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Mol Biol Cell. Mar 1, 2009; 20(5): 1565–1575.
PMCID: PMC2649262

TORC2 Plasma Membrane Localization Is Essential for Cell Viability and Restricted to a Distinct Domain

Carole Parent, Monitoring Editor

Abstract

The conserved target of rapamycin (TOR) kinases regulate many aspects of cellular physiology. They exist in two distinct complexes, termed TOR complex 1 (TORC1) and TOR complex 2 (TORC2), that posses both overlapping and distinct components. TORC1 and TORC2 respond differently to the drug rapamycin and have different cellular functions: whereas the rapamycin-sensitive TORC1 controls many aspects of cell growth and has been characterized in great detail, the TOR complex 2 is less understood and regulates actin polymerization, cell polarity, and ceramide metabolism. How signaling specificity and discrimination between different input signals for the two kinase complexes is achieved is not understood. Here, we show that TORC1 and TORC2 have different localizations in Saccharomyces cerevisiae. TORC1 is localized exclusively to the vacuolar membrane, whereas TORC2 is localized dynamically in a previously unrecognized plasma membrane domain, which we term membrane compartment containing TORC2 (MCT). We find that plasma membrane localization of TORC2 is essential for viability and mediated by lipid binding of the C-terminal domain of the Avo1 subunit. From these data, we suggest that the TOR complexes are spatially separated to determine downstream signaling specificity and their responsiveness to different inputs.

INTRODUCTION

During cell growth and division, many physiological processes need to be coordinated and modified according to nutrient availability. Signaling through target of rapamycin (TOR) kinases plays a central role in this regulation. Even though TOR kinases are related to phosphoinositide lipid kinases, they are Ser/Thr protein kinases, with a small number of known targets that regulate many cellular processes. Together, the diverse TOR signaling outputs regulate cell growth, both spatially and temporally (De Virgilio and Loewith, 2006 blue right-pointing triangle; Reiling and Sabatini, 2006 blue right-pointing triangle; Wullschleger et al., 2006 blue right-pointing triangle).

The characterization of TOR signaling was greatly aided by the discovery of the antifungal and immunosuppressant macrocyclic lactone rapamycin. The search for targets of this drug in the yeast Saccharomyces cerevisiae lead to the discovery of the TOR kinases and subsequently helped to identify their molecular function (Heitman et al., 1991 blue right-pointing triangle).

The rapamycin-sensitive branch of TOR signaling regulates processes that collectively modulate the rate of cell growth. TOR signaling is active under conditions of excess nutrients, blocking catabolic processes, such as autophagy and stress responses. In contrast, exposure to rapamycin or withdrawal of nitrogen or carbon sources lead to inactivation of TOR signaling and down-regulation of anabolic processes, such as protein synthesis. Together these processes coordinate cell growth with nutrient availability (De Virgilio and Loewith, 2006 blue right-pointing triangle; Wullschleger et al., 2006 blue right-pointing triangle). This function of TOR signaling is performed by TOR complex 1 (TORC1) that consists of the kinase plus three additional subunits in yeast, named Lst8, Kog1, and Tco89 or two subunits named raptor and mLst8 in mammals. Its molecular targets include translation initiation factors (e.g., eukaryotic initiation factor [eIF]4E, eIF4G, and eIF2) and the Sch9-kinase (homologue of PKB/Akt and p70S6 in mammals), which are thought to mediate its effect on translation. Sch9 is a member of the AGC-family of protein kinases (protein kinases A, G, and C) that form one level of the signaling network downstream of TOR kinases.

In addition, TOR kinase and Lst8 are present in a second complex that is not responsive to rapamycin and contains several different subunits, encoded by AVO1, AVO2, AVO3, and BIT61 in yeast (Loewith et al., 2002 blue right-pointing triangle; Wedaman et al., 2003 blue right-pointing triangle; Reinke et al., 2004 blue right-pointing triangle). Similarly to TORC1, the general architecture of this complex is conserved in evolution, and homologous subunits corresponding to Avo1 and Avo3 have been characterized in mammals and were named hSin1 and rictor, respectively (Jacinto et al., 2004 blue right-pointing triangle; Sarbassov et al., 2004 blue right-pointing triangle). Less is known about the regulation and targets of this second TOR complex, named TOR complex 2 (TORC2). In yeast, it is required for actin organization, efficient endocytosis and normal cell polarization (Schmidt et al., 1997 blue right-pointing triangle, 1998 blue right-pointing triangle; Loewith et al., 2002 blue right-pointing triangle; deHart et al., 2003 blue right-pointing triangle; Aronova et al., 2008 blue right-pointing triangle). The effect on actin organization is conserved in mammals (Jacinto et al., 2004 blue right-pointing triangle). In addition, TORC2 participates in the regulation of sphingolipid metabolism as functional TORC2 is required for maintaining normal ceramide levels in yeast (Beeler et al., 1998 blue right-pointing triangle; Tabuchi et al., 2006 blue right-pointing triangle; Aronova et al., 2008 blue right-pointing triangle). This regulatory pathway is a component of an intricate signaling network emanating from TORC2 and the sphingolipid responsive Pkh-kinases (homologues of the mammalian PDK1) that regulate many aspects of cellular physiology through the combinatorial phosphorylation of AGC kinases, such as Ypk1/2 or Sch9 in yeast and Akt in mammals, respectively. Phosphorylation by both the TORC2 branch and the Pkh-kinases is required for full activation of AGC-kinases (Inagaki et al., 1999 blue right-pointing triangle; Kamada et al., 2005 blue right-pointing triangle; Lee et al., 2005 blue right-pointing triangle; Sarbassov et al., 2005 blue right-pointing triangle; Urban et al., 2007 blue right-pointing triangle).

The importance of both TORC1 and TORC2 for cellular physiology is highlighted by the fact that several components of each complex (Kog1, Lst8, Avo1, Avo3, and Tor2) are essential for viability of yeast.

