Developmental Cell
Short Article A Dynamic Interface between Vacuoles and Mitochondria in Yeast Yael Elbaz-Alon,1 Eden Rosenfeld-Gur,2 Vera Shinder,3 Anthony H. Futerman,2 Tamar Geiger,4 and Maya Schuldiner1,* 1Department
of Molecular Genetics, Weizmann Institute of Science, Rehovot 7610001, Israel of Biological Chemistry, Weizmann Institute of Science, Rehovot 7610001, Israel 3Electron Microscopy Unit, Weizmann Institute of Science, Rehovot 7610001, Israel 4Department of Human Molecular Genetics and Biochemistry, Sackler Faculty of Medicine, Tel Aviv University, Tel Aviv 6997801, Israel *Correspondence: [email protected] http://dx.doi.org/10.1016/j.devcel.2014.06.007 2Department
SUMMARY
Cellular life depends on continuous transport of lipids and small molecules between mitochondria and the endomembrane system. Recently, endoplasmic reticulum-mitochondrial encounter structure (ERMES) was identified as an important yet nonessential contact for such transport. Using a high-content screen in yeast, we found a contact site, marked by Vam6/Vps39, between vacuoles (the yeast lysosomal compartment) and mitochondria, named vCLAMP (vacuole and mitochondria patch). vCLAMP is enriched with ion and aminoacid transporters and has a role in lipid relay between the endomembrane system and mitochondria. Critically, we show that mitochondria are dependent on having one of two contact sites, ERMES or vCLAMP. The absence of one causes expansion of the other, and elimination of both is lethal. Identification of vCLAMP adds to our ability to understand the complexity of interorganellar crosstalk. INTRODUCTION Mitochondria generate the majority of cellular energy and house enzymes required for the synthesis, breakdown, and interconversion of various species of amino acids, lipids, iron/sulfur clusters, and other small molecules. Due to their diverse functions and essential roles in cellular metabolism, mitochondria serve as hubs for signaling in events such as growth, differentiation, or cell death. Loss of optimal mitochondrial activity is therefore, not surprisingly, implicated in a growing number of human diseases as well as in aging (Bratic and Larsson, 2013; Costa and Scorrano, 2012; Griffiths, 2012; Vives-Bauza and Przedborski, 2011; Yu et al., 2012). The central tasks of mitochondria in cells necessitate constant communication and transport of small molecules with other organelles. However, mitochondria are not connected to the endomembrane system via the vesicular pathway. Instead, mitochondria have been shown to communicate with the endomembrane system by virtue of a membrane contact site (MCS) where membranes of mitochondria come into close proximity to membranes of the endoplasmic reticulum (ER). This MCS, also
termed mitochondria-associated-membranes (Achleitner et al., 1999; Vance, 1990), enables ions and lipids to be rapidly transported in a nonvesicular manner (Elbaz and Schuldiner, 2011; Levine and Loewen, 2006; Tatsuta et al., 2014). Understanding the molecular machineries creating and regulating this MCS has been the arena of intense investigations in the past decade. In yeast, the molecular identity of the ER-mitochondria tethering complex was recently uncovered and is mediated by a four-protein complex termed the ER-mitochondria encounter structure (ERMES) (Kornmann et al., 2009). One of its hypothesized functions was to enable phospholipid transport (Kopec et al., 2010; Kornmann and Walter, 2010) required for building mitochondrial membranes as well as for the three-step biosynthetic pathway of aminoglycerophospholipids. Therefore, it was of great surprise when the loss of ERMES subunits had very little effect on cellular levels of aminoglycerophospholipids (Kornmann et al., 2009; Nguyen et al., 2012; Voss et al., 2012). Hence, it became clear that alternate routes of phospholipid transport must exist in the cell, and uncovering them should shed light on novel modes of communication between mitochondria and the endomembrane system. We report here our findings of an MCS between mitochondria and vacuoles (the yeast lysosomal compartment) and its functional significance in lipid transport between the endomembrane system and mitochondria. RESULTS A High-Content Screen Uncovers that Vam6/Vps39 Influences the Number of MCSs between the ER and Mitochondria We set out to uncover alternate paths for communication between mitochondria and the endomembrane system using a systematic genetic screening approach. We reasoned that the extent of ERMES mediated contact sites must be regulated so that loss of a parallel pathway would result in an increase in ERMES mediated contacts. Therefore, we screened for genetic backgrounds yielding a pronounced increase in the amount of ERMES foci per cell (Figure 1A). Using the synthetic genetic array (SGA) methodology (Cohen and Schuldiner, 2011; Tong and Boone, 2006; Tong et al., 2001), we created a library in which each strain harbors a green fluorescent protein (GFP)tagged ERMES subunit (Mdm34-GFP) on the background of a single gene mutation (Breslow et al., 2008; Giaever et al., Developmental Cell 30, 95–102, July 14, 2014 ª2014 Elsevier Inc. 95
Developmental Cell A Dynamic Mitochondria/Vacuole Interface
A
Figure 1. A Genome-wide Screen to Uncover Mutants Affecting the Number of ERMES-Mediated Mitochondria-ER Contact Sites (A) Schematic representation of the screen flow. 1. A strain harboring a GFP-tagged ERMES subunit (Mdm34) was crossed against yeast KO and DAmP collections, and a library was created so that each strain harbors a tagged ERMES on the background of a single mutation. 2. Live cells were imaged using an automated image acquisition system. 3. Images were manually inspected for an increase in the number of ERMES foci. (B) Fluorescence images of the increase in Mdm34-GFP foci in deletion of either vps39 or vam7 relative to a WT background. Scale bars, 5 mm. See also Figure S1.
B
2002), covering the entire yeast genome. Following the automated imaging of all strains generated (Breker et al., 2013), we manually examined the resulting images to uncover strains with the phenotype of choice (Figure 1A). The screen resulted in identification of over 100 different genetic backgrounds in which the Mdm34-GFP signal was altered (Table S1 available online). Strains that altered Mdm34-GFP morphology included deletion strains of ERMES components themselves that caused loss of the junction altogether, reduction in proteasomal subunits that caused an increase in protein levels, and loss of a large repertoire of mitochondrial proteins that caused a decrease in the intensity of foci. We focused on the four mutant backgrounds that displayed elevated number of foci per cell: Ddnm1, Dfis1 (Figure S1A), Dvam6/vps39 (hereinafter referred to as vps39), and Dvam7 (Figures 1B; Figures S1B and S1C). Dnm1 and Fis1 are essential components of the mitochondrial fission machinery (Bleazard et al., 1999; Mozdy et al., 96 Developmental Cell 30, 95–102, July 14, 2014 ª2014 Elsevier Inc.
