

Activated PLD catalyzes the hydrolysis of phosphatidylcholine to generate PA. However, the downstream consequences of PA generation are not well understood. Although it is clear that the principal effects of PA in some systems may be mediated by its conversion to diacylglycerol (DAG) or lysophosphatidic acid (LPA), PA may also be a potent second messenger. Several laboratories have identified putative targets for PA in growth factor signal transduction, including a protein tyrosine phosphatase (17), phospholipase C-gamma (18), and Ras-GAP (19). However, the physiological relevance of these interactions has not been established.

Recently, Ghosh et al. (20) reported that PA interacts directly with the serine-threonine kinase Raf-1, an important component of the MAPK signaling cascade. Here we report that the stimulation of the MAPK pathway by insulin is dependent on PLD activation, and this effect is mediated through the induction of Raf-1 translocation to the plasma membrane by PA. Furthermore, overexpression of a catalytically inactive variant of PLD2, but not PLD1, blocks insulin-induced phosphorylation of MAPK. We also show that PA is required for complete activation of Raf-1 in response to insulin. However, PA alone cannot activate Raf kinase in vivo, does not have any effect on Raf kinase activity in vitro, and has no effects on the MAPK cascade. We also show that PA induces Raf-1 translocation to the plasma membrane and that the generation of PA is essential for the induction of Raf-1 translocation by insulin. PA also induced Raf-1 translocation to the plasma membrane in Ha-Ras(Q61L)-transformed Rat-1 fibroblasts, suggesting that PA may act concurrently with activated Ras in stimulating Raf-1 translocation. Raf-1 was found associated with intracellular vesicles containing the insulin receptor and clathrin after stimulation with insulin. Furthermore, Raf-1 association to endocytic vesicles was dependent on the generation of PA, suggesting a model in which Raf-1 migrates along with endocytic vesicles during receptor-mediated endocytosis via its interaction with PA.

    EXPERIMENTAL PROCEDURES
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Introduction
Procedures
Results
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References

Cell Culture-- Rat-1 fibroblasts overexpressing the human insulin receptors (HIRcB cells) were grown in Dulbecco's modified Eagle's medium/Ham's F-12 (1:1) supplemented with 10% fetal bovine serum and 100 nM methotrexate. Cells were grown to 80% confluency and serum-starved for at least 18 h prior to stimulation. Cells were treated with 200 nM insulin and/or 200 µM PA for 10 min. When indicated, cells were pretreated with brefeldin A (BFA) for 10 min prior to stimulation with insulin or PA. Ha-Ras(Q61L)-transformed Rat-1 fibroblasts were treated as described above.

Phospholipase D Activity-- HIRcB cells were labeled with [3H]palmitate (5 µCi/ml) overnight and treated with insulin as described above. The reaction was stopped after 1, 2, 5, 10, 15, and 20 min of insulin stimulation by the addition of cold PBS. Cells were then scraped in cold PBS and pelleted by centrifugation. Lipids were then extracted with chloroform/methanol (1:1). The lipid phase was collected and developed by thin layer chromatography (TLC) on silica gel 60 plates using ethyl acetate/trimethylpentane/acetic acid (9:5:2) as the solvent. The position of PA was measured by autoradiography and by the position of lipid standards (Avanti Polar Lipids). Lipids were then scraped from the TLC plates and counted via liquid scintillation. In other assays, cells were stimulated with insulin in the presence of 0.3% butanol for 20 min to determine the total activity of PLD by the standard transphosphatidylation assay described by Shome et al. (10). Levels of PA or phosphatidylbutanol were normalized to total fatty acid label incorporated into lipid.

Assessment of MAPK Phosphorylation-- HIRcB cells were treated as described above or as described in the figure legends. Cells were washed with cold PBS, scraped into microcentrifuge tubes in Buffer A (10 mM Hepes, pH 7.4, 2 mM EDTA, 1 mM Na3VO4, and 1 mM phenylmethylsulfonyl fluoride), and centrifuged. Cells were resuspended in a 0.5-ml detergent lysis buffer (50 mM Hepes, pH 7.4, 0.1 M NaCl, 1.5% sodium cholate, 1 mM EDTA, 1 mM EGTA, 5 µg/ml leupeptin, 1 mM phenylmethylsulfonyl fluoride, and 1 mg/ml soybean trypsin inhibitor) and lysed for 30 min at 4 °C. Cell lysates were then boiled in SDS sample buffer, and equal amounts of protein were resolved by SDS-PAGE followed by transfer to a nitrocellulose membrane. The nitrocellulose membrane was then probed with a phospho-specific anti-MAPK antibody (New England Biolabs). Immunocomplexes were detected by enhanced chemiluminescence.

Phospholipase D Mutants-- Wild-type and catalytically inactive variants of PLD1 and PLD2 (K898R-PLD1 and K758R-PLD2) were made as described previously (21) and fused to Aequorea victoria green fluorescent protein by subcloning into pEFGP-C1 (CLONTECH). The enzymatic activity of the GFP-wild-type enzyme chimeras expressed in COS-7 cells was determined in vitro to confirm that the GFP tag did not generate an inactive phenotype.2 The structure of the chimeras was verified directly by sequencing. Cells were transfected using LipofectAMINE transfection reagent. Transfection efficiency was assessed by conventional epifluorescence microscopy prior to experimentation using filters appropriate for the detection of GFP.

Construction of Raf-GFP-- A pUC13 expression vector encoding the human Raf-1 cDNA was a gift from Dr. Said Sebti. The human Raf-1 cDNA was amplified by polymerase chain reaction and subcloned into pEGFP-N1 (CLONTECH) using the EcoRI and BamHI restriction sites, successfully fusing GFP to the C terminus of Raf-1. Escherichia coli (DH5-alpha ) were transformed with the ligated vector and selected using kanamycin-containing LB agar plates. Plasmid DNA was obtained from transformed bacteria. The presence of the fusion construct was verified by digestion with restriction enzymes and direct sequencing.

Raf Kinase Assay-- HIRcB cells were transfected with Raf-1-GFP and treated as described above. After stimulation, the cells were washed with ice-cold PBS, scraped in 0.5 ml of Buffer A, and centrifuged. Cell pellets were then resuspended in 0.5 ml of detergent lysis buffer and lysed for 30 min on ice. 3 µl of monoclonal anti-GFP antibody (CLONTECH) were conjugated to 30 µl of agarose-anti-mouse IgG (Sigma) for 1 h at room temperature in detergent lysis buffer and washed twice. Cells lysates were incubated with agarose-conjugated antibody for 3 h at 4 °C with end-over-end rotation. Immunocomplexes were pelleted (10 min at 10,000 × g) and washed 3 times with detergent lysis buffer and 2 times with kinase buffer (50 mM Tris, pH 7.3, 150 mM NaCl, 12 mM MnCl2, 1 mM dithiothreitol, 0.2% Tween 20). The activity of Raf-1 kinase present in Raf-GFP immunoprecipitates was determined using a standard procedure (22-24). To test whether PA activated Raf-GFP in vitro, 200 µM PA was included in the reaction buffer where indicated. Following the kinase assay, immunocomplexes were boiled in SDS sample buffer and resolved on a 10% acrylamide gel followed by transfer to a nitrocellulose membrane. Membranes were probed with an anti-Raf-1 antibody (Transduction Laboratories) and resolved via an appropriate horseradish peroxidase-conjugated secondary antibody and ECL. Kinase activity was normalized to the relative amounts of Raf-GFP in the immunoprecipitate as determined by densitometry.