How the two different TOR complexes achieve their different essential functions is not known. It is also unclear how cross-talk from the identical kinases is prevented and how specific signaling in response to signals for either complex is achieved. To begin to address these fundamental questions, we investigated the subcellular localization and dynamics of the TOR complexes in vivo by using S. cerevisiae as a model.

MATERIALS AND METHODS

Yeast Strains

All yeast strains and their genotypes used in this study are listed in Supplemental Table 1. AVO1-GFP::HIS, AVO2-GFP::HIS, AVO3-GFP::KANR, BIT61-GFP::HIS, LST8-GFP::HIS, and KOG1-GFP::HIS were generated in the W303 wild-type strain TWY138 by homologous recombination of polymerase chain reaction (PCR)-generated fragments as described previously (Janke et al., 2004 blue right-pointing triangle). TWY776, TWY777, and TWY778 strains were generated by PCR-mediated tagging of LSP1, with a RFPmars::NATR fragment in TWY680, TWY701, and TWY696 strains, respectively. Analogously, TWY815, TWY699, and TWY780 strains were generated by transforming SLA1-RFPmars, Abp1-RFPmars, and CRN1-RFPmars fragments with NATR marker in TWY680. TWY801 was generated by transforming PMA1-RFPmars::NATR in TWY696. AVO1-GFP::KANR was transformed in TWY779 to yield TWY808. TWY722 was similarly generated by transformation and homologous recombination of AVO3-GFP::KANR in the EDE1-RFP–harboring strain TWY367, which was described previously (Toshima et al., 2006 blue right-pointing triangle).

The wild-type strains TWY138 and TWY139 were crossed, and zygotes were pulled to obtain the diploid wild-type strain TWY806. TWY825 was generated by homologous recombination of the PCR fragment of avo1-CΔ-GFP::HIS in the diploid wild type. TWY680 was crossed with the wild-type strain TWY139 to obtain TWY849, in which a avo1-CΔ-CaaX::NATR fragment was transformed (TWY860). Sporulation and dissection yielded TWY877. This strain was then crossed with TWY779, and zygotes were pulled to obtain TWY882. Dissection of the latter yielded TWY891. TWY918 was generated by transformation and homologous recombination of NATR::GAL::GFP-avo1-C-term in TWY806.

Yeast Culture

Yeast strains were grown according to standard procedures. For growth curves, cells were diluted in 200 μl of YPD to OD600 = 0.1, and OD600 was measured every 20 min at 30°C under constant shaking (Bioscreen; Labsystems, Helsinki, Finland). Yeast spotting was performed on YPD plates, which were incubated at 24, 30, and 37°C, respectively. For vacuole membrane staining, cells were incubated 15 min on ice with 10 μM N-[3-triethylammoniumpropyl]-4-[p-diethylaminophenylhexatrienyl]pyridinium dibromide (FM4-64) (in dimethyl sulfoxide), washed three times with media, and incubated at 30°C for 2 h. For actin disassembly, cells were incubated 10 min at 30°C with 10 μM latrunculin A (LAT-A).

Microscopy

For fluorescence microscopy, cells were grown in synthetic media to OD600 = 0.6. Cells were then mounted on coverslips coated with concanavalin A. Cells were imaged with a laser-based spinning disk confocal microscope (Andor Technology, Belfast, Northern Ireland), built from an iMIC body (TiLL/Agilent), a CSU22 spinning disk (Yokogawa, Amersfoort, The Netherlands), diode pumped solid state lasers (488 and 561 nm), and a FRAPPA module (TiLL/Agilent, Gräfelfing, Germany). Filtered images (Samrock emission filters in a Sutter filter wheel) were recorded with a D-977 iXon EMCCD+ camera (Andor Technology) after twofold magnification (Andor Technology) by using a 100xTIRFM objective with a numerical aperture of 1.45 (Olympus, Tokyo, Japan), resulting in a measured pixel size of 0,086 μm at a magnification of 200×.

For surface and midsection snapshots, single frames with exposure times ranging between 400 and 500 ms were averaged four times. To examine the dynamic behavior of TORC2 components, exposure time and laser intensity were reduced to a minimum, and only two times averaging of single frames was applied. Time-lapse movies were recorded with variable delays of 1–30 s between exposures.

For quantification of the green fluorescent protein (GFP) foci size of all TORC2 components, snaps without or with two times averaging were taken from the cell surface. Identical settings were used to obtain images of the nuclear spot of 128 GFPs (TWY804) and the kinetochor cluster of Cse4-GFP and Spc105-GFP, respectively. Next, we tested whether our microscopy setup linearly records fluorescence intensity over the measurement range by determining fluorescence intensities of serial dilutions of a fluorophore with similar absorbance and emission spectra as GFP. To this end, we imaged serial dilutions of the fluorescent dye Alexa488 (Invitrogen, Carlsbad, CA) two times each. The mean fluorescence intensity of each acquired image was measured, background intensity (water sample) was subtracted, and values of the duplicate sample were averaged. Intensity values were plotted against the concentration of Alexa488 by using logarithmic scales, and a power trend line was added (Supplemental Figure 1G).

We found that the measured fluorescence intensity was linearly proportional to the amount of fluorophores. From these experiments, we calculated an arbitrary GFP fluorescence unit and related these measurements to the intensities observed for individual TORC2 foci labeled with each of the specific subunits.

Image Processing and Calculation

Images were processed using ImageJ software (http://rsbweb.nih.gov/ij/) and the MBF ImageJ for Microscopy collection of plug-ins (http://www.macbiophotonics.ca/imagej/). Single channels were separately processed using a median filter with 1-pixel radius and adjusting brightness and contrast. For time-lapse series and stacks, a median 3D filter was applied.