2000). Interestingly, it has recently been shown that mitochondrial/ER contact sites mark the sites for fission to occur (Friedman et al., 2011; Murley et al., 2013). We therefore reasoned that the increase in ERMES foci on these backgrounds is most probably a result of an indirect effect on mitochondrial morphology or due to crosstalk with the ERMES complex. Vps39 and Vam7 have been previously characterized as components of the vacuolar fusion process (Price et al., 2000; Stroupe et al., 2006). These hits were surprising because no apparent direct link was reported between vacuolar morphology and ER-mitochondria connections or function. It has, however, previously been reported that the absence of Vps39 leads to impaired respiration capacity (Merz and Westermann, 2009). Therefore, we decided to examine the connection between the absence of Vps39 to the increase in ERMES foci. Vps39 Localizes to Contact Sites between Mitochondria and Vacuoles Termed vCLAMPs To see if Vps39 has any spatial connection to mitochondria, we tagged Vps39 with an N-terminal GFP (hereinafter referred to as GFP-Vps39), since tagging on the N terminus was previously shown to retain the function of the protein (Angers and Merz, 2009). Following GFP-Vps39 localization by fluorescence microscopy, we found that it was highly enriched in defined patches on the vacuolar membrane, which colocalized with a mitochondrial marker (a matrix-targeted blue fluorescent protein; MTS-BFP) (Figure 2A). We therefore attempted to uncover whether such an overlap between vacuoles and mitochondria implies an undescribed MCS. Electron microscopy demonstrated that indeed there exists a tight interface between the vacuolar membrane and the
Developmental Cell A Dynamic Mitochondria/Vacuole Interface
A
B
Figure 2. Vps39 Marks a Contact Site between Mitochondria and Vacuoles (A) GFP-tagged Vps39 is enriched in defined patches on the vacuolar membrane that colocalize with mitochondria. Scale bars, 5 mm. (B) Immunoelectron microscopy analysis of a GFP-Vps39 strain, using antibodies against GFP, demonstrates that Vps39 accumulates specifically at the mitochondria-vacuole MCS that we termed vCLAMP. Distribution analysis of gold particles (n = 400): 64% vCLAMP; 15% vacuole; 10% mitochondria; 7% cytosol; 4% nucleus. V, vacuole; M, mitochondria.
mitochondrial outer membrane. Similarly to what was observed by fluorescence microscopy, this MCS was labeled by GFPVps39. The membranes of the two organelles maintained a separate identity and were not fused, as would be expected from a bona fide MCS (Figure 2B). We therefore referred to this MCS as the vCLAMP (vacuole and mitochondria patch). This MCS was discovered and characterized in parallel from studies on Vps39 (see Ho¨nscher et al., 2014 [this issue of Developmental Cell]). Vps39 has been studied in depth for many years as part of the homotypic fusion and vacuole protein sorting (HOPS) tethering complex (Stroupe et al., 2006). In order to start and characterize its function at the vCLAMP, we decided to identify proteins interacting with GFP-Vps39 specifically at the contact site. To do that, we isolated cellular fractions enriched for mitochondria and performed pull-downs with an anti-GFP antibody. We reasoned that this should enrich for proteins that lie at the interface of the two organelles. Enriched proteins were identified by mass spectrometry analysis. (The complete list of significantly enriched proteins is given in Table S2.) The strongest interactions observed were among Vps39 and the additional five HOPS complex proteins (Figure S2A), as would be expected from the established role of Vps39. Interestingly, in addition to these known interactions, we found nine small molecule transporters that were enriched in our samples in a statistically significant manner (false discovery rate [FDR] = 0.05; Table S3). We tagged all nine transporters with a GFP tag in order to verify their
exact cellular localization. Five of the eight vacuolar proteins were not homogenously distributed over the vacuolar membrane but rather localized to specific patches (Figure S2B). Further colocalization with a mitochondrial marker showed that GFP-Mnr2 and GFP-Pho91 partially colocalized with mitochondria. (Figures S2C and S2D). This could suggest that the vCLAMP, marked by GFP-Vps39, could serve as a hub for interorganelle transport of small molecules. The Two Contact Sites of Mitochondria with the Endomembrane System Are in Dynamic Equilibrium and Serve as Mutual Backup Pathways Our original hypothesis for uncovering the vCLAMP was that optimal communication of mitochondria and the endomembrane system requires coregulation with ERMES. Indeed, our screen showed that loss of vCLAMP caused an increase in the number of ERMES mediated contact sites (Figure 1B and Figures S1A and S1B). Therefore, we wanted to explore the effect of deleting ERMES on the vCLAMP. Loss of ERMES (caused by deletion of mdm34 [Figures 3A and S3A] or any of the other three complex members [Figure 3C]) caused a dramatic expansion of the vCLAMP interface. In fact, in this genetic background, vCLAMP completely surrounded mitochondria, despite the fact that the cells retained normal morphology (Figure S3C). Electron microscopy demonstrated that the contact site expanded to the entire circumference of each mitochondrion (Figures 3B and S3B). This finding demonstrates that ERMES and vCLAMP are tightly coregulated, which would ensure that mitochondria does not detach from the endomembrane system and, as a consequence, no reduction in endomembrane/mitochondrial transport occurs in the absence of one MCS. The tight interplay between ERMES and vCLAMP would indeed ensure that, in the absence of one structure, the increase in the other would serve as a ‘‘backup’’ path for small-molecule transport. This predicts that disturbing both MCSs should, in fact, cause a dramatic cellular phenotype. To obtain double mutants, we crossed a Dvps39 strain with a strain harboring a deletion in each of the ERMES subunits. Diploid yeast were sporulated, and the haploid progeny from single meiosis events were dissected. We found that, in all cases, the combination of both mutations was lethal (Figure S4A), indicating that both MCSs cannot be simultaneously disturbed. Thus, as would be expected, mitochondria must be in contact with at least one organelle in the endomembrane system to maintain proper cellular functions. Loss of Both Mitochondrial Contact Sites Causes Defects in Phospholipid Transport Phospholipid transport between the endomembrane system and mitochondria is essential for the three-step enzymatic pathway generating aminoglycerophospholipids (see model in Figure 4H). In this pathway, phosphatidylserine (PtdSer) that is synthesized in the ER is transported to mitochondria to be converted to phosphatidylethanolamine (PtdEtn) by the inner mitochondrial membrane enzyme, PtdSer decarboxylase 1 (Psd1). Some of the PtdEtn is then transported back to the ER, where it can serve as a source for generation of phosphatidylcholine (PtdCho) (Birner and Daum, 2003; Daum et al., 1998). Although 5% of PtdEtn is created in the late endomembrane system by the Developmental Cell 30, 95–102, July 14, 2014 ª2014 Elsevier Inc. 97
Developmental Cell A Dynamic Mitochondria/Vacuole Interface
A
B
C
Figure 3. vCLAMP and ERMES Are in Dynamic Equilibrium (A) In a strain deleted for mdm34, mitochondria (visualized by MTS-RFP) become large and spherical, and the GFP-Vps39 marked vCLAMP expands to surround mitochondria. Scale bars, 5 mm.