Subcellular Fractionation-- HIRcB cells were transfected with the plasmid encoding Raf-GFP using LipofectAMINE (Life Technologies, Inc.). Cells were stimulated as described above and were scraped in 0.5 ml of Buffer A, pelleted by centrifugation, and resuspended in Buffer B (Buffer A + 50 mM NaF, 1 µg/ml aprotinin, and 10 µg/ml leupeptin). Cells were lysed by sonication with 3 series of 5 × 5-s bursts using a microtip sonicator. The nuclei and cell debris were subsequently pelleted (3,000 RPM, 10 min, 4 °C). The remaining supernatant fraction was spun at 100,000 × g for 80 min at 4 °C to separate membrane and cytosolic fractions. Membrane pellets were solubilized in Buffer B containing 1% Triton X-100 and incubated for 30 min at 4 °C. Supernatant and pellet fractions were boiled in Laemmli sample buffer, and equal amounts of protein were run on a 10% acrylamide gel followed by transfer to nitrocellulose. Membranes were probed with an anti-Raf-1 antibody (Transduction Laboratories). Immunocomplexes were then detected chemiluminescence. Densitometry was performed as described above.

Fluorescence Microscopy-- HIRcB cells or Rat-1 fibroblasts transformed with Ha-Ras(Q61L) (25) were plated on poly-L-lysine-coated glass coverslips and transfected with the plasmid encoding Raf-GFP using Superfect Transfection Reagent (Qiagen) as per the manufacturer's instructions. Cells were stimulated as described above. Live cells were imaged at 37 °C using a Molecular Dynamics 2001 laser scanning confocal microscope equipped with a 60× oil immersion objective and a microperfusion incubator (Medical Systems Corp.) attached to the microscope stage. Samples were excited at a wavelength of 488 nm, and emitted light was filtered through a 530DF30 double bandpass filter prior to detection with a photomultiplier tube. Quantitation of fluorescence intensity was performed with Molecular Dynamics ImageSpace software.

Internalization Assay-- HIRcB cells were stimulated as described above and treated as described in Shome et al. (26). Briefly, stimulated cells were washed with PBS and Dulbecco's modified Eagle's medium, 0.1% bovine serum albumin (pH 4.0) at 4 °C and incubated with 1 mg/ml trypsin in PBS for 30 min at 4 °C to cleave the alpha -subunit of insulin receptors on the cell surface. Trypsin treatment was stopped with buffer containing 0.4% Triton X-100, 0.5 mg/ml bacitracin, 25 mg/ml soybean trypsin inhibitor, and 100 µg/ml leupeptin and solubilized for 30 min at 4 °C. Antibody 83.7, a monoclonal antibody raised against the alpha -subunit of the insulin receptor (27), was conjugated to agarose-anti-mouse IgG as described above and incubated with cell lysates and used to immunoprecipitate internalized (i.e. intact, not exposed to trypsin hydrolysis) insulin receptors as described by Shome et al. (26). Immunoprecipitates were extensively washed as described above, resolved by SDS-PAGE, and immunoblotted with CT-1, a monoclonal antibody that recognizes the beta -subunit of the insulin receptor (27).

Immunoisolation of Vesicles-- Cells were treated as described above with BFA, insulin, and/or PA. After stimulation, cells were placed on ice and washed with cold PBS. Cells were scraped into Buffer A and pelleted. Cells obtained from two 100-mm dishes were combined and resuspended in 1 ml of Buffer C (Buffer A + 5% sucrose, 1 µg/ml aprotinin, and 10 µg/ml leupeptin) and homogenized with 4 passes through a ball homogenizer as described by Martin (28). Unbroken cells and nuclei were pelleted by centrifugation at 3000 rpm for 10 min (4 °C). Equal amounts of protein were layered on a 35% sucrose solution (in Buffer A) and centrifuged for 90 min at 150,000 × g (4 °C) in an SW55 TI swing bucket rotor. Following centrifugation, the vesicle fraction was collected at the interface of the two layers. Antibody CT-1 was conjugated to agarose-anti-mouse IgG as described above and incubated with the vesicle fractions in order to isolate internalized vesicles containing the insulin receptor. Following immunoprecipitation for 3 h at 4 °C, immunoprecipitates were washed 3× in cold Buffer C and 2× in cold PBS. The proteins of interest (Clathrin, Raf-1, and the insulin receptor) were resolved via SDS-PAGE and detected with specific antibodies.

Cell Preparation for Immunoelectron Microscopy-- Cells were grown to confluence in 100-mm dishes, treated as described above with insulin and PA, and immediately fixed for 1 h in 2% paraformaldehyde, 0.01% glutaraldehyde in PBS. The cells were scraped from the Petri dish, spun, and resuspended in 2% gelatin (300 bloom), fixed for 10 min in the same fix as above, and cryoprotected overnight in 2.3 M sucrose in 0.1 M PBS. Subsequently, the cell pellet was diced into 1 mm3, mounted on cutting stubs, shock-frozen, and stored in liquid nitrogen. Thin sections (70-100 nm) were cut using a Reichert ultracut S Ultramicrotome with a FC4S cryo-attachment and lifted in a small drop of sucrose and mounted on Formvar-coated carbon grids. The sections were washed three times in PBS containing 0.5% bovine serum albumin and 0.15% glycine, pH 7.4 (Buffer D), followed by a 30-min incubation with purified goat IgG (50 mg/ml) at 25 °C and three additional washes with buffer. All the preceding steps are designed to ensure minimal nonspecific reaction to the antibodies used. Sections were then incubated for 60 min in the first primary antibody, a murine IgG1 directed against Raf-1 (Transduction Laboratories) followed by three washes and a 60-min incubation with 5 nm of gold-labeled protein G (0.1-2 µg/ml). The sections were then washed six times (5 min/wash), and the second primary antibody, a rabbit polyclonal IgG directed against the alpha -subunit of the insulin receptor (Santa Cruz Biotechnology), was applied and followed by a similar washing and labeling strategy as above, although detection was with a 10-nm protein A-gold conjugate. Finally, sections were washed thoroughly in Buffer D (5 changes) and in PBS (3 changes), followed by a brief fixation step in 2.5% glutaraldehyde in PBS. Subsequent steps were 3 further washes in PBS, 5 washes in water, counterstaining with uranyl acetate and embedment in 1.25% methylcellulose. Observation was with a Jeol 1210CX electron microscope.

