Correspondence to Anne Kenworthy: anne.kenworthy{at}vanderbilt.edu
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Introduction |
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The hypervariable domain additionally functions to regulate the subcellular distribution, intracellular trafficking, and membrane microenvironment of Ras. The CAAX motif targets Ras to the cytosolic face of the ER and Golgi apparatus, and exit of Ras from these compartments requires either palmitoylation or the polybasic domain (Choy et al., 1999; Apolloni et al., 2000). How trafficking of Ras to the cell surface is accomplished depends on the nature of the second signal. The palmitoylated Ras isoforms HRas and NRas are delivered from the Golgi complex to the cell surface as part of the secretory pathway, whereas KRas 4B reaches the plasma membrane by an unknown mechanism that is independent of vesicular transport. The COOH-terminal membrane targeting motifs of Ras appear to contain the relevant signals for Ras trafficking, as these motifs traffic GFP to the plasma membrane in a similar manner as the full-length protein (Choy et al., 1999; Apolloni et al., 2000). In adipocytes and yeast, HRas can traffic to the plasma membrane by a nonclassical secretory transport pathway as well as the classic secretory pathway (Dong et al., 2003; Watson et al., 2003). Palmitoylated forms of Ras are also often found associated with the Golgi complex where they can signal, providing a mechanism for regulation of isoform-specific Ras signaling via their distinct subcellular localizations (for review see Bivona and Philips, 2003; Hancock, 2003). The hypervariable domain further contributes to specificity of Ras signaling through different isoforms by targeting the proteins to spatially and compositionally distinct plasma membrane microdomains (for review see Hancock, 2003; Parton and Hancock, 2004).
How palmitoylation contributes to the isoform-specific trafficking and signaling of Ras has not been fully established. One proposed function of palmitoylation is to enhance Ras binding to membranes (Silvius and l'Heureux, 1994; Shahinian and Silvius, 1995; Silvius, 2002). Palmitoylation may also regulate the sorting of Ras into vesicles destined for the cell surface or targeted for clathrin-independent endocytosis (Smotrys and Linder, 2004). Both mechanisms could be potentially modulated in a dynamic manner, as the palmitates on NRas and HRas undergo dynamic turnover within minutes to hours (Magee et al., 1987; Lu and Hofmann, 1995; Baker et al., 2000, 2003). How this turnover is regulated and its significance for Ras biology is not yet known, as the enzymes involved in the regulation of Ras palmitoylation and depalmitoylation have only recently begun to be identified (Linder and Deschenes, 2003, 2004; Dietrich and Ungermann, 2004; Smotrys and Linder, 2004).
In this study, we examined the role of palmitoylation in the intracellular transport of HRas and NRas to and from the Golgi complex. Using quantitative fluorescence microscopy and photobleaching techniques, we show that GFP-tagged mutants of HRas and NRas lacking functional palmitoylation sites undergo rapidly reversible binding to the ER and Golgi complex. We also provide evidence that wild-type NRas and HRas undergo a cycle of depalmitoylation and repalmitoylation that allows them to recycle to the Golgi complex. We propose that the reversible palmitoylation of Ras allows the protein to shift between vesicular and nonvesicular modes of transport, and ultimately controls the location and time course of intracellular Ras signaling.
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Results |
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To test this hypothesis, we first confirmed that GFP-NRas (Fig. 4 A) and GFP-HRas (unpublished data) remain Golgi associated after cycloheximide treatment. We next asked whether Golgi-associated Ras was actively trafficked to the Golgi complex in the absence of new protein synthesis by photobleaching the entire Golgi-associated pool of protein, then monitoring recovery of fluorescence in the area over time (Fig. 4 B). Both GFP-NRas and GFP-HRas recovered after photobleaching, with similar half times (48 ± 9 s vs. 36 ± 5 s, respectively) but to differing extents (64 ± 4% vs. 39 ± 3%, n = 33 and 23 cells, respectively). Thus, a fraction of both wild-type GFP-HRas and GFP-NRas appear to be actively and rapidly recycled to the Golgi complex; the remainder of the protein, which does not recover after the bleach, appears to represent a stably bound pool.