For quantification of the GFP foci size of all TORC2 components, the mean fluorescence intensity of each single detectable GFP foci was measured in a region of 4 × 4 pixels. The same region of interest was used to obtain mean values of the background and the nuclear background, respectively. Mean background values were averaged and subtracted from each measured value of the foci. Histograms for all values of each TORC2 component, 128 GFPs, Cse4-GFP, and Spc105-GFP were compiled using Excel (Microsoft, Redmond, WA). To get an approximate fluorescence intensity value for a single GFP, the average intensity of the 128 GFP spot was divided by 128, the average value for a Cse4-GFP cluster was divided by 32, and the average intensity of a Spc105-GFP kinetochore cluster was divided by 80. Box plots were used for comparison of the signal intensity of GFP foci (of 4 different TORC2 components) with the approximate value of a single GFP.

For quantification of the dynamic behavior of TORC2 foci, time-lapse movies (10 frames; 3-s intervals) were recorded, and the background was subtracted. For every single cell, a region as small as possible containing all Avo3-GFP foci was designed, and the area was measured. All foci were tracked manually over time using the Manual Tracking Plugin (ImageJ). Obtained data were used for further analysis of density and velocity of foci and their direction of movement using Excel (Microsoft).

Protein Expression and Purification

For bacterial expression of the HIS6-tagged avo1 C terminus, the sequence coding for the last 123 amino acids of AVO1 were amplified by polymerase chain reaction (PCR) from purified yeast genomic DNA and cloned into a pETM14-ccdB vector. Positive plasmids were identified by sequencing and transformed in Escherichia coli BL21 (DE3) Rosetta.

The N-terminal HIS6-tagged avo1 C-term was expressed and purified by nickel-nitrilotriacetic acid chromatography (QIAGEN, Hilden, Germany), followed by a buffer exchange step using HiTrap desalting column (GE Healthcare, Little Chalfont, Buckinghamshire, United Kingdom). The protein was stored in buffer containing 150 mM NaCl, 50 mM Tris, pH 8.0, and 10% glycerol.

Liposome Preparation

Liposomes were made of either 100% dioleoyl phosphatidylcholine (DOPC, Avanti Polar Lipids), or DOPC combined in a 97:3 molar ratio with dioleoyl phosphatidylserine (DOPS, Avanti Polar Lipids), dipalmitoyl phosphatidylinositol (DPPI), dipalmitoyl phosphatidylinositol-3-phosphate (DPPI(3)P), dipalmitoyl phosphatidylinositol-(4)-phosphate (DPPI(4)P), dipalmitoyl phosphatidylinositol-3,4-phosphate (DPPI(3,4)P2) and dipalmitoyl phosphatidylinositol-4,5,-phosphate (DPPI(4,5)P2) (all Matreya, State College, PA), respectively. Lipids were dissolved in CHCl3-ethanol (1:1, vol/vol) with 0.1% HCl and vortexed in a round-bottomed glass flask. Lipids were mixed according to the above-mentioned molar ratio to a final concentration of 25 mM. The solvent was evaporated under constant stream of argon gas for 10 min and subsequently dried in a desiccator for 4 h. Dried lipids were hydrated in 150 mM NaCl, 50 mM Tris, pH 8.0, and 10% glycerol and vortexed extensively until solution was clear. After three freeze thaw cycles in liquid nitrogen, lipids were extruded (Mini extruder; Avanti Polar Lipids) 15 times through a 100-nm filter (Whatman, Maidstone, United Kingdom) and kept at 4°C.

Liposome Flotation Assay

Purified HIS6-avo1 C-term (10 μM) was incubated in the absence or presence of 4 mM lipids in liposomes in a final volume of 50 μl for 1 h at 4°C. The liposome flotation assay was performed in a discontinuous sucrose gradient as described previously (Narayan and Lemmon, 2006 blue right-pointing triangle). After incubation, 50 μl of 80% sucrose in 150 mM NaCl, 50 mM Tris pH 8.0, and 10% glycerol was added to the samples, mixed, and overlaid with 300 μl of 30% sucrose in the same buffer. The gradient was centrifuged at 55,000 rpm for 1 h at 4°C. Fractions of 50 μl were taken from the top and the bottom and analyzed by SDS-polyacrylamide gel electrophoresis (PAGE) (4–12% gradient NuPAGE; Invitrogen), Coomassie staining, and silver staining.

RESULTS

TORC1 and TORC2 Complexes Localize to Distinct Cellular Compartments

Previous experiments investigating the subcellular localization of TOR complexes have relied on immune detection of tagged components after subcellular fractionation or electron microscopy and indirect immune fluorescence (Cardenas and Heitman, 1995 blue right-pointing triangle; Kunz et al., 2000 blue right-pointing triangle; Chen and Kaiser, 2003 blue right-pointing triangle; Wedaman et al., 2003 blue right-pointing triangle; Aronova et al., 2007 blue right-pointing triangle). Because these techniques do not allow the visualization of protein localization with respect to other organelles in live cells, we aimed to fluorescently tag components of TORC1 and TORC2. We chose the yeast S. cerevisiae as a model system because genomic tagging in this organism allows one to express only the fluorescently labeled protein from their endogenous promoter. To follow the localization of both TOR complexes independently, we fused GFP tags to the exclusive subunits of TORC2 (Avo1, Avo2, Avo3, and Bit61), the TORC1-specific Kog1, and the shared subunit Lst8. The resulting strains expressing C-terminally GFP-tagged TOR complex components all grew normally at 30°C (Figure 1a, left). In contrast, C-terminally tagged alleles of TOR2 that can be found in both complexes were not viable (data not shown), consistent with a recent report (Sturgill et al., 2008 blue right-pointing triangle).

Figure 1.
Functional GFP-tagged TORC1 and TORC2 subunits localize to the vacuolar and the plasma membrane, respectively. (a) GFP-tagged TORC2 subunits Avo1, Avo2, Avo3, and Bit61 support normal yeast growth, whereas LST8-GFP and KOG1-GFP are temperature-sensitive ...