98 Developmental Cell 30, 95–102, July 14, 2014 ª2014 Elsevier Inc.
paralogous enzyme Psd2 (Trotter and Voelker, 1995), the mitochondrial Psd1 is the major enzyme, and so a large lipid flux between the ER and mitochondria should exist. If indeed vCLAMP and ERMES are coregulated to sustain constant contact area between the endomembrane system and mitochondria, then this would explain the lack of phenotype on PtdEtn levels caused by losing ERMES alone (Kornmann et al., 2009; Nguyen et al., 2012; Voss et al., 2012). It would also predict that concomitant loss of both MCSs would result in a more dramatic decrease in the phospholipids that require such transport events. To measure this effect, we created a conditional strain that is deleted for vps39 and expresses MDM34 under the control of a repressible GalS promoter. We eliminated additional pathways for de novo PtdEtn synthesis by growing cells in synthetic media depleted for ethanolamine, so as to limit the Kennedy pathway (Daum et al., 1998), and by deleting the late endomembrane localized PtdSer decarboxylase, Psd2 (Trotter and Voelker, 1995). Under these conditions, the only existing pathway for de novo PtdEtn synthesis involves the mitochondrial Psd1 and the obligation of transporting ER synthesized PtdSer into mitochondria. The phospholipid composition of the triple mutant compared to control cells was examined after growth for 24 hr in glucose (to repress GalSp-MDM34 expression), at which point the cells are still alive (Figure S4B) but mitochondrial shape is already affected (Figure S4C), indicating a depletion in Mdm34 levels. Thin-layer chromatography (TLC) of phospholipids extracted from whole cells (Figure 4A, independent quadruplicate quantified in Figure 4B) or enriched mitochondrial samples (Figure 4C, independent triplicate quantified in Figure 4D) demonstrated that the mutant has up to 40% decrease in the amounts of PtdEtn and cardiolipin (CL) (CL observed only in the enriched mitochondrial samples) concomitantly with near doubling in phosphatidylinositol (PtdIns) levels. Specifically, the decrease in PtdEtn could only be seen in this condition, whereas in the single mutants (Dvps39 or GalSp-MDM34 in glucose; data not shown) or after short-term repression of GalSp-MDM34 in the triple mutant (Figures S4D and S4E), this reduction was not seen. This demonstrates that only the concomitant loss of both MCSs causes a halt in shuttling of PtdSer. More generally, PtdEtn, CL, and PtdIns are all synthesized from a common precursor, phosphatidic acid (PA); however, PtdEtn and CL are synthesized in mitochondria, whereas PtdIns is synthesized in the ER. Thus, accumulation of PtdIns and reduction of CL most probably also reflect a defect in PA transport from ER to mitochondria (Figure 4H). In order to monitor the direct conversion of PtdSer to PtdEtn and, consequently, to PtdCho, we metabolically labeled the cells using 3H-serine. The radioactive serine was incorporated in the ER to PtdSer, and, following phospholipid extraction and separation by TLC, we could follow its fate in control versus mutant cells. In the mutant cells, where vCLAMP is absent and ERMES is gradually depleted, there is a massive accumulation of PtdSer
(B) Electron microscopy analysis shows that, upon deletion of ERMES, the vCLAMP is enlarged to surround mitochondria (visualized by immunogold label against Por1). Cristae are marked by white arrows. (C) Deletion of each of the ERMES subunits yields a similar effect on vCLAMP dynamics. See also Figure S3.
Developmental Cell A Dynamic Mitochondria/Vacuole Interface
A
E
B
C
F
D
G
H
I
Figure 4. The vCLAMP Serves as a Route for Phospholipid Transport (A) TLC of phospholipids extracted from whole cells of control (WT) and a mutant disrupted for both ERMES and vCLAMP. PA, phosphatidic acid; PI, phosphatidylinositol; PS, phosphatidylserine; PC, phosphatidylcholine; PE, phosphatidylethanolamine. (B) Quantitation of four biologically independent repeats of whole cell extracts. Bars represent means ± SD. The disruption of both ERMES and vCLAMP causes a reduction in PE levels, although Psd1 is present in mitochondria. (C) TLC of phospholipids levels from mitochondrial (Mito) enriched fractions of control (WT) and a mutant disrupted for both ERMES and vCLAMP. Abbreviations for lipids are described in (A). (D) Quantitation of three biologically independent repeats of mitochondrial enriched fractions. Bars represent means ± SD. (E) TLC of phospholipids extracted from whole cells following a 2 hr pulse of 3H-serine to control (WT) and a mutant disrupted for both ERMES and vCLAMP. (F) Quantitation of four biologically independent repeats. Bars represent means + SD. Upon disruption of both vCLAMP and ERMES cells, phospholipid transport both into and out of mitochondria is impaired. As a result, high levels of de novo synthesized PtdSer accumulate in the ER, while very low levels of PtdCho are generated. (G) Autoradiograph of TLC of phospholipids extracted from whole cells of control (WT) and a mutant strain (Dvps39, GalSp-MDM34, Dpsd2) after 24 hr growth in glucose-containing media. Cells were labeled with 3H-serine for indicated time points, followed by 1 mM cold L-serine addition and phospholipid extraction. (H) Phospholipid biosynthesis pathways involve enzymes in both the ER and mitochondria, thus necessitating transport of substrates between both compartments. Etn, ethanolamine; Cho, choline; CDP-DAG, cytidine diphosphate diacylglycerol; DAG, diacylglycerol. (I) A schematic model representing the tight coregulation of ERMES and vCLAMP. Loss of one causes an enlargement of the other, and loss of both is lethal. See also Figure S4.