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Discussion |
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Because nonpalmitoylated Ras has access to the cytosol, our findings raise the question of why it does not significantly bind the plasma membrane (Choy et al., 1999; Apolloni et al., 2000). One possibility is that the lipid composition of the ER and Golgi membranes is preferable for insertion of prenyl moieties. However, in vitro binding of prenylated peptides to liposomes show little dependence on membrane composition (Silvius and l'Heureux, 1994). Alternatively, membrane potential and/or electrostatic interactions, which presumably are responsible for the specific binding of KRas to the cell surface (Choy et al., 1999; Apolloni et al., 2000), may play an inhibitory role in preventing plasma membrane binding in the absence of a second plasma membrane binding signal. Finally, as discussed in the previous paragraph, putative prenyl binding proteins localized to the ER and Golgi complex could potentially act as sites for transient binding interactions.
The palmitoylation state of Ras determines whether the protein undergoes vesicular or nonvesicular transport
Our data suggest that Ras can shift between vesicular and nonvesicular transport by regulating its palmitoylation state in a manner consistent with the "kinetic trap" model of palmitoylation (Shahinian and Silvius, 1995). This model proposes that farnesylated (but nonpalmitoylated) peptides can efficiently but reversibly bind membranes until they are palmitoylated, trapping them on the membrane. Such kinetic trapping can readily explain why Ras palmitoylation mutants are rapidly and reversibly bound to membranes, whereas a substantial fraction of palmitoylated HRas and NRas is stably bound to the Golgi complex.
As an extension of this model, we propose that the intracellular sites at which Ras palmitoyl acyl transferase (PAT) enzymes reside define the entry points for Ras into vesicular transport pathways. For example, if PAT activity is present in the ER or intermediate compartment, newly synthesized Ras would enter into vesicles early in the secretary pathway. Conversely, if the PAT activity is localized exclusively at the plasma membrane, Ras could potentially traffic to the plasma membrane completely independently of vesicular transport. The latter possibility may explain why in adipocytes, delivery of HRas to the cell surface occurs in the presence of BFA or after a 20° block (Watson et al., 2003). This nonclassical cell surface transport pathway is an excellent candidate for regulation via a cycle of palmitoylation and depalmitoylation (Watson et al., 2003). A similar model could explain how peptides mimicking the NRas COOH terminus reach to the plasma membrane independent of the secretory pathway (Schroeder et al., 1997).
Our understanding of the role of palmitoylation in Ras trafficking will be greatly enhanced by the identification of the proteins responsible for this process. Although palmitoylation can occur by nonenzymatic means, there is evidence that in cells this is a regulated event (Dietrich and Ungermann, 2004; Smotrys and Linder, 2004). Although the enzymes responsible for palmitoyl transferase activity have long been sought (Kasinathan et al., 1990; Gutierrez and Magee, 1991; Berthiaume and Resh, 1995; Das et al., 1997), only recently have candidate Ras PATs been identified. Studies in yeast first identified an ER-localized protein complex, Erf2/Erf4, that stimulates palmitoylation of Ras2 in vitro (Lobo et al., 2002). These proteins contain a DHHC cysteine-rich domain that has been postulated to be a signature of proteins involved in palmitoylation (Smotrys and Linder, 2004). Very recently, several candidate mammalian Ras palmitoyltransferases have been identified (Ducker et al., 2004; Fukata et al., 2004; Huang et al., 2004). Given these recent breakthroughs, it should be possible to begin to directly dissect the role of these proteins in Ras localization and function in the near future.
It is important to note that the kinetic trapping model does not exclude other potential roles for palmitoylation in the regulation of Ras trafficking. For example, the yeast Ras homologue Ras2p is trafficked to the plasma membrane in the absence of a functional secretory pathway in a process requiring Erf2p, a putative ER-localized palmitoyltransferase, by an as-yet-unidentified mechanism (Dong et al., 2003). It is possible that palmitoylation-dependent interactions allow Ras to interact with chaperone proteins that traffic the protein by vesicle-independent pathways. Alternatively, palmitoylation-dependent targeting of Ras to raft-enriched or other types of membrane microdomains may be important for allowing it either to exit the Golgi complex (Magee and Marshall, 1999) or to be internalized from the cell surface by either clathrin-dependent or -independent endocytic pathways (Roy et al., 2002).