Because TOR kinases react to stress, we also tested whether the tagged TOR complex subunits support growth at elevated temperature. Under heat stress at 37°C, all cells harboring tagged subunits of the TORC2 grew as well as the wild-type strain, but mutants carrying Kog1-GFP or Lst8-GFP did not grow (Figure 1a, right). This might indicate that the GFP interferes with the essential interaction of either Lst8 or Kog1 with the TOR kinase (Kim et al., 2003 blue right-pointing triangle; Wullschleger et al., 2005 blue right-pointing triangle).

In summary, GFP-tagged versions of essential Avo1 and Avo3 are fully functional, whereas KOG1-GFP and LST8-GFP are novel temperature-sensitive alleles of the TORC1 complex and both TOR complexes, respectively.

Initial imaging of TOR complexes on a conventional epifluorescence microscope showed only dim and barely detectable signals. To analyze the subcellular localization of TOR complexes, we therefore made use of recent advances in fluorescence microscopy, by using a highly sensitive electron multiplying charge-coupled device (EM-CCD) camera (99% quantum yield) combined with a spinning disk confocal microscope, a high numerical aperture objective (1.45 numerical aperture), and high optical magnification (200× after all components; see Materials and Methods).

Images of exponentially growing cells expressing Kog1-GFP or Bit61-GFP revealed distinct localizations of both TOR complexes. Whereas TORC1 marked by Kog1-GFP localized exclusively to one or a few central rings in all cells (100% of cells; n > 200), which we suspected were vacuoles (Figure 1b, left), TORC2 labeled by Bit61-GFP localized in a peculiar, punctate pattern at the cell surface forming small foci in all cells observed (100%, n > 200; Figure 1b, right). The localization of neither TORC1 nor TORC2 components was dependent on growth conditions (Berchtold and Walther, unpublished observation). Consistent with its localization in both TOR complexes, Lst8-GFP was found in a pattern overlaying the two localizations at the cell periphery and the central rings (Figure 1b, middle). However, fewer foci were found at the cell periphery in Lst8-GFP compared with Bit61-GFP–expressing cells. The formation of fewer spots at the cell periphery might indicate that the GFP-tagged Lst8 protein does not get incorporated into TORC2 as efficiently as the wild type, consistent with the reduced functionality of LST8-GFP seen in growth assays at 37°C.

TORC1 Localizes to the Yeast Vacuole

To determine to which site in the cell the TOR complexes are localized, we performed double labeling experiments with different organelle markers. Because the TORC1 component Kog1-GFP localized in one or a few rings in the cell interior, suggesting a vacuolar localization, we tested whether the Kog1-GFP signal was located at the vacuolar delimiting membrane. Indeed, the Kog1-GFP signal colocalized completely with the vital dye FM4-64 used to label the vacuolar membrane (Figure 1c), confirming previous findings (Araki et al., 2005 blue right-pointing triangle; Huh et al., 2003 blue right-pointing triangle). Similarly, we observed a precise overlay for the ring-like Lst8-GFP signal in the center of the cells and vacuolar FM4-64 (data not shown), as it was proposed previously (Chen and Kaiser, 2003 blue right-pointing triangle). This confirms that TORC1 indeed localizes to the vacuolar membrane in live cells, a finding supported by a report that the nonconserved TORC1 subunit Tco89 is also localized to the vacuolar membrane (Reinke et al., 2004 blue right-pointing triangle; Urban et al., 2007 blue right-pointing triangle).

TORC2 Localizes to the Plasma Membrane

TORC2 foci marked by Bit61-GFP localize exclusively in the cell periphery (Figure 1b, right). To test whether Bit61 localizes to the plasma membrane, we expressed a red fluorescently tagged plasma membrane protein Pma1-RFPmars in Bit61-GFP–expressing cells and imaged them by fluorescence microscopy. Midsection images show that Bit61-GFP foci completely overlap with the plasma membrane signal of Pma1-RFPmars (Figure 1d), confirming that Bit61-GFP is a plasma membrane protein. From these data, we conclude that the two TOR complexes have distinct and mutually exclusive localizations in the cell.

All TORC2 Components Localize As Oligomeric Foci to the Plasma Membrane

To investigate whether the other subunits of the TORC2 complex are localized in a pattern equivalent to Bit61, we analyzed fluorescence images of all tagged members of TORC2. Midsection (Figure 2a, left) and cell surface images (Figure 2a, right) revealed that they all localize to the plasma membrane in small foci of identical appearance.

Figure 2.
All subunits of TORC2 localize to the plasma membrane in a similar pattern. (a) All exclusive TORC2 subunits show a similar pattern at the plasma membrane. Strains carrying the indicated subunits of TORC2 as the only allele of the corresponding gene were ...

To estimate the amount of each protein in a TORC2 focus, we related the GFP signal from single foci to a GFP standard. For this, we first analyzed the fluorescence intensity signal from GFP molecules bound to the Lac-operator in the nucleus that form one spot (Brickner and Walter, 2004 blue right-pointing triangle). For each of the components, we observed a very similar median fluorescence. In summary, for all components, this value corresponds to 3.8 ± 0.4 GFPs (Figure 2b). In histograms of fluorescence intensities, we observed a trail in the distribution toward higher fluorescence intensities that might indicate formation of complexes with more subunits (see Supplemental Figure 1, A–F). To confirm these conclusion, we also GFP-tagged kinetochore proteins Cse4 and Spc105 as markers to quantify in vivo protein numbers (Joglekar et al., 2008 blue right-pointing triangle). One kinetochore cluster of Cse4-GFP represents the fluorescence of 32 GFP molecules, whereas a kinetochore cluster of Spc105-GFP represents 80 GFP molecules. From this experiment, the median fluorescence of the four measured TORC2 components corresponded to 3.3 ± 0.4 GFPs (see Supplemental Figures 2 and 3, a–h), consistent with our measurements using the Lac-operator. Together, these data suggest that TORC2 foci at the plasma membrane represent higher order oligomers, with two to six copies of each subunit present in each focus.