Developmental Cell 30, 95–102, July 14, 2014 ª2014 Elsevier Inc. 99
Developmental Cell A Dynamic Mitochondria/Vacuole Interface
(PtdSer content mutant/wild-type = 1.75), suggesting that as both contacts between mitochondria and the endomembrane system are gradually collapsing, more and more PtdSer, which cannot be transported to mitochondria, is accumulating in the ER (Figure 4E and independent quadruplicate quantified in Figure 4F; see also Figure 4G). Quantification of the various lipid species demonstrates a large difference in the amounts of de novo synthesized PtdCho. While PtdCho accumulates in control cells, it is hardly synthesized in the mutant. This difference probably reflects the inability of PtdEtn that was synthesized in the mitochondria to be transported back to the ER for further processing. Taken together, the results of steady-state and de novo phospholipid amounts demonstrate the role of both ERMES and vCLAMP in phospholipid transport. More importantly, they highlight how well coordinated these back-up pathways are, as only elimination of both yields a dramatic change in cellular phospholipid levels. DISCUSSION Functional cooperation between mitochondria and other organelles is essential. Since mitochondria is not connected to the endomembrane system through vesicular trafficking, the ERmitochondria MCS serves as an important interorganellar interaction, allowing for fast and direct transport of phospholipids and Ca+2. We show here that, in yeast, mitochondria maintain a second MCS, vCLAMP, linking it to the endomembrane system through the vacuole. Since vacuoles are linked to the ER—both directly, through the nuclear vacuolar junction (NVJ), and indirectly, through vesicular traffic—this MCS can serve as a bypass for flow of information and nutrients between the ER and mitochondria. Indeed, we show that ERMES and vCLAMP are coregulated and that vCLAMP serves as a backup for ERMES, as only in the case where both are missing do yeast suffer from dramatic alternations in phospholipid levels and death. Loss of both mitochondrial MCSs is lethal, and although a dramatic reduction in PtdEtn is observed when there is concomitant loss of ERMES and vCLAMP, it is possible that there exist additional essential functions for the two MCSs. In yeast, where the major ion stores (such as calcium, iron, and copper) reside in vacuoles, an additional essential function may be ion transport. Condition-specific direct traffic between endosomes and mitochondria in reticulocytes have been shown to facilitate iron transfer (Sheftel et al., 2007). Recently, it has been reported that mitochondria and melanosomes, pigment-synthesizing lysosome-like organelles, establish physical contacts. The proximities observed between both organelles correlated with melanosome biogenesis and maturation (Daniele et al., 2014). Hence, the need for sustaining a contact with later endomembrane organelles may be conserved from yeast to higher eukaryotes. More broadly, a better understanding of vCLAMP resident proteins must be obtained before the full extent of crosstalk between mitochondria and the endomembrane system can be determined. Previously described MCSs have been shown to be dynamic and respond to the needs of the cell. For example, the NVJ enlarges upon entry to stationary growth phase (Pan et al., 2000), the formation of new plasma membrane-ER MCSs is induced upon Ca2+ store depletion (Wu et al., 2006), and a late endo-
some/lysosome-ER MCS forms solely under conditions of increased cholesterol levels (Du et al., 2011). vCLAMP is dynamic too, in response to both loss of ERMES and changes in carbon source (see Ho¨nscher et al., 2014). It is this dynamic and condition-specific nature that can most probably account for the fact that the vCLAMP has remained elusive for so long. The dynamic nature of vCLAMP and its apparent regulation through metabolic cues (see Ho¨nscher et al., 2014) marks yet another link between intracellular organelle positioning, morphology, and degree of contact as determinants in the cell’s adaptation to changing metabolic requirements (Liesa and Shirihai, 2013) and cell fate (Csorda´s et al., 2006). To summarize, we demonstrate an MCS between vacuoles and mitochondria. This contact site, which we term vCLAMP, is in dynamic equilibrium with the ERMES mediated junction between mitochondria and the ER and works in parallel to enable the shuttling of small molecules (such as lipids) between the endomembrane system and mitochondria (model in Figure 4I). More generally, identification of the vCLAMP demonstrates the true complexity of interorganellar crosstalk: not only must an interface between two organelles be formed, but bypass systems must also be in place, and their size should be coregulated to serve the changing needs of the cell. Future efforts should be directed to elucidate the nature of the vCLAMP tether. Our proteomic analysis did not yield a definite answer as to how Vps39 is tethered to the mitochondria; whether through an adaptor protein or a yet-to-be-discovered mitochondrial partner. Future studies aimed at understanding the coregulation of the vCLAMP and ERMES should provide novel insights into what the cells sense to ensure optimal MCS surface area. Having visual markers and the genetic capacity to alter junction size should now give a platform for in depth investigations of this new cellular organization module. EXPERIMENTAL PROCEDURES Strains and Plasmids Strains created in this study are listed in a table in the Supplemental Experimental Procedures. All yeast strains in this study are based on the BY4741 laboratory strains (Brachmann et al., 1998). Genetic manipulations were performed using the Li-acetate, polyethylene glycol, single-stranded DNA method for transforming yeast strains (Gietz and Woods, 2006) using integration plasmids described elsewhere (Janke et al., 2004; Longtine et al., 1998). For mitochondria staining, we used either an MTS-red fluorescent protein (RFP) plasmid (kindly provided by Jodi Nunnari) or an MTS-BFP plasmid (kindly provided by Christian Ungermann and Benedikt Westermann). SGA and High-Content Screening The SGA technique was used to efficiently introduce an ERMES marker (GFPtagged Mdm34) into systematic yeast deletion and hypomorphic allele libraries (Breslow et al., 2008; Giaever et al., 2002). SGA was performed as described elsewhere (Cohen and Schuldiner, 2011; Tong and Boone, 2006; Tong et al., 2001). Microscopic screening was performed using an automated microscopy setup as described elsewhere (Cohen and Schuldiner, 2011). Further information is present in the Supplemental Experimental Procedures. Manual Fluorescence Microscopy Imaging was performed using an Olympus IX71 microscope controlled by the DeltaVision SoftWoRx 3.5.1 software with a 603 or 1003 oil lens. Images were captured by a Phoetometrics Coolsnap HQ camera with excitation at 490/ 20 nm and emission at 528/38 nm (for GFP); excitation at 555/28 nm and emission at 617/73 nm (for mCherry/RFP); or excitation at 402 nm and emission at