Regulation of the rate of depalmitoylation of NRas versus HRas
The potential for Ras to undergo multiple rounds of depalmitoylation and subsequent repalmitoylation were first suggested by studies of NRas, which indicated that the half-life of palmitate is shorter than the life span of the protein (Magee et al., 1987). One enzyme that has been demonstrated to remove palmitate from HRas in vitro is acyl-thioesterase 1 (Smotrys and Linder, 2004). However, much remains to be learned about how this event is regulated. Although it is tempting to speculate that the primary pool of Ras undergoing depalmitoylation is localized to the cell surface, it is unclear whether specific subcellular pools of Ras are preferred substrates for depalmitoylation.
Interestingly, we found that the rate of palmitate turnover differs for HRas and NRas. First, a larger fraction of wild-type NRas than HRas was reversibly bound to the Golgi complex (Fig. 4). This implies that a larger fraction of NRas than HRas is present in a depalmitoylated state under steady-state conditions. In addition, after 2BP treatment NRas redistributed to the ER and Golgi more rapidly than HRas (Fig. 6), suggesting that NRas undergoes a more rapid rate of depalmitoylation. These observations are consistent with reports that the half-life of palmitate on HRas, ranging from 90 min to 2.4 h (Lu and Hofmann, 1995; Baker et al., 2003), is relatively long compared with that of NRas (20 min) (Magee et al., 1987). Because HRas contains two palmitoylation sites compared with NRas's one, it is likely that the difference in overall half-lives reflects a higher probability that at least one palmitate will be present on HRas. Indeed, the half-life of palmitate on overexpressed HRas was previously shown to be reduced from 90 min to 15 min upon mutation of one of the palmitoylation sites, with slower turnover correlating with stronger membrane binding (HRas Ser181) (Lu and Hofmann, 1995). This same study showed little evidence for specific recognition of palmitoylated proteins, thus suggesting that access to a depalmitoylating enzyme determined the palmitate turnover rate. Our finding that depalmitoylation plays a role in determining the subcellular distribution and trafficking of HRas and NRas highlights the need for further characterization of the regulation of these events.
Nature of the soluble pool of Ras
The presence of a soluble pool of Ras has been noted by several groups, with the fraction of soluble protein ranging from 1020% to upwards of 4050% (Magee et al., 1987; Hancock et al., 1990; Lu and Hofmann, 1995; Choy et al., 1999; Webb et al., 2000; Baker et al., 2003). Our data suggest that this soluble pool corresponds to transiently depalmitoylated Ras, in agreement with a previous study showing that soluble NRas is farnesylated but not palmitoylated (Magee et al., 1987). How Ras is solubilized in the presence of a farnesyl moiety is not yet known. Delivery of geranylgeranylated Rab proteins to membranes is mediated by Rab escort protein (REP) or a Rab GDP dissociation inhibitor (GDI) (for review see Seabra and Wasmeier, 2004). The prenylated Rab acceptor protein (PRA1) and phosphodiesterase- are two candidate Ras escort proteins (Figueroa et al., 2001; Hanzal-Bayer et al., 2002; Nancy et al., 2002). However, it should be noted that in vitro peptide binding experiments show that farnesylated proteins have an intrinsically weak affinity for membranes, and thus may not require a specialized mechanism to allow them to become solubilized (Silvius and l'Heureux, 1994). Yet another possibility, suggested by biochemical studies in progress, is that soluble farnesylated Ras exists as a dimer (unpublished data). Our FCS studies show that soluble Ras has a diffusional mobility similar to cytoplasmic GFP (Fig. 3, Table I). Given the weak dependence of diffusion on protein size, these data are consistent with the possibility that soluble Ras exists either as a monomer or in small complexes with itself or other proteins. More work will be required to distinguish between these possibilities.