Plasma Membrane TORC2 Is Highly Dynamic

During our analysis of TORC2 foci, we observed that their plasma membrane pattern is highly dynamic. To characterize this behavior in more detail, we recorded time-lapse movies of cells expressing Avo3-GFP. The movies showed that within 10 s, the surface pattern of TORC2 changes almost completely (Figure 3a and Supplemental Movie 1). Further inspection of movies revealed that TORC2 complexes display random short-range lateral movements. We analyzed this movement and tracked individual TORC2 foci by using movies with 3-s intervals (Supplemental Movie 2). Figure 3b shows three such tracks from the first frame of the movies to the disappearance of the foci. Individual foci have quite a different overall behavior, but similar speeds of their movement, with an average of 0,039 μm/s (quantitated from n = 141 tracks). We only observed movement in the plane of the membrane and never detected detachment of a TORC2 focus from the plasma membrane. Besides the lateral movement four additional events contribute to the highly dynamic pattern: 1) appearance, 2) disappearance, 3) fusion, and 4) splitting of foci (Figure 3d and Supplemental Movies 3–6). Fusion was accompanied by doubling of fluorescence intensity, whereas splitting of foci resulted in a decrease of fluorescence (Figure 3d). To quantitate the behavior, we counted the number of spots occurring per time and found that TORC2 foci formed with a rate of 0.61 per μm2 membrane surface and minute. Consistent, but slightly higher, results were obtained for the rate of foci disappearance, possibly due to bleaching of some spots (data not shown). In contrast, fusion and splitting of the foci were much less frequent and only occurred in a minority of image sequences.

Figure 3.
TORC2 localization at the plasma membrane is highly dynamic. (a) The TORC2 pattern at the plasma membrane changes rapidly. Surface images of cells expressing Avo3-GFP were collected in 400-ms intervals. The image t = 0 s is shown in green (left); the ...

To confirm the appearance and disappearance of TORC2 foci independently, we bleached one-half of the cell and followed fluorescence recovery over time. In optical midsections (Figure 3f and Supplemental Movie 7) and top sections (data not shown), TORC2 foci reappeared in a previously bleached area. In few cases, these could be attributed to migration of foci from the unbleached area of the cell. In most cases, however, the TORC2 focus occurred de novo. This basic characterization demonstrates that TORC2 focus formation is dynamic. To begin to analyze the molecular requirements for TORC2 localization, we performed analogous analysis of cells expressing Avo3-GFP in which we deleted either AVO2 or BIT61. TORC2 plasma membrane signals were not changed as measured by the density, velocity, and intensity of Avo3 foci, showing that these nonessential subunits are not required for TORC2 localization and dynamics. For bit61Δ cells, the rate of appearance of the TORC2 foci was elevated slightly, maybe indicating that Bit61 stabilizes the interaction with the plasma membrane (Supplemental Figure 4).

TORC2 Foci at the Plasma Membrane Are Distinct from Actin Patches

TORC2 regulates actin in both yeast and mammalian systems. In yeast, actin localizes in patches at the cell cortex and forms actin cables, found mostly in the cell interior. Actin patches mediate at least a subset of endocytic events. Because TORC2 is also required for efficient endocytosis (deHart et al., 2003 blue right-pointing triangle), we hypothesized that TORC2 foci are part of actin patches. To tests this, we analyzed the localization of Avo3-GFP in cells expressing different fluorescently labeled actin patch proteins. Because actin patches are highly dynamic structures of changing protein composition, we selected several proteins that are recruited at the early (Ede1, Sla1), later (Crn1), and at final stage (Abp1) of an actin patch assembly cycle (Kaksonen et al., 2003 blue right-pointing triangle, 2005 blue right-pointing triangle). Consistent with previous findings by Wedaman et al. (2003) blue right-pointing triangle, we did not find any colocalization of Avo3-GFP with any of these proteins, either in surface snapshots (Figure 4a) or when we followed actin patches over time (Supplemental Movies 8 and 9). Furthermore, TORC2 did not localize at the sites of actin patch formation before or after their residence at the plasma membrane (Supplemental Movies 8 and 9). Thus, TORC2 is not a component of actin patches.

Figure 4.
TORC2 foci are different from actin patches. (a) TORC2 foci do not colocalize with actin patches. Avo3-GFP (green) was expressed in cells expressing either Sla1, Ede1, Crn1, or Abp1 tagged with the RFPmars fluorophore (red). Representative surface images ...

To determine whether the movement of TORC2 at the plasma membrane is dependent on actin, we incubated cells with the actin depolymerizing drug latrunculin A. This treatment efficiently disassembled actin patches as expected. For example, Abp1-RFPmars lost its normal patch localization and was present in a diffuse cytoplasmic signal (Figure 4b, top right). When we investigated the motility of TORC2 under these conditions, we did not detect any changes compared with untreated controls, i.e., neither TORC2 localization at the plasma membrane nor its dynamics were significantly changed compared with untreated controls (Figure 4b, bottom). From these results, we conclude that the TORC2 foci are independent from actin patches and that actin is not required for TORC2 motility.

TORC2 Localizes in a Distinct Plasma Membrane Domain

To further characterize the membrane environment of TORC2, we asked whether it localizes to a specific membrane subcompartment. The plasma membrane of S. cerevisiae is organized in two mutually exclusive membrane domains that are composed of different lipids and proteins. One of these domains is organized by large cytosolic protein complexes termed eisosomes and harbors several transporter proteins, such as Can1 (Malinska et al., 2003 blue right-pointing triangle; Malinska et al., 2004 blue right-pointing triangle; Walther et al., 2006 blue right-pointing triangle; (Grossmann et al., 2007 blue right-pointing triangle). Accordingly, it was termed membrane compartment occupied by Can1 (MCC). The MCC is organized by large protein complexes termed eisosomes that localize there beneath the plasma membrane and consist of two major subunits, Pil1 and Lsp1. Deletion of Pil1 results in the collapse of the MCC to one or a few clusters at the plasma membrane (Grossmann et al., 2006 blue right-pointing triangle; Walther et al., 2006 blue right-pointing triangle). Besides several specific proteins, the MCC also has a distinct lipid composition, as visualized by accumulation of the sterol binding fluorescent dye filipin (Grossmann et al., 2007 blue right-pointing triangle). A mutually exclusive domain is characterized by the presence of Pma1 and was therefore named membrane compartment occupied by Pma1 (MCP).