100 Developmental Cell 30, 95–102, July 14, 2014 ª2014 Elsevier Inc.
Developmental Cell A Dynamic Mitochondria/Vacuole Interface
457 nm (for BFP). Images were transferred to Adobe Photoshop CS3 for slight contrast and brightness adjustments. Electron Microscopy For immunoelectron microscopy, cells were fixed in 4% paraformaldehyde with 0.1% glutaraldehyde in 0.1 M cacodylate buffer (pH, 7.4) for 1 hr at room temperature and kept at 4 C during 1–2 days. The samples were soaked overnight in 2.3 M sucrose and rapidly frozen in liquid nitrogen. Frozen ultrathin (70–90 nm) sections were cut with a diamond knife at 120 C on a Leica EM UC6 ultramicrotome. The sections were collected on 200-mesh formvar-coated nickel grids. Sections were blocked by a blocking solution containing 1% BSA, 0.1% glycine, 0.1% gelatin, and 1% Tween20. Immunolabeling was performed using rabbit polyclonal anti-GFP antibody (ab6556, 1:100; Abcam) during 1.5–2 hr at room temperature followed by goat anti-rabbit immunoglobulin G coupled to 10 nm (or 15 nm) gold particles (1:20 dilution) for 30 min at room temperature. Contrasting and embedding were performed as described elsewhere (Tokuyasu, 1986). The embedded sections were scanned and digitally viewed on a Tecnai Spirit transmission electron microscope (FEI) at 120 kV using a CCD Eagle camera with TIA software (FEI). Phospholipid Analysis Phospholipids were extracted (Bligh and Dyer, 1959) from mitochondria-enriched fractions (Daum et al., 1982) or from whole-cell homogenates (Folch et al., 1957) and analyzed by TLC using chloroform:acetone:methanol:acetic acid:water (50:20:10:15:5, v:v:v:v:v) as the developing solvent. Lipids were visualized using copper sulfate (6.25 mM in 9.4 ml phosphoric acid) and heated at 110 C for 30–60 min, identified using lipid standards (Sigma, Avanti Polar Lipids), and quantified using ImageGauge Ver 4.0 software (FUJIFILM). For measuring de novo phospholipid synthesis, we metabolically labeled cells with 3H-serine as follows: cells were grown to early logarithmic phase and pelleted, and 15 optical density units were resuspended in 25 ml synthetic media (S-dextrose) supplemented with 0.5 mM ethanolamine and 1 mg/ml myriocin. 3H-serine (1 mCi/ml) was then added to a final concentration of 2 mCi/ml. Cells continued growth at 30 C until pulse was stopped by the addition of 1 mM cold L-serine at various time points (as indicated in the figures). Phospholipids were extracted (Folch et al., 1957) and separated by TLC using chloroform:methanol:acetic acid:water (25:15:4:2, v/v/v) as the developing solvent. [3H]-labeled lipids were visualized using a phosphorimaging screen (Fuji) and quantified using ImageGauge Ver 4.0 software (FUJIFILM). Interaction Proteomics Pull-down of GFP-Vps39 was performed from mitochondria-enriched preparations followed by liquid chromatography-tandem mass spectrometry analysis. Detailed information is present in the Supplemental Experimental Procedures. SUPPLEMENTAL INFORMATION
with the Israel Ministry of Science, and a recipient of Marie Curie International Reintegration Grant 239224 of the European Union. A.H.F. is the Joseph Meyerhoff Professor of Biochemistry at the Weizmann Institute of Science. T.G. is supported by the Israel Science Foundation. Received: October 15, 2013 Revised: April 14, 2014 Accepted: June 9, 2014 Published: July 14, 2014 REFERENCES Achleitner, G., Gaigg, B., Krasser, A., Kainersdorfer, E., Kohlwein, S.D., Perktold, A., Zellnig, G., and Daum, G. (1999). Association between the endoplasmic reticulum and mitochondria of yeast facilitates interorganelle transport of phospholipids through membrane contact. Eur. J. Biochem. 264, 545–553. Angers, C.G., and Merz, A.J. (2009). HOPS interacts with Apl5 at the vacuole membrane and is required for consumption of AP-3 transport vesicles. Mol. Biol. Cell 20, 4563–4574. Birner, R., and Daum, G. (2003). Biogenesis and cellular dynamics of aminoglycerophospholipids. Int. Rev. Cytol. 225, 273–323. Bleazard, W., McCaffery, J.M., King, E.J., Bale, S., Mozdy, A., Tieu, Q., Nunnari, J., and Shaw, J.M. (1999). The dynamin-related GTPase Dnm1 regulates mitochondrial fission in yeast. Nat. Cell Biol. 1, 298–304. Bligh, E.G., and Dyer, W.J. (1959). A rapid method of total lipid extraction and purification. Can. J. Biochem. Physiol. 37, 911–917. Brachmann, C.B., Davies, A., Cost, G.J., Caputo, E., Li, J., Hieter, P., and Boeke, J.D. (1998). Designer deletion strains derived from Saccharomyces cerevisiae S288C: a useful set of strains and plasmids for PCR-mediated gene disruption and other applications. Yeast 14, 115–132. Bratic, A., and Larsson, N.G. (2013). The role of mitochondria in aging. J. Clin. Invest. 123, 951–957. Breker, M., Gymrek, M., and Schuldiner, M. (2013). A novel single-cell screening platform reveals proteome plasticity during yeast stress responses. J. Cell Biol. 200, 839–850. Breslow, D.K., Cameron, D.M., Collins, S.R., Schuldiner, M., Stewart-Ornstein, J., Newman, H.W., Braun, S., Madhani, H.D., Krogan, N.J., and Weissman, J.S. (2008). A comprehensive strategy enabling high-resolution functional analysis of the yeast genome. Nat. Methods 5, 711–718. Cohen, Y., and Schuldiner, M. (2011). Advanced methods for high-throughput microscopy screening of genetically modified yeast libraries. Methods Mol. Biol. 781, 127–159. Costa, V., and Scorrano, L. (2012). Shaping the role of mitochondria in the pathogenesis of Huntington’s disease. EMBO J. 31, 1853–1864. Csorda´s, G., Renken, C., Va´rnai, P., Walter, L., Weaver, D., Buttle, K.F., Balla, T., Mannella, C.A., and Hajno´czky, G. (2006). Structural and functional features and significance of the physical linkage between ER and mitochondria. J. Cell Biol. 174, 915–921.
Supplemental Information includes Supplemental Experimental Procedures, four figures, and three tables and can be found with this article online at http://dx.doi.org/10.1016/j.devcel.2014.06.007.
Daniele, T., Hurbain, I., Vago, R., Casari, G., Raposo, G., Tacchetti, C., and Schiaffino, M.V. (2014). Mitochondria and melanosomes establish physical contacts modulated by Mfn2 and involved in organelle biogenesis. Curr. Biol. 24, 393–403.
ACKNOWLEDGMENTS
Daum, G., Bo¨hni, P.C., and Schatz, G. (1982). Import of proteins into mitochondria. Cytochrome b2 and cytochrome c peroxidase are located in the intermembrane space of yeast mitochondria. J. Biol. Chem. 257, 13028–13033.