    RESULTS
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Procedures
Results
Discussion
References

The Generation of PA by PLD Is Required for Receptor-induced MAPK Phosphorylation-- In order to investigate the functional role of PLD activation in insulin signaling, we used a Rat-1 fibroblast cell line that overexpresses the human insulin receptor (HIRcB cells). Insulin-induced activation of PLD in these cells appears to be exclusively dependent on ARF activation (10). We have shown that BFA, an inhibitor of ARF activation (29-31), blocks completely insulin-dependent PLD activation in these cells (10).

Fig. 1A shows the effects of insulin on the generation of PA by HIRcB cells. Also shown in the figure is the blockade of the generation of PA by treatment with BFA. To assess the physiological relevance of insulin-induced PA production, we examined its effect on the MAPK signaling pathway. Activation of this cascade is characterized by a series of phosphorylation events, one of which is the phosphorylation of MAPK. We therefore used an antibody specific for phosphorylated MAPK as a marker for the activation of this signaling cascade (Fig. 1B). As shown, insulin induced the phosphorylation of MAPK. The effect of insulin was inhibited by pretreatment with BFA at doses consistent with the inhibition of PLD activity (Fig. 1A (10)). Addition of PA had no effects on the phosphorylation of MAPK. However, the addition of PA reversed the inhibitory effects of BFA on insulin-induced MAPK phosphorylation.


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Fig. 1.   The effects of BFA on insulin-induced PA generation and MAPK phosphorylation. A, HIRcB cells were labeled with [3H]palmitate and stimulated with 200 nM insulin (black-square) or pretreated with 50 µg/ml BFA 10 min prior to stimulation with insulin (black-triangle). Insulin was added at 0 min, and the incubation was stopped at the time points indicated. Lipids were then extracted, separated by TLC, and counted via liquid scintillation. The percentage of PA was calculated as the percentage of total counts of PA compared with the total counts of lipid isolated. Results shown represent at least three separate experiments. B, HIRcB cells were pretreated for 10 min with a dose of BFA as indicated prior to stimulation 200 nM insulin or 200 µM PA for 10 min. Cell lysates were resolved via SDS-PAGE and Western blot-probed with phospho-specific anti-MAPK antibodies.

The role of PLD in the insulin-induced phosphorylation of MAPK was further examined using cells that had been transiently transfected with catalytically inactive mutants of PLD1 and PLD2. These variants contain a conservative Lys to Arg mutation in the active site, effectively disrupting the ability of PLD to hydrolyze phosphatidylcholine. These mutants have been shown to be devoid of activity in vivo and in vitro using PC as a substrate and to localize similarly to the wild-type proteins (21). In these assays, GFP-PLD1 and GFP-PLD2 constructs were used to transfect HIRcB cells. These chimeric proteins were used instead of the original proteins because the expression of the GFP constructs facilitated significantly the analysis of the transfection efficiency. Thus, we verified the expression of these constructs by conventional epifluorescence analysis prior to experimentation. A transfection efficiency of approximately 60% was obtained for each of the constructs (data not shown). As shown in Fig. 2A, overexpression of GFP-tagged constructs of the catalytically inactive K758R-PLD2 mutant in HIRcB blocked insulin-induced PLD activity, demonstrating the ability of this mutant to act as a dominant negative. Furthermore, expression of the K758R-PLD2 also blocked insulin-induced MAPK activation (Fig. 2B). In contrast, overexpression of the catalytically inactive K898R-PLD1 mutant failed to inhibit insulin-dependent PLD activity and insulin-induced MAPK phosphorylation in HIRcB cells (Fig. 2, A and B). This demonstrates that, in HIRcB cells, insulin signals preferentially through PLD2 and that a functional PLD2 is required for insulin-induced activation of MAPK. Taken altogether, these data lead us to conclude that the generation of PA by PLD2 is essential, but not sufficient, to mediate the effects of insulin on the MAPK signaling cascade.


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Fig. 2.   The effect of catalytically inactive PLD mutants on insulin-dependent PLD activity and MAPK phosphorylation. HIRcB cells were transfected with plasmids encoding EGFP (Control) or the catalytically inactive GFP-PLD mutants (K898R-PLD1; K758R-PLD2). Expression of these constructs was verified by conventional epifluorescence prior to experimentation. A, cells were serum-starved and labeled overnight with [3H]palmitate. PLD activity was then measured as described under "Experimental Procedures" in cells stimulated with 200 nM insulin (black bars) or without any stimulation (white bars). The data are expressed as the number of counts obtained from the phosphatidylbutanol (PtdBu) spot-normalized by the total counts of lipid. Results shown represent at least three separate experiments. Samples denoted with * were significantly different as determined by analysis of variance Bonferroni multiple comparisons test (p < 0.05). B, cells treated with and without 200 nM insulin were lysed and resolved via SDS-PAGE and Western blot. Nitrocellulose membranes were probed with phospho-specific anti-MAPK antibodies and resolved as described above.

PA Induces Raf-GFP Translocation, but Does Not Activate Raf Kinase Activity-- It has been shown that Raf-1 binds PA in vitro (20). Therefore, one putative site of action for PA in the MAPK signaling cascade is the activation of Raf-1. To assess the role of PA on the regulation of Raf-1 activity, we constructed a Raf-GFP fusion protein and transiently transfected HIRcB cells. In order to demonstrate that the fusion protein behaved as the native kinase, enzymatic assays were performed on materials immunoprecipitated with an antibody specific for the GFP epitope. Treatment of HIRcB cells with insulin induced a 2-fold activation of Raf kinase activity, whereas treatment with PA did not have statistically significant effects on Raf kinase activity (Fig. 3A). As predicted by our hypothesis, the activation of Raf by insulin was inhibited by BFA. This inhibition was reversed by the addition of PA, suggesting that the generation of PA by PLD is necessary for Raf-1 activation and that PA alone cannot activate Raf-1. Because Ghosh et al. (20) showed a direct interaction between Raf-1 and PA, we tested the ability of PA to activate directly immunoprecipitated Raf-GFP. As shown, PA was unable to stimulate Raf kinase activity in immunoprecipitated Raf-GFP. Interestingly, some endogenous Raf-1 co-immunoprecipitated with Raf-GFP, suggesting that Raf-GFP can oligomerize with endogenous Raf-1 in vivo (data not shown). This supports the idea that the Raf-GFP construct is physiologically active.