To determine to which of these plasma membrane domains TORC2 localizes, we imaged surfaces of cells expressing one of the TORC2-specific subunits fused to GFP and the RFPmars-tagged eisosome component Lsp1. We never observed colocalization between the two proteins (Figure 5a). This is readily apparent in fluorescence intensity line profiles showing clearly separated peaks for each channel (Figure 5b). Simultaneous imaging of GFP-tagged TORC2 subunits and Lsp1-RFPmars over time clearly shows that TORC2 foci do not overlap with eisosomes during the course of the experiment (Supplemental Movie 10). Consistent with these observations, TORC2 localizes normally in pil1Δ cells, where the MCC collapses to one or a few remnants per cell (Supplemental Figure 5; Walther et al., 2006 blue right-pointing triangle; Grossmann et al., 2007 blue right-pointing triangle).

Figure 5.
TORC2 foci localize to distinct plasma membrane domains. (a) TORC2 foci localize to a plasma membrane domain distinct from MCC/eisosomes. The indicated subunits of TORC2 were expressed as GFP fusion proteins (green) in cells also expressing a RFPmars-tagged ...

Because the MCC is mutually exclusive with MCP, our data predicted that TORC2 would occupy the latter plasma membrane domain. To test this directly, we analyzed the localization of Bit61-GFP in cells expressing Pma1-RFPmars. To our surprise, we found that TORC2 marked by Bit61-GFP localized in areas excluding Pma1 (Figure 5c). This is particularly evident in fluorescence intensity line profiles through the cell surface (Figure 5d). Instead of being a component of MCP, TORC2 occupied domains that where devoid of Pma1 (Figure 5c, top, see magnified inset). In addition, we always observed plasma membrane regions devoid of fluorescence signal from either Pma1 or TORC2, and thus presumably representing MCC (Figure 5c, bottom, see magnified inset). TORC2 therefore defines a previously unrecognized plasma membrane compartment that we name membrane compartment containing TORC2 (MCT).

Plasma Membrane Targeting of TORC2 through the C-Terminal Domain of Avo1 Is Essential

To investigate the function of TORC2 plasma membrane binding and to begin analyzing the mechanism of TORC2 membrane binding, we wanted to identify its membrane anchor. From sequence inspection, homology modeling, and previous studies in mammalian systems, we suspected that the C terminus of Avo1 could potentially form a Pleckstrin homology (PH)-like domain (Schroder et al., 2007 blue right-pointing triangle; and Ziolkowska, Berchtold, and Walther, unpublished observation). PH domains most often bind phosphoinositide-(4,5)-bisphosphate [PI(4,5)P2], a plasma membrane-specific lipid in yeast (Lemmon, 2008 blue right-pointing triangle; Odorizzi et al., 2000 blue right-pointing triangle). Therefore, the C terminus of Avo1 was a good candidate to mediate TORC2 plasma membrane anchoring.

To test whether the C terminus of Avo1 is required for localization and/or function of TORC2, we deleted the last 117 amino acids of Avo1 in one of the genomic copies of diploid cells. Analysis of the haploid progeny from a sporulation of these heterozygous cells showed that only spores with the wild-type allele of AVO1 were viable, whereas the avo1-CΔ allele was lethal (Figure 6a). The C terminus of Avo1 is therefore essential for yeast viability.

Figure 6.
Plasma membrane localization of TORC2 is essential. (a) The C-terminal domain of Avo1 is essential. The C-terminal 117 amino acids of Avo1 were deleted in a diploid strain; the resulting heterozygote sporulated and the progeny were dissected. A representative ...

The requirement for the C terminus could be explained by at least three alternative hypotheses: it could be necessary for stability or folding of Avo1, its stable incorporation into TORC2, or if it indeed formed a PH-domain, for lipid binding and its localization to the plasma membrane. To distinguish between these possibilities, we tested whether replacing the C terminus by a heterologous plasma membrane targeting signal was sufficient to support yeast growth. To achieve this, we replaced the C terminus of Avo1 with a CaaX-motif, which gets enzymatically lipidated (Onken et al., 2006 blue right-pointing triangle). Surprisingly, given that the Avo1 C terminus is required for viability, cells harboring the deletion allele replaced by a short CaaX-box had no discernible growth defect under any condition investigated, as seen for example in solid media growth assays (Figure 6b) or in growth curves obtained from liquid cultures (data not shown).

To examine the dynamics of artificially plasma membrane targeted TORC2 complexes, we investigated avo1-CΔ-CaaX cells with GFP-tagged Avo3. To our surprise, the localization and dynamics of TORC2 complexes targeted to the plasma membrane by the CaaX-box membrane anchor was indistinguishable from wild type, both in the pattern of the foci (Figure 6c), their size (Figure 6d), their density on the plasma membrane and their dynamics of movement (Figure 6f), and turnover (Figure 6g). From these data, we conclude that the C-terminal domain of Avo1 mediates the essential plasma membrane targeting of TORC2.

To further determine whether the Avo1 C-terminal domain is sufficient for plasma membrane targeting in vivo, we fused the avo1 C-term sequence in frame with the C terminus of GFP and expressed it from the GAL promoter. After induction of GFP-avo1 C-term expression, we detected cytoplasmic GFP fluorescence and accumulation of GFP signal at the plasma membrane (Figure 7a, left). This was corroborated by line scans through midsections of confocal images (Figure 7a, right).

Figure 7.
The C-terminal domain of Avo1 binds to plasma membrane PI(4,5)P2. (a) The C-terminal domain of Avo1 is sufficient to mediate plasma membrane targeting. The C-terminal 117 amino acids of Avo1 were fused to GFP and expressed from the Gal promoter. A representative ...