We thank Jodi Nunnari for sharing the MTS-RFP plasmid and Christian Ungermann and Benedikt Westermann for the mBFP plasmid. We thank Oren Elbaz for immense help with the graphic design, Michal Breker and Silvia Chuartzman for technical assistance with the screening system, and Shai Fuchs for technical assistance with the flow cytometer. We thank Oren Schuldiner, Tslil Ast, and Shai Fuchs for critical reading of the manuscript. The electron microscopy studies were conducted at the Irving and Cherna Moskowitz Center for Nano and Bio-Nano Imaging at the Weizmann Institute of Science. This study was funded by a European Research Council Starting Grant (260395). Y.E.-A. was the recipient of the Weizmann Institute Dean’s postdoctoral fellowship. M.S. is an awardee of the EMBO Young Investigator Program, in conjunction
Daum, G., Lees, N.D., Bard, M., and Dickson, R. (1998). Biochemistry, cell biology and molecular biology of lipids of Saccharomyces cerevisiae. Yeast 14, 1471–1510. Du, X., Kumar, J., Ferguson, C., Schulz, T.A., Ong, Y.S., Hong, W., Prinz, W.A., Parton, R.G., Brown, A.J., and Yang, H. (2011). A role for oxysterol-binding protein-related protein 5 in endosomal cholesterol trafficking. J. Cell Biol. 192, 121–135. Elbaz, Y., and Schuldiner, M. (2011). Staying in touch: the molecular era of organelle contact sites. Trends Biochem. Sci. 36, 616–623.
Developmental Cell 30, 95–102, July 14, 2014 ª2014 Elsevier Inc. 101
Developmental Cell A Dynamic Mitochondria/Vacuole Interface
Folch, J., Lees, M., and Sloane Stanley, G.H. (1957). A simple method for the isolation and purification of total lipides from animal tissues. J. Biol. Chem. 226, 497–509.
Murley, A., Lackner, L.L., Osman, C., West, M., Voeltz, G.K., Walter, P., and Nunnari, J. (2013). ER-associated mitochondrial division links the distribution of mitochondria and mitochondrial DNA in yeast. eLife 2, e00422.
Friedman, J.R., Lackner, L.L., West, M., DiBenedetto, J.R., Nunnari, J., and Voeltz, G.K. (2011). ER tubules mark sites of mitochondrial division. Science 334, 358–362.
Nguyen, T.T., Lewandowska, A., Choi, J.Y., Markgraf, D.F., Junker, M., Bilgin, M., Ejsing, C.S., Voelker, D.R., Rapoport, T.A., and Shaw, J.M. (2012). Gem1 and ERMES do not directly affect phosphatidylserine transport from ER to mitochondria or mitochondrial inheritance. Traffic 13, 880–890.
Giaever, G., Chu, A.M., Ni, L., Connelly, C., Riles, L., Ve´ronneau, S., Dow, S., Lucau-Danila, A., Anderson, K., Andre´, B., et al. (2002). Functional profiling of the Saccharomyces cerevisiae genome. Nature 418, 387–391. Gietz, R.D., and Woods, R.A. (2006). Yeast transformation by the LiAc/SS Carrier DNA/PEG method. Methods Mol. Biol. 313, 107–120. Griffiths, E.J. (2012). Mitochondria and heart disease. Adv. Exp. Med. Biol. 942, 249–267. Ho¨nscher, C., Mari, M., Auffarth, K., Bohnert, M., Griffith, J., Geerts, W., van der Laan, M., Cabrera, M., Reggiori, F., and Ungermann, C. (2014). Cellular metabolism regulates contact sites between vacuoles and mitochondria. Dev. Cell 30, this issue, 86–94. Janke, C., Magiera, M.M., Rathfelder, N., Taxis, C., Reber, S., Maekawa, H., Moreno-Borchart, A., Doenges, G., Schwob, E., Schiebel, E., and Knop, M. (2004). A versatile toolbox for PCR-based tagging of yeast genes: new fluorescent proteins, more markers and promoter substitution cassettes. Yeast 21, 947–962. Kopec, K.O., Alva, V., and Lupas, A.N. (2010). Homology of SMP domains to the TULIP superfamily of lipid-binding proteins provides a structural basis for lipid exchange between ER and mitochondria. Bioinformatics 26, 1927– 1931. Kornmann, B., and Walter, P. (2010). ERMES-mediated ER-mitochondria contacts: molecular hubs for the regulation of mitochondrial biology. J. Cell Sci. 123, 1389–1393. Kornmann, B., Currie, E., Collins, S.R., Schuldiner, M., Nunnari, J., Weissman, J.S., and Walter, P. (2009). An ER-mitochondria tethering complex revealed by a synthetic biology screen. Science 325, 477–481.
Pan, X., Roberts, P., Chen, Y., Kvam, E., Shulga, N., Huang, K., Lemmon, S., and Goldfarb, D.S. (2000). Nucleus-vacuole junctions in Saccharomyces cerevisiae are formed through the direct interaction of Vac8p with Nvj1p. Mol. Biol. Cell 11, 2445–2457. Price, A., Wickner, W., and Ungermann, C. (2000). Proteins needed for vesicle budding from the Golgi complex are also required for the docking step of homotypic vacuole fusion. J. Cell Biol. 148, 1223–1229. Sheftel, A.D., Zhang, A.S., Brown, C., Shirihai, O., and Ponka, P. (2007). Direct interorganellar transfer of iron from endsome to mitochondrion. Blood 110, 125–132. Stroupe, C., Collins, K.M., Fratti, R.A., and Wickner, W. (2006). Purification of active HOPS complex reveals its affinities for phosphoinositides and the SNARE Vam7p. EMBO J. 25, 1579–1589. Tatsuta, T., Scharwey, M., and Langer, T. (2014). Mitochondrial lipid trafficking. Trends Cell Biol. 24, 44–52. Tokuyasu, K.T. (1986). Application of cryoultramicrotomy to immunocytochemistry. J. Microsc. 143, 139–149. Tong, A.H., and Boone, C. (2006). Synthetic genetic array analysis in Saccharomyces cerevisiae. Methods Mol. Biol. 313, 171–192. Tong, A.H., Evangelista, M., Parsons, A.B., Xu, H., Bader, G.D., Page´, N., Robinson, M., Raghibizadeh, S., Hogue, C.W., Bussey, H., et al. (2001). Systematic genetic analysis with ordered arrays of yeast deletion mutants. Science 294, 2364–2368.
Levine, T., and Loewen, C. (2006). Inter-organelle membrane contact sites: through a glass, darkly. Curr. Opin. Cell Biol. 18, 371–378.