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Fig. 3.   The role of PLD-generated PA on insulin-induced Raf-GFP kinase activity and translocation. In order to assess the role of PLD-generated PA on Raf-1 kinase activity, HIRcB cells were transiently transfected with a plasmid encoding the Raf-GFP construct. A, transfected cells were stimulated with 200 nM insulin (Ins), 200 µM PA, and/or 50 µg/ml BFA as indicated. Kinase activity of immunoprecipitated Raf-GFP was assessed as described under "Experimental Procedures." In order to test the ability of PA to activate directly Raf-1 in vivo, immunoprecipitates (Ippt) from serum-starved cells were treated with 200 µM PA prior to the kinase assay. Results shown represent at least 3 separate experiments. Samples denoted with * were significantly different as determined by analysis of variance Bonferroni multiple comparisons test (p < 0.01). B, HIRcB cells transiently transfected with the Raf-GFP construct were stimulated as shown and fractionated by centrifugation at 100,000 × g for 80 min as described under "Experimental Procedures." The membrane pellet and supernatant fractions were analyzed by SDS-PAGE and immunoblotting with an antibody raised against Raf-1. The pellet fraction was quantitated by densitometry, and the optical density of the bands was plotted as shown.

The effects of anionic phospholipid-protein interactions, such as those between phosphatidylinositol 4,5-bisphosphate (PIP2) and proteins containing pleckstrin homology domains, have been shown to stimulate the recruitment of proteins to cellular membranes with or without stimulation of catalytic activity. Since activation of Raf-1 in vivo is highly dependent upon the translocation of Raf-1 to the plasma membrane, we examined the relationship between PA-induced Raf-1 translocation and the activation of Raf-1. In these experiments, we fractionated cells expressing GFP-tagged Raf-1 by centrifugation and separated cytosolic (supernatant) and membrane (pellet) fractions. PA alone was sufficient to induce Raf-GFP translocation from the supernatant to the pellet fraction (Fig. 3B). Insulin-induced Raf-GFP translocation was inhibited by BFA in a dose-dependent manner consistent with the inhibition of PLD. Addition of PA to cells treated with BFA and insulin restored Raf-GFP translocation into the pellet fraction, efficiently reversing the effects of BFA. PA was also found to be at least as efficient as insulin in the recruitment of Raf-1 to cell membranes. In contrast, as shown above, PA failed to stimulate Raf kinase activity. This suggests the hypothesis that the recruitment of Raf to cell membranes is essential but not sufficient to completely activate the kinase.

Live Cell Dynamics of Raf-GFP Translocation-- Raf-GFP translocation in live cells was studied using a Molecular Dynamics confocal microscope equipped with a microperfusion incubator to maintain a constant temperature of 37 °C. Confocal sections of the plasma membrane adjacent to the coverslip in HIRcB cells expressing Raf-GFP were taken in order to track Raf-GFP translocation to the plasma membrane. Cells were imaged every 1 min following the addition of insulin or PA. Representative plasma membrane sections of a cell stimulated with insulin are shown in Fig. 4A. Total intensity of the section was then quantitated using Molecular Dynamics ImageSpace software, and the results were normalized for background intensity. Insulin induced a transient Raf-1 translocation to the plasma membrane peaking at approximately 10 min (Fig. 4B).


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Fig. 4.   Live cell dynamics of insulin-induced Raf-GFP translocation. HIRcB cells transiently transfected with Raf-GFP were imaged using fluorescent confocal microscopy at 37 °C. A, a confocal section of the plasma membrane adjacent to the coverslip was obtained by scanning optical sections along the z axis of the cell. The plasma membrane adjacent to the coverslip was identified by choosing a z section below the nucleus of the cell. This allows quantitation of Raf-GFP translocation from the cytoplasm (out of focus sections) to the plasma membrane (confocal section in focus). Cells were imaged at 1-min intervals for 20 min following the addition of 200 nM insulin. Representative sections are shown. B, fluorescence intensity of the plasma membrane sections as shown in A was quantitated using Molecular Dynamics ImageSpace software and plotted versus time after the addition of insulin. The arrow indicates point that insulin (200 nM) was administered. Results were representative of at least 4 separate experiments. C, HIRcB cells imaged as in A were pretreated with BFA (50 µg/ml) and stimulated with PA (200 µM) and insulin (200 nM) together (gray lines) or separately (black lines). Arrows indicate addition of agonists.

Pretreatment of HIRcB cells with BFA did not alter endogenous levels of membrane-bound Raf-GFP but blocked insulin-induced Raf-GFP translocation (Fig. 4C, black line). However, the addition of PA restored Raf-1 translocation to cells treated with BFA and stimulated with insulin. Furthermore, Raf-GFP translocation was dependent on the addition of PA. When PA was included along with insulin, Raf-GFP translocation occurred immediately (Fig. 4C, gray line). PA alone was also found to induce Raf-1 translocation, and this effect was insensitive to the addition of BFA (data not shown).

PA Induces Raf-GFP Translocation in Ras-transformed Cells-- Raf-1 translocation to the plasma membrane has been previously shown to be stimulated by the activation of the small G-protein Ras. Ras is constitutively bound to the plasma membrane and binds Raf-1 upon the exchange of GDP for GTP. In order to examine whether PA-induced translocation of Raf-1 can occur in the presence of activated Ras, we imaged confocal plasma membrane sections in Rat-1 cells transformed with the GTPase-deficient (constitutively active) Ha-Ras(Q61L) (25). PA was found to induce rapid and transient Raf-GFP in these cells (Fig. 5). Levels of Raf-GFP at the plasma membrane were sustained until approximately 8 min and decreased to control levels by approximately 12 min after stimulation with PA. Thus, PA also stimulated the transient recruitment of Raf-1 to the plasma membrane in Ras-transformed cells.


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Fig. 5.   PA induces transient Raf-GFP translocation in Ha-ras(Q61L)-transformed Rat-1 fibroblasts. In order to test the ability of PA to induce Raf-1 translocation in the presence of activated Ras, Ha-Ras(Q61L)-transformed Rat-1 fibroblasts were transiently transfected with the Raf-GFP construct, and a confocal section of the plasma membrane was imaged as described in Fig. 3. Cells were treated with 200 µM PA and imaged at 1-min intervals for 20 min. Sections shown are representative of three separate trials.

Insulin and PA Induce Raf-1 Translocation to Endocytic Vesicles-- In mid-cell confocal sections of insulin and PA-treated HIRcB cells, it was noted that Raf-1 accumulated in vesicular intracellular structures in addition to the plasma membrane. Interestingly, BFA did not inhibit PA-induced translocation to the plasma membrane or intracellular structures, whereas the effects of insulin were sensitive to BFA. PA has been implicated in the generation of transport vesicles from the endoplasmic reticulum and Golgi (12, 32), and recent work by Chung et al. (33) has further suggested that PLD-generated PA promotes formation of endocytic vesicles and vesicle coat assembly via a positive feedback mechanism with PIP2 biosynthesis. These findings suggested to us the hypothesis that the generation of PA is important for receptor-mediated endocytosis and formation of endocytic vesicles and that Raf-1 leaves the plasma membrane while still bound to an endocytic vesicle via its association with PA.