We next asked whether the C-terminal sequence of Avo1 directly mediates lipid binding, as its similarity to a PH-domain suggests. Because PH-domains often bind PI(4,5)P2 or other phosphoinositides, we focused on these lipids. To investigate a possible lipid interaction, we expressed the Avo1 C-terminal domain (amino acids 1054–1176) as a 6xHis-fusion in E. coli, purified it to apparent homogeneity, and performed liposome flotation experiments. Figure 7b shows that Avo1 C-terminal domain floated to the top of a sucrose gradient (fraction T) bound to phosphatidylcholine (PC) liposomes containing 3% PI(4,5)P2, and to a much lesser extent to liposomes containing 3% PI(3,4)P2. No flotation of the Avo1 C-terminal domain was observed in reactions containing no liposomes or liposomes containing only PC, PC/3% phopsphatidylinositol (PI), PC/3% phosphatidylserine (PS), PC/3% phosphatidylinositide (3) phosphate [PI(3)P], or PC/3% phosphatidylinositide (4) phosphate [PI(4)P] (Figure 7b). Similar results were obtained with an independently generated glutathione transferase-tagged construct (Berchtold and Walther, data not shown).

Together, our in vivo data, biochemical observations, and the structural homology strongly suggest that the Avo1 C terminus forms a PH-like domain that can bind PI(4,5)P2 and is sufficient and necessary for plasma membrane targeting of TORC2.

DISCUSSION

TOR kinases are important regulators of cellular physiology and exist in two distinct complexes. We show that TORC2 localizes exclusively at the plasma membrane, segregated from TORC1, which is found only at the vacuolar membrane. TORC2 did not colocalize with previously known functional membrane compartments and instead defines a distinct compartment that we name the MCT.

The spatial segregation of TOR kinases in different complexes is conceptually similar to the targeting of protein kinase A (PKA) to a variety of subcellular localizations (Smith et al., 2006 blue right-pointing triangle). In that case, the same PKA is recruited by A-kinase anchoring proteins, which are in turn bound to different organelles or the cytoskeleton. This segregation helps to provide specificity for G protein-coupled signals that relay information for example on growth factors availability via cAMP to PKA. Analogously, one could see Avo1 as a TOR plasma membrane anchoring protein within TORC2. Similarly, TORC1 must contain (a) targeting subunit(s) for the vacuole. Because Kog1 is the only TORC1-specific component that is conserved through evolution, it is a good candidate for such an anchor.

Scaffolding and subcellular segregation of kinase complexes is a recurrent theme in signal transduction. In yeast, also the mitogen-activated kinase kinase (MAKK) Ste11 is present in two complexes that mediate the response to pheromone and changes in osmolarity, respectively. Again, this segregation is important for signaling specificity and to prevent cross talk between pathways (Schwartz and Madhani, 2006 blue right-pointing triangle). From these examples, it seems likely that one function of TOR anchoring to the plasma membrane and the vacuole is to separate the output of the kinases. Consistent with this notion, the TORC2 target Ypk1 is also localized to the plasma membrane, but not to the vacuolar membrane (Berchtold and Walther, unpublished observation; Sun et al., 2000 blue right-pointing triangle). Conversely, the TORC1 target Sch9 is localized exclusively to the vacuolar surface (Jorgensen et al., 2004 blue right-pointing triangle; Urban et al., 2007 blue right-pointing triangle). Because both of these target kinases are quite similar (44% identity) and could therefore be recognized by each TORC, their separation is likely required to ensure signaling fidelity.

In addition, TOR localization might be important for the detection of input signals. Consistent with its function in nutrient sensing, we found TORC1 localized exclusively to the yeast vacuole, which is a major nutrient reservoir in yeast. This is in agreement with previous data, reporting Tor2 kinase (a member of both TORC1 and TORC2) localization at the vacuolar membrane (Cardenas and Heitman, 1995 blue right-pointing triangle). Indeed, several studies underscore a prominent role for vacuolar functions in TORC1 signaling (Aronova et al., 2007 blue right-pointing triangle; Zurita-Martinez et al., 2007 blue right-pointing triangle; Puria et al., 2008 blue right-pointing triangle). Contrary to previous reports, we did not detect Kog1-GFP in the nucleus or the plasma membrane (Aronova et al., 2007 blue right-pointing triangle; Tsang and Zheng, 2007 blue right-pointing triangle). However, our results do not exclude the possibility of a small nondetectable, but biologically significant pool of Kog1 at those locations or that TORC1 gets targeted to other locations after altering conditions. At the vacuolar membrane, TORC1 patches are less pronounced than those of TORC2. Together with the high mobility of yeast vacuoles and the lack of spatial landmarks in the vacuolar membrane, this precludes further analysis of TORC1 dynamics.

Compared with TORC1, far less is known about the input signals of TORC2. Because it is required for maintaining normal ceramide levels (Beeler et al., 1998 blue right-pointing triangle; Tabuchi et al., 2006 blue right-pointing triangle; Aronova et al., 2008 blue right-pointing triangle), one model suggests that TORC2 senses ceramide levels to maintain their level in a feedback loop. In this scenario, its localization at the plasma membrane where most cellular ceramides are localized could be important for sensing. However, ceramides are mostly present in the outer leaflet of the plasma membrane (van Meer and Lisman, 2002 blue right-pointing triangle), so it is unlikely that TORC2 bound to the cytoplasmic face is regulated by directly binding ceramides. Ceramides are the major class of sphingolipids in yeast and are thought to form specialized liquid ordered plasma membrane domains together with ergosterol, the main yeast sterol (Simons and Ikonen, 1997 blue right-pointing triangle; Bagnat and Simons, 2002 blue right-pointing triangle). TORC2 could therefore indirectly sense sphingolipid levels by responding to alterations in plasma membrane structure.