Trotter, P.J., and Voelker, D.R. (1995). Identification of a non-mitochondrial phosphatidylserine decarboxylase activity (PSD2) in the yeast Saccharomyces cerevisiae. J. Biol. Chem. 270, 6062–6070.
Liesa, M., and Shirihai, O.S. (2013). Mitochondrial dynamics in the regulation of nutrient utilization and energy expenditure. Cell Metab. 17, 491–506.
Vance, J.E. (1990). Phospholipid synthesis in a membrane fraction associated with mitochondria. J. Biol. Chem. 265, 7248–7256.
Longtine, M.S., McKenzie, A., 3rd, Demarini, D.J., Shah, N.G., Wach, A., Brachat, A., Philippsen, P., and Pringle, J.R. (1998). Additional modules for versatile and economical PCR-based gene deletion and modification in Saccharomyces cerevisiae. Yeast 14, 953–961. Merz, S., and Westermann, B. (2009). Genome-wide deletion mutant analysis reveals genes required for respiratory growth, mitochondrial genome maintenance and mitochondrial protein synthesis in Saccharomyces cerevisiae. Genome Biol. 10, R95. Mozdy, A.D., McCaffery, J.M., and Shaw, J.M. (2000). Dnm1p GTPase-mediated mitochondrial fission is a multi-step process requiring the novel integral membrane component Fis1p. J. Cell Biol. 151, 367–380.
Vives-Bauza, C., and Przedborski, S. (2011). Mitophagy: the latest problem for Parkinson’s disease. Trends Mol. Med. 17, 158–165. Voss, C., Lahiri, S., Young, B.P., Loewen, C.J., and Prinz, W.A. (2012). ER-shaping proteins facilitate lipid exchange between the ER and mitochondria in S. cerevisiae. J. Cell Sci. 125, 4791–4799. Wu, M.M., Buchanan, J., Luik, R.M., and Lewis, R.S. (2006). Ca2+ store depletion causes STIM1 to accumulate in ER regions closely associated with the plasma membrane. J. Cell Biol. 174, 803–813. Yu, E., Mercer, J., and Bennett, M. (2012). Mitochondria in vascular disease. Cardiovasc. Res. 95, 173–182.
102 Developmental Cell 30, 95–102, July 14, 2014 ª2014 Elsevier Inc.
Short Article A Dynamic Interface between Vacuoles and Mitochondria in Yeast Yael Elbaz-Alon,1 Eden Rosenfeld-Gur,2 Vera Shinder,3 Anthony H. Futerman,2 Tamar Geiger,4 and Maya Schuldiner1,* 1Department
of Molecular Genetics, Weizmann Institute of Science, Rehovot 7610001, Israel of Biological Chemistry, Weizmann Institute of Science, Rehovot 7610001, Israel 3Electron Microscopy Unit, Weizmann Institute of Science, Rehovot 7610001, Israel 4Department of Human Molecular Genetics and Biochemistry, Sackler Faculty of Medicine, Tel Aviv University, Tel Aviv 6997801, Israel *Correspondence: [email protected] http://dx.doi.org/10.1016/j.devcel.2014.06.007 2Department
SUMMARY
Cellular life depends on continuous transport of lipids and small molecules between mitochondria and the endomembrane system. Recently, endoplasmic reticulum-mitochondrial encounter structure (ERMES) was identified as an important yet nonessential contact for such transport. Using a high-content screen in yeast, we found a contact site, marked by Vam6/Vps39, between vacuoles (the yeast lysosomal compartment) and mitochondria, named vCLAMP (vacuole and mitochondria patch). vCLAMP is enriched with ion and aminoacid transporters and has a role in lipid relay between the endomembrane system and mitochondria. Critically, we show that mitochondria are dependent on having one of two contact sites, ERMES or vCLAMP. The absence of one causes expansion of the other, and elimination of both is lethal. Identification of vCLAMP adds to our ability to understand the complexity of interorganellar crosstalk. INTRODUCTION Mitochondria generate the majority of cellular energy and house enzymes required for the synthesis, breakdown, and interconversion of various species of amino acids, lipids, iron/sulfur clusters, and other small molecules. Due to their diverse functions and essential roles in cellular metabolism, mitochondria serve as hubs for signaling in events such as growth, differentiation, or cell death. Loss of optimal mitochondrial activity is therefore, not surprisingly, implicated in a growing number of human diseases as well as in aging (Bratic and Larsson, 2013; Costa and Scorrano, 2012; Griffiths, 2012; Vives-Bauza and Przedborski, 2011; Yu et al., 2012). The central tasks of mitochondria in cells necessitate constant communication and transport of small molecules with other organelles. However, mitochondria are not connected to the endomembrane system via the vesicular pathway. Instead, mitochondria have been shown to communicate with the endomembrane system by virtue of a membrane contact site (MCS) where membranes of mitochondria come into close proximity to membranes of the endoplasmic reticulum (ER). This MCS, also
termed mitochondria-associated-membranes (Achleitner et al., 1999; Vance, 1990), enables ions and lipids to be rapidly transported in a nonvesicular manner (Elbaz and Schuldiner, 2011; Levine and Loewen, 2006; Tatsuta et al., 2014). Understanding the molecular machineries creating and regulating this MCS has been the arena of intense investigations in the past decade. In yeast, the molecular identity of the ER-mitochondria tethering complex was recently uncovered and is mediated by a four-protein complex termed the ER-mitochondria encounter structure (ERMES) (Kornmann et al., 2009). One of its hypothesized functions was to enable phospholipid transport (Kopec et al., 2010; Kornmann and Walter, 2010) required for building mitochondrial membranes as well as for the three-step biosynthetic pathway of aminoglycerophospholipids. Therefore, it was of great surprise when the loss of ERMES subunits had very little effect on cellular levels of aminoglycerophospholipids (Kornmann et al., 2009; Nguyen et al., 2012; Voss et al., 2012). Hence, it became clear that alternate routes of phospholipid transport must exist in the cell, and uncovering them should shed light on novel modes of communication between mitochondria and the endomembrane system. We report here our findings of an MCS between mitochondria and vacuoles (the yeast lysosomal compartment) and its functional significance in lipid transport between the endomembrane system and mitochondria. RESULTS A High-Content Screen Uncovers that Vam6/Vps39 Influences the Number of MCSs between the ER and Mitochondria We set out to uncover alternate paths for communication between mitochondria and the endomembrane system using a systematic genetic screening approach. We reasoned that the extent of ERMES mediated contact sites must be regulated so that loss of a parallel pathway would result in an increase in ERMES mediated contacts. Therefore, we screened for genetic backgrounds yielding a pronounced increase in the amount of ERMES foci per cell (Figure 1A). Using the synthetic genetic array (SGA) methodology (Cohen and Schuldiner, 2011; Tong and Boone, 2006; Tong et al., 2001), we created a library in which each strain harbors a green fluorescent protein (GFP)tagged ERMES subunit (Mdm34-GFP) on the background of a single gene mutation (Breslow et al., 2008; Giaever et al., Developmental Cell 30, 95–102, July 14, 2014 ª2014 Elsevier Inc. 95
Developmental Cell A Dynamic Mitochondria/Vacuole Interface
A
Figure 1. A Genome-wide Screen to Uncover Mutants Affecting the Number of ERMES-Mediated Mitochondria-ER Contact Sites (A) Schematic representation of the screen flow. 1. A strain harboring a GFP-tagged ERMES subunit (Mdm34) was crossed against yeast KO and DAmP collections, and a library was created so that each strain harbors a tagged ERMES on the background of a single mutation. 2. Live cells were imaged using an automated image acquisition system. 3. Images were manually inspected for an increase in the number of ERMES foci. (B) Fluorescence images of the increase in Mdm34-GFP foci in deletion of either vps39 or vam7 relative to a WT background. Scale bars, 5 mm. See also Figure S1.