To study the relationship between PA production and endocytosis, we first examined the effects of BFA on the internalization of the insulin receptor. The internalization of the insulin receptor was measured using a limited trypsin proteolysis assay described previously (26). Pretreatment of cells with BFA blocked insulin-induced internalization of the insulin (Fig. 6A). This is consistent with a role of ARF proteins and PA production in the formation of the endocytic vesicle (33). To confirm that the internalization of the insulin receptor proceeded via the classic endocytic pathway, we immunoisolated vesicles using a specific antibody that recognizes the C terminus of the beta -subunit of the insulin receptor (27). As shown, these vesicles also contained clathrin heavy chains (Fig. 6B). Also shown is the effect of BFA, which reduced substantially the co-immunoprecipitation of the clathrin heavy chain with the insulin receptor-containing vesicles (Fig. 6B). When these vesicles were examined for the presence of Raf-1 a consistent pattern emerged: very little Raf-1 was associated to vesicles isolated from untreated cells, whereas a very significant amount of Raf-1 was found in the vesicles obtained from cells that had been exposed to insulin (Fig. 6B). In consistency with the data in Fig. 6, A and B, a short pretreatment with BFA abolished the co-localization of Raf-1 with clathrin and the insulin receptor.


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Fig. 6.   A, BFA inhibits the internalization of the insulin receptor. HIRcB cells treated as described were trypsinized after stimulation for 30 min at 4 °C in order to cleave the alpha -subunit insulin (Ins) receptors localized at the cell surface. Cleaved receptors were separated from intact, internalized receptors by immunoprecipitation with an antibody raised to the N terminus of the alpha -subunit of the insulin receptor. The immunoprecipitated receptors were detected using a specific antibody against the beta -subunit. Western blots of immunoprecipitates detecting the beta -subunit of the insulin receptor were quantitated via densitometry and Molecular Dynamics ImageQuant software. The intensity of the bands is directly proportional to the number of intact, internalized receptors. Samples denoted with * were significantly different as determined by analysis of variance Bonferroni multiple comparisons test (p < 0.01). B, insulin induces co-localization of Raf-1 and clathrin in vesicles containing the insulin receptor. Vesicles were isolated from cells following stimulation as indicated by sucrose gradient purification of cell lysates. Vesicles containing the insulin receptor were then immunoisolated with an antibody recognizing the C terminus of the beta -subunit of the insulin receptor as described under "Experimental Procedures." immunoprecipitates were resolved via SDS-PAGE and Western blot. Nitrocellulose membranes were probed for Raf-1 and clathrin heavy chain as indicated.

Ultrastructural data using dual staining electron microscopy against the insulin receptor and Raf-1 epitopes in insulin- and PA-treated HIRcB cells were obtained to confirm the co-localization of Raf-1 and the insulin receptor in endocytic vesicles. Both insulin (Fig. 7, B and C) and PA (Fig. 7D) induced co-localization of Raf-1 with the insulin receptor in endosomes, indicating that Raf-1 migrates along with endocytic vesicles in response to stimulation with insulin or PA. Therefore, both the internalization of the insulin receptor and the recruitment of Raf-1 to the plasma membrane are dependent on the generation of PA. Furthermore, PA alone can induce the internalization of insulin receptors and recruitment of Raf-1 to vesicles containing insulin receptors, suggesting that Raf-1 is bound to the membranes of endocytic vesicles via its association with PA.


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Fig. 7.   Raf-1 co-localizes with the insulin receptor in endocytic vesicles. HIRcB cells were left untreated (A) or stimulated with 200 nM insulin (B and C) or 200 µM PA (D). Cells were then fixed and processed for frozen sectioning prior to labeling for Raf-1 (5 nm gold particles, small arrow in A) and the alpha -subunit of the insulin receptor (10 nm gold particles, large arrow in A). Arrows in B-D show vesicles that contain both insulin receptors and Raf-1, indicating that Raf-1 is internalized with endocytic vesicles. Bar, 200 nm.


    DISCUSSION
Top
Abstract
Introduction
Procedures
Results
Discussion
References

The activation of Raf-1 kinase activity by growth factors requires its translocation from the cytoplasm to the plasma membrane where it is activated through a complex mechanism which includes the interaction with the GTP-bound form of Ras, and possibly phosphorylation by PKC, and tyrosine kinases (34-36). Whereas the precise nature of the events occurring at the plasma membrane remains unresolved, it is clear that the translocation of Raf-1 is crucial. Targeting Raf-1 to the plasma membrane by attaching a protein prenylation motif to the C terminus of Raf-1 is sufficient for activation of the kinase (37), whereas trapping Raf-1 in the cytoplasm with cytosolic Ras prevents activation (22). However, translocation itself does not bring about the full activation of Raf-1. Mineo et al. (38) used a mutant Ras protein that is deficient in binding to wild-type Raf-1, but binds Raf-1(257L), to show that the interaction between Ras and Raf-1 stimulates Raf-1 kinase activity 3-fold better than targeting Raf-1 to the membrane alone. Furthermore, Roy et al. (39) showed that recruitment of Raf-1 to the plasma membrane by Ras was not sufficient for full activation of Raf-1 and that a second interaction between the cysteine-rich domain (CRD) on Raf-1 and the GTP-bound form of Ras was necessary for full activation. They also showed that deletion of the CRD from membrane-targeted Raf-1 abrogated Raf-1 kinase activity, suggesting that plasma membrane localization of Raf-1 by itself is insufficient for activation of Raf-1 but that a second regulatory event affecting the CRD must occur for Raf-1 activation. Thus, Raf-1 translocation and Raf-1 kinase activation are closely related but distinct phenomena.

It has been assumed for some time that the interactions of Raf-1 with Ras are the primary mechanism driving the recruitment of Raf-1 to the cell membrane. Recently, other mechanisms that may play an important role in the recruitment of Raf-1 to the membrane have been investigated. For instance, Ghosh et al. (20) have explored the interactions of Raf-1 with phosphatidylserine and PA in vitro. Phosphatidylserine appears to bind to the cysteine-rich domain (CRD) of Raf-1, which is analogous to the zinc finger on PKC. Luo et al. (40) replaced the Raf-1 CRD with the analogous zinc finger domain found on PKC and found that DAG activated this chimera independently of Ras activation, demonstrating that interaction of an effector with the CRD is critical in the activation of Raf-1. Other effectors, such as ceramide (41) and Rap1A (42), interact with Raf-1 at this site and consequently have effects on its activation. The PA-binding site proposed by Ghosh et al. (20) does not lie in this crucial lipid binding regulatory domain on Raf-1 but on a second lipid-binding site near the catalytic domain of Raf-1. The influence of effector binding at this site on Raf-1 kinase activity, if any, has not been fully characterized at the present time. It has also been shown that inhibition of PLD-mediated generation of PA with ethanol inhibited phorbol ester-induced Raf-1 translocation to cell membranes (20). This suggests that the generation of PA by a receptor-sensitive PLD may play an important role in the recruitment and/or activation of Raf-1 kinase.