At the plasma membrane, TORC2 localizes to neither of the previously recognized plasma membrane domains (Malinska et al., 2003 blue right-pointing triangle, 2004 blue right-pointing triangle; Grossmann et al., 2006 blue right-pointing triangle; Walther et al., 2006 blue right-pointing triangle) but instead occupies a distinct compartment that we name MCT. The TORC2 foci at the plasma membrane represent higher oligomeric forms of the complex. One possible model of TORC2 assembly and dynamics is that individual TORC2 complexes are bound to the plasma membrane where they interact to form dynamic higher order complexes. This model is supported by our observation that TORC2 foci are formed, fall apart, and can fuse and segregate.

The formation of TORC2 foci might be important for the regulation of its activity because multimerization was correlated previously with kinase activity biochemically (Wullschleger et al., 2005 blue right-pointing triangle). TORC2 foci formation in the MCT is probably mediated by a mechanism distinct from its membrane anchoring by PI(4,5)P2, because targeting to the plasma membrane by a heterologous lipid anchor also results in MCT localization.

How many proteins are exclusively targeted to MCT, MCP, or MCC and whether these domains are really segregated or represent examples from a continuous spectrum of behaviors of plasma membrane proteins are not yet clear. It is also not clear whether MCC, MCP, and MCT have distinct lipid compositions. The MCC was shown previously to be enriched in ergosterol, as visualized by the fluorescent sterol-binding dye filipin (Grossmann et al., 2007 blue right-pointing triangle). Also, the MCP might be rich in these lipids as its highly abundant Pma1 component is used as a marker for lipid rafts, lateral membrane domains rich in ergosterol and sphingolipids (Bagnat et al., 2000 blue right-pointing triangle; Lee et al., 2002 blue right-pointing triangle). It is therefore possible that only the MCT represents the nonraft plasma membrane domain in yeast, enriched in phospholipids, but poor in sterols and sphingolipids. Consistent with this notion, proteins lipidated at a CaaX-box similar to the one targeting TORC2 to the MCT, were found previously in such raft-deficient membrane domains (Zacharias et al., 2002 blue right-pointing triangle).

CaaX-box anchored TORC2 was constitutively bound to the membrane, presumably due to embedding of the lipid moiety in the bilayer. This artificially anchored TORC2 is fully functional and its dynamics are indistinguishable from the wild type, arguing that TORC2 does not have to go through association/dissociation cycles with the plasma membrane for its function. Consistently, we never observed dissociation of wild-type TORC2 foci from the membrane into the cytoplasm.

Downstream of TORC2 at least some signals are transduced to the conserved Ypk-kinases, which also localize to the plasma membrane (Kamada et al., 2005 blue right-pointing triangle; Berchtold and Walther, unpublished observation). For full activation, Ypk-kinases need both phosphorylations by TORC2 at the conserved HM-site and by Pkh-kinases at the activation loop (Kamada et al., 2005 blue right-pointing triangle). Even though both TORC2 and Pkh-kinases localize to the cell cortex, they are in different compartments: Pkh-kinases were previously found in the MCC/eisosomes, separated from the MCT (Walther et al., 2007 blue right-pointing triangle). This sequestration of kinases of the same signaling network into distinct plasma membrane compartments provides opportunity for an additional level of regulation. It also necessitates communication between compartments to achieve complete phosphorylation of targets such as Ypk2. In the case of the TORC2/Pkh-signaling network, this regulation could be mediated at least in part by the Slm-proteins that are targets of both TORC2 and Pkh-signaling (Audhya et al., 2004 blue right-pointing triangle; Fadri et al., 2005 blue right-pointing triangle; Daquinag et al., 2007 blue right-pointing triangle). Slm-proteins are required for some of the output of the network, e.g., actin polarization, placing them downstream of TORC2/Pkh-kinases in the network (Audhya et al., 2004 blue right-pointing triangle; Daquinag et al., 2007 blue right-pointing triangle). The notion that Slm-proteins mediate between the TORC2 and Pkh-branch of the signaling network is supported by their localization in both the MCC/eisosomes and also outside these domains in an MCT pattern (Berchtold and Walther, unpublished observation). Further work will be needed to determine their role in TORC2/Pkh signaling.

Interestingly, the Pkh-kinases are regulated by sphingolipids (specifically sphingoid long chain bases (Friant et al., 2001 blue right-pointing triangle; Liu et al., 2005 blue right-pointing triangle). This suggests that the TORC2/Pkh-kinase network integrates information on sphingolipid levels and transduces the information via Ypk-kinases to regulate many cellular functions, such as lipid metabolism, cell wall integrity, and actin polarization. Ceramides and long-chain bases often have opposing effects on cells. The balance of their amounts is therefore critical and was termed the ceramide/long chain base rheostat. The physical basis of this control mechanism might be the TORC2/Pkh-kinase signaling module.

The overall features of this signaling network and the TOR complex architecture are evolutionary conserved (Casamayor et al., 1999 blue right-pointing triangle; Jacinto et al., 2004 blue right-pointing triangle; Sarbassov et al., 2004 blue right-pointing triangle). Therefore, it is likely that our findings are also relevant for higher eukaryotes. Indeed, it was observed that TORC2 can localize to the plasma membrane in human embryonic kidney 293 cells (Schroder et al., 2007 blue right-pointing triangle). Whether and how the homologous mTORC2 and PDK1 signaling complexes are organized spatially at the plasma membrane, particularly in situations relevant to disease, such as insulin signaling, remains a fascinating topic for future research.

Supplementary Material

[Supplemental Materials]

ACKNOWLEDGMENTS

We thank Robert V. Farese, Pablo Mardones, Pablo Aguilar, and Karen Moreira for critical reading of the manuscript and members of the Walther laboratory for discussions. We thank Roland Wedlich-Söldner for the generous gift of pFA6aNATR-RFPmars. This work was supported by the Max Planck Society and a career development award of the International Human Frontiers Science Program (to T.C.W.).

Footnotes

This article was published online ahead of print in MBC in Press (http://www.molbiolcell.org/cgi/doi/10.1091/mbc.E08-10-1001) on January 14, 2009.

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