Implications for Ras signaling
We envision several mechanisms by which the regulation of the palmitoylation state of Ras could control the intracellular location and time course of Ras signaling. First, the loss of palmitate on Ras may allow for the regulated release of the protein from the cell surface. Such an event could even be regulated by Ras activation itself, as the depalmitoylation of GTP-bound HRas is accelerated compared with the GDP-bound form (palmitate half-life of 10 min vs. 2.4 h, respectively) (Baker et al., 2003). Given that depalmitoylated Ras can still efficiently bind intracellular membranes, such an event may not have a large effect on the subcellular distribution of Ras as assessed by biochemical criteria. This could explain why 2BP treatment causes only a 720% increase in the soluble pool of HRas (Webb et al., 2000). After depalmitoylation, activated Ras could itself act as a diffusible signaling intermediate, allowing the protein to rapidly redistribute to other intracellular compartments. Evidence supporting this model was very recently reported (Peyker et al., 2005; Rocks et al., 2005). The reversible binding of depalmitoylated Ras may also explain the rapid (2 min after stimulation) recruitment of GFP-RBD to ER membranes in cells overexpressing HRas palmitoylation mutants (Chiu et al., 2002). This also implies that reversible membrane binding of Ras is sufficient to enable efficient signaling. Finally, a cycle of depalmitoylation and repalmitoylation may regulate intracellular Ras signaling by maintaining a steady-state pool of Ras on the Golgi complex (Chiu et al., 2002; Bivona et al., 2003; Caloca et al., 2003; Mitin et al., 2004; Perez de Castro et al., 2004; Rocks et al., 2005). This may also contribute to the specific outcomes of HRas signaling in response to altered membrane-targeting signals (Booden et al., 1999, 2000; Coats et al., 1999). Disruption of Ras recycling to the Golgi complex may thus offer a potential mechanism to interfere with oncogenic Ras activity.
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Materials and methods |
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Nocodazole and cycloheximide treatments
To disrupt microtubules, cells were preincubated for 5 min on ice in DME containing 10% fetal calf serum and 50 mM Hepes. The cells were then treated with 5 µg/ml nocodazole (Sigma-Aldrich) for 15 min on ice, warmed for 5 min to 37°C, and imaged in the continued presence of nocodazole at 37°C. Control experiments were performed using vehicle alone (DMSO). To inhibit new protein synthesis, cells were treated with 200 µg/ml cycloheximide (Sigma-Aldrich) in DME, 10% fetal calf serum, and 50 mM Hepes for 4 h at 37°C. The cells were then imaged at 37°C in the cycloheximide solution.
Fluorescence microscopy and photobleaching measurements
Cells were imaged with an inverted laser scanning confocal microscope (model 510; Carl Zeiss MicroImaging, Inc.) equipped with the Confocor2 for FCS (Carl Zeiss MicroImaging, Inc.). Where indicated, an Air Stream Stage Incubator (Nevtek) was used for imaging at 37°C. GFP was excited with an argon laser with excitation at 488 nm and emission was detected with a GFP long-pass (LP) 505 or 530 or band-pass (BP) 505530 filter. A Plan-Neofluar 40x/1.3 oil immersion lens was used for imaging all samples. Cells were maintained in phenol-red free DME containing 10% fetal calf serum and 50 mM Hepes for live-cell imaging experiments.
Confocal FRAP measurements were performed using a previously described protocol (Kenworthy et al., 2004). In brief, a strip 4 µm wide was photobleached using high laser intensity and fluorescence recovery monitored at low intensity. Diffusion coefficients were calculated from whole-cell recoveries using a program that simulates diffusion (Siggia et al., 2000). Mf was calculated as described previously (Ellenberg et al., 1997). Statistical differences were evaluated using the t test.
In experiments measuring kinetics of Golgi refilling, an area containing the entire Golgi was bleached (Nichols et al., 2001). Halftimes of recovery were calculated as described in Feder et al. (1996), and the final percentage of recovered fluorescence was calculated as for Mf after correcting for the loss of fluorescence due to the photobleaching event. Control experiments on fixed cells confirmed that the loss of fluorescence was confined to the bleached region.
All quantitative image analysis was performed using unprocessed images. For presentation purposes, image contrast was adjusted using Adobe Photoshop.