The data reported here strongly support this view. By taking advantage of the fact that insulin-dependent PLD activation in HIRcB cells is mediated by ARF proteins in a BFA-sensitive manner (10), we have shown that the blockade of PLD-dependent generation of PA disrupts the activation of Raf-1, the translocation of Raf-1 to membranes, and the phosphorylation of MAPK. We also demonstrated that overexpression of a catalytically inactive variant of PLD2 blocks insulin-induced activation of PLD and MAPK phosphorylation. In consistency with the hypothesis that the effects of BFA are a consequence of the blockade of the generation of PA by receptor-sensitive PLD, we have also shown that all the effects of BFA can be reversed by the addition of exogenous PA. However, our data show that PA alone cannot activate MAPK phosphorylation in live cells and that it cannot activate Raf-1 in vitro or in cultured cells. Taking all these data together, we conclude that the generation of PA is required but not sufficient for the activation of Raf-1 by insulin.

On the other hand, we show here that PA is sufficient for induction of Raf-1 translocation and reverses the blockade of insulin-induced Raf-1 translocation by BFA to intracellular vesicles. These facts suggest a model in which PA directly facilitates Raf-1 translocation but does not activate the kinase and is insufficient to completely stimulate Raf-1 kinase activity in intact cells. Furthermore, we show that PA induces Raf-1 translocation in Ha-Ras(Q61L)-transformed cells, suggesting that PA and Ras may act concurrently and by parallel pathways in stimulating Raf-1 translocation to the plasma membrane. We therefore conclude that the main role of PA in the activation of the MAPK cascade is the induction of Raf-1 translocation to the cell membrane.

Much of the attention on receptor-sensitive PLD has focused on PLD1, primarily because recombinant PLD1 may be activated by ARF, Rho, and PKCalpha in vitro (43-45). In vitro studies on recombinant PLD2, on the other hand, have demonstrated that purified PLD2 has a high basal activity that is largely insensitive to ARF and Rho (44, 46) and thus was thought to be an unlikely candidate for the receptor-sensitive PLD activity. However, our findings suggest that in HIRcB cells PLD2 is the main species involved in insulin-dependent PLD signaling. This conclusion is based on the fact that a catalytically inactive variant of PLD2 functions as a dominant negative and blocks insulin-induced phosphorylation of MAPK, whereas a catalytically inactive PLD1 does not. Interestingly, this further suggests that PLD2 is regulated by ARF in vivo, since the insulin-dependent PLD activity in HIRcB cells requires ARF activation (10). Recent evidence from the Sung et al.3 agrees with this model. Although immunopurified PLD2 was found to be largely unresponsive to ARF, a preparation of crude membranes containing PLD2 overexpressed in COS-7 cells was activated by ARF preloaded with GTPgamma S, suggesting that PLD2 may be regulated by ARF in vivo. Furthermore, a PLD2 mutant lacking the N-terminal 308 amino acids displays both reduced in vitro and in vivo basal activity and is stimulated more than 10-fold by ARF. These results are consistent with a model for ARF-mediated PLD2 activation in response to insulin activation.

Our data do not rule out the possibility that PA-stimulated Raf-1 translocation is mediated by metabolites of PA, specifically DAG or LPA. However, we do not believe this to be the case. DAG-mediated activation of PKC results in potent activation of Raf-1 (48), and thus a significant conversion of PA to DAG would result in MAPK activation as well as potent activation of Raf-1. Since treatment of cells with PA alone failed to activate either Raf-1 or MAPK, we conclude that the generation of DAG does not play a significant role in the mechanism by which PA modulates the MAPK cascade. Likewise, a significant accumulation of LPA would also elicit a strong activation of the MAPK cascade (49). Therefore, conversion of PA to either LPA or DAG should have dramatic effects on the Raf-1-MAPK signaling cascade. Since these effects were not seen after the addition of PA in our experiments, it is likely that the effects of PA on Raf-1 translocation are due to a direct interaction between PA and Raf-1 and not a consequence of its conversion to other lipid second messengers.

Previously, immunocytochemical characterization of Raf-1 translocation has been limited to a few studies in which Raf-1 was microinjected into Ras-transformed cells (37, 50, 51). However, little work has been done to characterize growth factor-induced Raf-1 translocation. Here, we have studied the dynamics of insulin-induced Raf-1 translocation to the plasma membrane in live cells. Our data demonstrate that Raf-GFP undergoes growth factor-induced kinase activation and translocation, indicating that Raf-GFP is an appropriate model for growth factor-induced Raf-1 dynamics. In order to assess translocation of Raf-GFP to the plasma membrane in live cells, we imaged a confocal section of the plasma membrane adjacent to the cover glass. To select this section, several images along the z axis of the cell were collected. Because Raf-1 does not enter the nucleus, the plasma membrane sections adjacent to the coverslip were identified by choosing a confocal section below the nucleus. These sections were imaged at 37 °C in order to examine the kinetics of Raf translocation in response to insulin or PA stimulation.

By using this experimental approach, we show here that the association of Raf-1 to the plasma membrane is transient. However, Raf-1 does not simply dissociate from the plasma membrane. Dual staining immunogold labeling for the insulin receptor and Raf-1 resolved by electron microscopy shows that Raf-1 co-localized with the insulin receptor in intracellular vesicular structures in response to stimulation with both insulin and PA. The structure of these vesicles suggests that they are endosomes. To confirm that Raf-1 binds endocytic vesicles, cells were treated with insulin, and vesicles containing the insulin receptor were isolated using a specific anti-insulin receptor antibody. Both Raf-1 and the heavy chain of clathrin were present in these preparations. Consistent with our model, the association between Raf-1 and the isolated vesicles also appeared to be dependent on the presence of PA. This conclusion is based on the observation that BFA blocked the internalization of the insulin receptor and abolished the localization of Raf-1 in the immunoisolated endocytic vesicles. Therefore, we propose that PA is required for endocytosis of the insulin receptor and that the association between Raf-1 and endosomes is mediated by a direct interaction between PA and Raf-1. This is consistent with current models of vesicle formation mediated by acidic phospholipids (32, 33, 52-55) which suggest that PA may form an integral part of newly formed vesicles. These models suggest that PLD-mediated generation of PA, through participation in a positive feedback loop concurrently with the generation of PIP2, may sufficiently perturb membrane structure and facilitate the formation of a vesicle from a planar membrane. Consequently, the membranes of newly formed vesicles are enriched with the acidic phospholipids PA and PIP2. This acidic surface may serve as a binding matrix for a number of signaling molecules such as Raf-1.

Recent work by Daaka et al. (47) also supports this model. By using dominant suppressor mutants of beta -arrestin or dynamin, they showed that inhibition of G-protein-coupled receptor endocytosis blocked MAPK phosphorylation. Furthermore, they isolated Raf-1-containing vesicles that co-isolated with clathrin-coated vesicles, suggesting that Raf-1 associates with clathrin-coated vesicles. Our data suggest a very similar model for insulin-mediated activation of MAPK. We suggest that the activation of the MAPK cascade by insulin also requires endocytosis of the insulin receptor. Raf-1 is associated with endocytic vesicles in insulin-treated cells, and PLD activation appears to be necessary for receptor-mediated endocytosis and Raf-1 translocation to membranes.

Fig. 8 depicts the proposed role of insulin-induced generation of PA. According to this model, PA has the following two functions: 1) the recruitment of Raf-1 to the plasma membrane where it is activated by factors which reside on the plasma membrane, such as activated Ras and PKC, and 2) facilitation of endocytic vesicle formation. These two effects, acting in conjunction, may result in the recruitment of many important components of signal transduction to the plasma membrane and to the membranes of endocytic vesicles. Among these components are receptor tyrosine kinases, Raf-1 through its association with PA, and proteins that associate with PIP2 through pleckstrin homology domains. Our work and that of Daaka et al. (47) further suggest that the internalization of these signaling components is necessary for full activation of the MAPK signaling cascade, probably by bringing Raf-1 in contact with downstream targets such as MEK. In this paper, we demonstrate that this phenomenon is mediated by phosphatidic acid.


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Fig. 8.   A model for the contribution of PLD and its product, PA, in mitogenic signaling. Activation of insulin receptors results in the stimulation of PLD mediated by ARF and in the activation of Ras via the phosphorylation of insulin receptor substrate-1 (IRS-1). PLD acts to generate PA which then 1) aids in the recruitment of Raf-1 to the plasma membrane where it interacts with GTP-Ras; and 2) facilitates formation of the endocytic vesicle. Raf-1 then remains bound to the endocytic vesicle via its association with PA. The internalized Raf-1 then propagates the MAPK signaling cascade.


    ACKNOWLEDGEMENTS

We thank Dr. Adrienne D. Cox for the Ha-Ras(Q61L)-transformed cells and Dr. Saïd Sebti for the plasmid encoding c-Raf-1. We also thank Dr. Edwin Levitan for a critical review of this manuscript.

    FOOTNOTES

* This work was supported by National Institutes of Health Grants DK51183, DK02465, 5-T32-GM08424-04, and GM54813 and American Diabetes Association Grant 96-029.The costs of publication of this article were defrayed in part by the payment of page charges. The 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: Dept. of Pharmacology, University of Pittsburgh, Pittsburgh, PA 15261. Tel.: 412-648-9408; Fax: 412-648-1945; E-mail: ggr+{at}pitt.edu.

The abbreviations used are: ARF, ADP-ribosylation factor; BFA, brefeldin A; CRD, cysteine-rich domain; DAG, diacylglycerol; LPA, lysophosphatidic acid; MAPK, mitogen-activated protein kinase; PA, phosphatidic acid; PIP2, phosphatidylinositol 4,5-bisphosphate; PKC, protein kinase C; PLD, phospholipase D; Raf-GFP, Raf-green fluorescent protein fusion protein; PAGE, polyacrylamide gel electrophoresis; PBS, phosphate-buffered saline; GTPgamma S, guanosine 5'-3-O-(thio)triphosphate.