Quantitation of Ras localization after 2BP treatment
Cells were treated with 25 µM 2BP (Sigma-Aldrich) or vehicle (DMSO) at 37°C, either immediately after transfection or 18 h after transfection for the indicated times (30 min, 2 h, or 5 h). After treatment, cells were imaged live with the confocal pinhole fully open for quantitation or set at 12 Airy units for presentation purposes. The subcellular distribution of Ras in 20 cells/treatment was quantitated in two ways. (1) Localization of Ras in the ER/nuclear envelope versus plasma membrane. Cells were scored for the relative amounts of ER/nuclear envelope labeling ranging from ER labeling but no apparent plasma membrane stain (++++) to no apparent ER or nuclear envelope label (). These scores were converted to a numeric value as follows: ++++ (1 pt), +++/ (0.75 pt), ++/ (0.5 pt), +/ (0.25 pt), or (0 pt). The total numeric score for all cells at a given time point was calculated for each experiment. (2) Fraction of Ras localized to the Golgi complex. Images were converted to tiff format, and the average fluorescence in the Golgi region versus in the entire cell was calculated using NIH Image. After background subtraction, the ratio of fluorescent material in the Golgi region versus the entire cell was calculated.
Fluorescence correlation spectroscopy
FCS measures time-dependent fluorescence fluctuations in a diffraction-limited (0.1 femtoliter) volume defined using confocal microscope optics with a sensitive avalanche photodiode detector. Intensity fluctuations corresponding to the movements of individual molecules in and out of the confocal volume are recorded over time. Fluorescence fluctuations reflect the average residence time of the fluorescent species in the confocal volume, which in turn are a function of its characteristic diffusional mobility. The diffusion of fluorescently tagged proteins through the sampling volume occurs with a characteristic D. This is related to the diffusion coefficient D by
D = (
02)/(4D), where
0 is the radius of the laser beam. Thus, a longer
D corresponds to a slower D. The intensity fluctuations are characterized by their average value <I> and their fluctuations
I(t) = I(t) <I> and can be analyzed using an autocorrelation function G(
). The normalized autocorrelation function is given by G(
) = 1 + <
I(t) *
I(t +
)> * <I>2, where I(t) is the time-dependent fluorescence intensity,
is a short time interval after any arbitrary time t, and <I> is the mean value of fluorescence intensity.
FCS experiments were performed on a microscope (LSM 510; Carl Zeiss MicroImaging, Inc.) outfitted with ConfoCor2 (Carl Zeiss MicroImaging, Inc.), combining both FCS and confocal laser scanning capabilities. A C-Apochromat 40x 1.2 NA water objective was used in conjunction with a dichroic filter and 520-nm long-pass filter to focus and separate exciting and emitting radiation. GFP-tagged constructs were excited at 488 nm with a 40-mW argon laser. Aqueous rhodamine 6G solutions were used to calculate the confocal volume radius 0 = 1.44 x 107 m, resulting in a confocal volume element of 0.1 fl.
Cells were imaged in LSM mode and, after selection of an appropriate cell, whole cells were repeatedly bleached to reduce the fluorescence to allow for FCS measurements. A line scan in the axial direction was performed to set the volume element at an appropriate position in the cell. FCS measurements were made for 10 s each and repeated 10 times per cell. The autocorrelation curves were fit using software provided by the manufacturer. The autocorrelation function G() for a two-component model is described by the following equation: G(
) = 1 + (1/N) [(1 Y) (1 +
/
D1)1 (1 +
/S2
D1)1/2 + Y (1 +
/
D2)1 (1 +
/S2
D2)1/2].
Here, N is the number of fluorescent particles in the confocal volume; S is the structure parameter (defined by the dimensions of the confocal volume); D1 and
D2 are the average residence times of the first and second component, respectively; and Y and 1 Y are the fraction of particles in the confocal volume with diffusion time
D2 and
D1, respectively. Data were fit assuming a constant structure parameter of 5.0. Diffusion coefficients were calculated from the fitted values of
D and the known confocal radius
0 as described above. Data were obtained from 1020 cells from two independent experiments.
Online supplemental material
The online supplemental material describes control experiments performed to test whether overexpressed GFP-Ras fusion proteins are quantitatively farnesylated, and is available at http://www.jcb.org/cgi/content/full/jcb.200502063/DC1.
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Acknowledgments |
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A. Kenworthy was supported by a fellowship from the National Research Council and a New Faculty Development Award from the Department of Molecular Physiology and Biophysics, Vanderbilt University School of Medicine.
Submitted: 10 February 2005
Accepted: 15 June 2005
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