2 T.-C. Sung, Y. Zhang, A. J. Morris, and M. A. Frohman, submitted for publication.

3 T. S. Sung, A. J. Morris, and M. A. Frohman, submitted for publication.

    REFERENCES
Top
Abstract
Introduction
Procedures
Results
Discussion
References

  1. Ben-Av, P., Eli, Y., Schmidt, U. S., Tobias, K. E., and Liscovitch, M. (1993) Eur. J. Biochem. 215, 455-463[Abstract]
  2. Donchenko, V., Zannetti, A., and Baldini, P. M. (1994) Biochim. Biophys. Acta 1222, 492-500[Medline] [Order article via Infotrieve]
  3. Price, B. D., Morris, J. D., and Hall, A. (1989) Biochem. J. 264, 509-515[Medline] [Order article via Infotrieve]
  4. Yeo, E.-J., and Exton, J. H. (1995) J. Biol. Chem. 270, 3980-3988[Abstract/Free Full Text]
  5. Brown, H. A., Gutowski, S., Moomaw, C. R., Slaughter, C., and Sternweis, P. C. (1993) Cell 75, 1137-1144[Medline] [Order article via Infotrieve]
  6. Hammond, S. M., Altshuller, Y. M., Sung, T.-C., Rudge, S. A., Rose, K., Engebrecht, J., Morris, A. J., and Frohman, M. A. (1995) J. Biol. Chem. 270, 29640-29643[Abstract/Free Full Text]
  7. Malcolm, K. C., Ross, A. H., Qui, R.-G., Symons, M., and Exton, J. H. (1994) J. Biol. Chem. 269, 25951-25954[Abstract/Free Full Text]
  8. Frohman, M. A., and Morris, A. J. (1996) Curr. Biol. 6, 945-947[Medline] [Order article via Infotrieve]
  9. Singer, W. D., Brown, H. A., and Sternweis, P. C. (1997) Annu. Rev. Biochem. 66, 475-509[CrossRef][Medline] [Order article via Infotrieve]
  10. Shome, K., Vasudevan, C., and Romero, G. (1997) Curr. Biol. 7, 387-396[Medline] [Order article via Infotrieve]
  11. Karnam, P., Standaert, M. L., Galloway, L., and Farese, R. V. (1997) J. Biol. Chem. 272, 6136-6140[Abstract/Free Full Text]
  12. Bi, K., Roth, M. G., and Ktistakis, N. T. (1997) Curr. Biol. 7, 301-307[Medline] [Order article via Infotrieve]
  13. English, D. (1996) Cell. Signalling 8, 341-347[CrossRef][Medline] [Order article via Infotrieve]
  14. Exton, J. H. (1997) Physiol. Rev. 77, 303-320[Abstract/Free Full Text]
  15. Ha, K. S., and Exton, J. H. (1993) J. Cell Biol. 123, 1789-1796[Abstract]
  16. Reinhold, S. L., Prescott, S. M., Zimmerman, G. A., and McIntyre, T. M. (1990) FASEB J. 4, 208-214[Abstract/Free Full Text]
  17. Zhao, Z., Shen, S. H., and Fischer, E. H. (1993) Proc. Natl. Acad. Sci. U. S. A. 90, 4251-4255[Abstract]
  18. Jones, G. A., and Carpenter, G. (1993) J. Biol. Chem. 268, 20845-20850[Abstract/Free Full Text]
  19. Tsai, M. H., Yu, C. L., Wei, F. S., and Stacy, D. W. (1989) Science 243, 522-526[Medline] [Order article via Infotrieve]
  20. Ghosh, S., Strum, J. C., Sciorra, V. A., Daniel, L., and Bell, R. M. (1996) J. Biol. Chem. 271, 8472-8480[Abstract/Free Full Text]
  21. Sung, T. C., Roper, R. L., Zhang, Y., Rudge, S. A., Temel, R., Hammond, S. M., Morris, A. J., Moss, B., Engebrecht, J., and Frohman, M. A. (1997) EMBO J. 16, 4519-4530[Abstract/Free Full Text]
  22. Lerner, E. C., Qian, Y., Blaskovich, M. A., Fossum, R. D., Vogt, A., Sun, J., Cox, A. D., Der, C. J., Hamilton, A. D., and Sebti, S. M. (1995) J. Biol. Chem. 270, 26802-26806[Abstract/Free Full Text]
  23. App, H., Hazan, R., Zilberstein, A., Ullrich, A., Schlessinger, J., and Rapp, U. (1991) Mol. Cell. Biol. 11, 913-919[Medline] [Order article via Infotrieve]
  24. Turner, B., Rapp, U., App, H., Greene, M., Dobashi, K., and Reed, J. (1991) Proc. Natl. Acad. Sci. U. S. A. 88, 1227-1231[Abstract]
  25. Der, C. J., Finkel, T., and Cooper, G. M. (1986) Cell 44, 167-176[Medline] [Order article via Infotrieve]
  26. Shome, K., Xu, X., and Romero, G. (1995) FEBS Lett. 357, 109-114[CrossRef][Medline] [Order article via Infotrieve]
  27. Clark, S., Eckardt, G., Siddle, K., and Harrison, L. C. (1991) Biochem. J. 276, 27-33[Medline] [Order article via Infotrieve]
  28. Martin, T. F. (1989) Methods Enzymol. 168, 225-33[Medline] [Order article via Infotrieve], 1989
  29. Donaldson, J. G., Finazzi, D., and Klausner, R. D. (1992) Nature 360, 350-352[CrossRef][Medline] [Order article via Infotrieve]
  30. Helms, J. B., and Rothman, J. E. (1992) Nature 360, 352-354[CrossRef][Medline] [Order article via Infotrieve]
  31. Randazzo, P. A., Yang, Y. C., Rulka, C., and Kahn, R. A. (1993) J. Biol. Chem. 268, 9555-9563[Abstract/Free Full Text]
  32. Ktistakis, N. T., Brown, H. A., Waters, M. G., Sternweis, P. C., and Roth, M. G. (1996) J. Cell Biol. 134, 295-306[Abstract]
  33. Chung, J.-K., Sekiya, F., Kang, H.-S., Lee, C., Han, J.-S., Kim, S. R., Bae, Y. S., Morris, A. J., and Rhee, S. G. (1997) J. Biol. Chem. 272, 15980-15985[Abstract/Free Full Text]
  34. Avruch, J., Zhang, X., and Kyriakis, J. M. (1994) Trends Biochem. Sci. 19, 279-283[CrossRef][Medline] [Order article via Infotrieve]
  35. Daum, G., Eisenmann-Tappe, I., Fries, H., Troppmair, J., and Rapp, U. R. (1994) Trends Biochem. Sci. 19, 474-479[CrossRef][Medline] [Order article via Infotrieve]
  36. Morrison, D. K., and Cutler, R. E., Jr. (1997) Curr. Biol. 9, 174-179[CrossRef]
  37. Leevers, S. J., Paterson, H. F., and Marshall, C. J. (1994) Nature 369, 411-414[CrossRef][Medline] [Order article via Infotrieve]
  38. Mineo, C., Anderson, R. G. W., and White, M. A. (1997) J. Biol. Chem. 272, 10345-10348[Abstract/Free Full Text]
  39. Roy, S., Lane, A., Yan, J., McPherson, R., and Hancock, J. F. (1997) J. Biol. Chem. 272, 20139-20145[Abstract/Free Full Text]
  40. Luo, Z., Diaz, B., Marshall, M. S., and Avruch, J. (1997) Mol. Cell. Biol. 17, 46-53[Abstract]
  41. Huwiler, A., Brunner, J., Hummel, R., Vervoordeldonk, M., Stabel, S., van der Bosch, H., and Pfeilschifter, J. (1996) Proc. Natl. Acad. Sci. U. S. A. 93, 6959-6963[Abstract/Free Full Text]
  42. Hu, C.-D., Kariya, K., Kotani, G., Shirouzu, M., Yokoyama, S., and Kataoka, T. (1997) J. Biol. Chem. 272, 11702-11705[Abstract/Free Full Text]
  43. Hammond, S. M., Altshuller, Y. M., Sung, T. C., Rudge, S. A., Rose, K., Engebrecht, J., Morris, A. J., and Frohman, M. A. (1995) J. Biol. Chem. 270, 29640-29643[Abstract/Free Full Text]
  44. Kodaki, T., and Yamashita, S. (1997) J. Biol. Chem. 272, 11408-11413[Abstract/Free Full Text]
  45. Hammond, S. M., Jenco, J. M., Nakashima, S., Cadwallader, K., Gu, Q., Cook, S., Nozawa, Y., Pystivich, G. D., Frohman, M. A., and Morris, A. J. (1997) J. Biol. Chem. 272, 3860-3868[Abstract/Free Full Text]
  46. Colley, W., Sung, T. C., Roll, R., Hammond, S. M., Altshuller, Y. M., Bar-Sagi, D., Morris, A. J., and Frohman, M. A. (1997) Curr. Biol. 7, 191-201[Medline] [Order article via Infotrieve]
  47. Daaka, Y., Luttrell, L. M., Ahn, S., Della Rocca, G. J., Ferguson, S. S. G., Caron, M. G., and Lefkowitz, R. J. (1998) J. Biol. Chem. 273, 685-688[Abstract/Free Full Text]
  48. van Dijk, M. C. M., Muriana, F. J. G., van der Hoeven, P. C. J., de Widt, J., Schaap, D., Moolenaar, W. H., and van Blitterswijk, W. J. (1997) Biochem. J. 323, 693-699[Medline] [Order article via Infotrieve]
  49. van Dijk, M. C. M., Postma, F., Hilkmann, H., Jalink, K., van Blitterswijk, W. J., and Moolenaar, W. H. (1998) Curr. Biol. 8, 386-392[Medline] [Order article via Infotrieve]
  50. Traverse, S., Cohen, P., Paterson, H., Marshall, C., Rapp, U., and Grand, R. J. A. (1993) Oncogene 8, 3175-3181[Medline] [Order article via Infotrieve]
  51. Marais, R., Light, Y., Paterson, H. F., and Marshall, C. J. (1995) EMBO J. 14, 3136-3145[Abstract]
  52. Liscovitch, M., and Cantley, L. C. (1995) Cell 81, 659-662[Medline] [Order article via Infotrieve]
  53. Rothman, J. E. (1994) Nature 372, 55-63[CrossRef][Medline] [Order article via Infotrieve]
  54. Takei, K., Haucke, V., Slepnev, V., Farsad, K., Salazar, M., Chen, H., and De Camilli, P. (1998) Cell 94, 131-141[Medline] [Order article via Infotrieve]
  55. Matsuoka, K., Orci, L., Amherdt, M., Bednarek, S., Hamamoto, S., Schekman, R., and Yeung, T. (1998) Cell 93, 263-275[Medline] [Order article via Infotrieve]


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