B
2002), covering the entire yeast genome. Following the automated imaging of all strains generated (Breker et al., 2013), we manually examined the resulting images to uncover strains with the phenotype of choice (Figure 1A). The screen resulted in identification of over 100 different genetic backgrounds in which the Mdm34-GFP signal was altered (Table S1 available online). Strains that altered Mdm34-GFP morphology included deletion strains of ERMES components themselves that caused loss of the junction altogether, reduction in proteasomal subunits that caused an increase in protein levels, and loss of a large repertoire of mitochondrial proteins that caused a decrease in the intensity of foci. We focused on the four mutant backgrounds that displayed elevated number of foci per cell: Ddnm1, Dfis1 (Figure S1A), Dvam6/vps39 (hereinafter referred to as vps39), and Dvam7 (Figures 1B; Figures S1B and S1C). Dnm1 and Fis1 are essential components of the mitochondrial fission machinery (Bleazard et al., 1999; Mozdy et al., 96 Developmental Cell 30, 95–102, July 14, 2014 ª2014 Elsevier Inc.
2000). Interestingly, it has recently been shown that mitochondrial/ER contact sites mark the sites for fission to occur (Friedman et al., 2011; Murley et al., 2013). We therefore reasoned that the increase in ERMES foci on these backgrounds is most probably a result of an indirect effect on mitochondrial morphology or due to crosstalk with the ERMES complex. Vps39 and Vam7 have been previously characterized as components of the vacuolar fusion process (Price et al., 2000; Stroupe et al., 2006). These hits were surprising because no apparent direct link was reported between vacuolar morphology and ER-mitochondria connections or function. It has, however, previously been reported that the absence of Vps39 leads to impaired respiration capacity (Merz and Westermann, 2009). Therefore, we decided to examine the connection between the absence of Vps39 to the increase in ERMES foci. Vps39 Localizes to Contact Sites between Mitochondria and Vacuoles Termed vCLAMPs To see if Vps39 has any spatial connection to mitochondria, we tagged Vps39 with an N-terminal GFP (hereinafter referred to as GFP-Vps39), since tagging on the N terminus was previously shown to retain the function of the protein (Angers and Merz, 2009). Following GFP-Vps39 localization by fluorescence microscopy, we found that it was highly enriched in defined patches on the vacuolar membrane, which colocalized with a mitochondrial marker (a matrix-targeted blue fluorescent protein; MTS-BFP) (Figure 2A). We therefore attempted to uncover whether such an overlap between vacuoles and mitochondria implies an undescribed MCS. Electron microscopy demonstrated that indeed there exists a tight interface between the vacuolar membrane and the
Developmental Cell A Dynamic Mitochondria/Vacuole Interface
A
B
Figure 2. Vps39 Marks a Contact Site between Mitochondria and Vacuoles (A) GFP-tagged Vps39 is enriched in defined patches on the vacuolar membrane that colocalize with mitochondria. Scale bars, 5 mm. (B) Immunoelectron microscopy analysis of a GFP-Vps39 strain, using antibodies against GFP, demonstrates that Vps39 accumulates specifically at the mitochondria-vacuole MCS that we termed vCLAMP. Distribution analysis of gold particles (n = 400): 64% vCLAMP; 15% vacuole; 10% mitochondria; 7% cytosol; 4% nucleus. V, vacuole; M, mitochondria.
mitochondrial outer membrane. Similarly to what was observed by fluorescence microscopy, this MCS was labeled by GFPVps39. The membranes of the two organelles maintained a separate identity and were not fused, as would be expected from a bona fide MCS (Figure 2B). We therefore referred to this MCS as the vCLAMP (vacuole and mitochondria patch). This MCS was discovered and characterized in parallel from studies on Vps39 (see Ho¨nscher et al., 2014 [this issue of Developmental Cell]). Vps39 has been studied in depth for many years as part of the homotypic fusion and vacuole protein sorting (HOPS) tethering complex (Stroupe et al., 2006). In order to start and characterize its function at the vCLAMP, we decided to identify proteins interacting with GFP-Vps39 specifically at the contact site. To do that, we isolated cellular fractions enriched for mitochondria and performed pull-downs with an anti-GFP antibody. We reasoned that this should enrich for proteins that lie at the interface of the two organelles. Enriched proteins were identified by mass spectrometry analysis. (The complete list of significantly enriched proteins is given in Table S2.) The strongest interactions observed were among Vps39 and the additional five HOPS complex proteins (Figure S2A), as would be expected from the established role of Vps39. Interestingly, in addition to these known interactions, we found nine small molecule transporters that were enriched in our samples in a statistically significant manner (false discovery rate [FDR] = 0.05; Table S3). We tagged all nine transporters with a GFP tag in order to verify their
exact cellular localization. Five of the eight vacuolar proteins were not homogenously distributed over the vacuolar membrane but rather localized to specific patches (Figure S2B). Further colocalization with a mitochondrial marker showed that GFP-Mnr2 and GFP-Pho91 partially colocalized with mitochondria. (Figures S2C and S2D). This could suggest that the vCLAMP, marked by GFP-Vps39, could serve as a hub for interorganelle transport of small molecules. The Two Contact Sites of Mitochondria with the Endomembrane System Are in Dynamic Equilibrium and Serve as Mutual Backup Pathways Our original hypothesis for uncovering the vCLAMP was that optimal communication of mitochondria and the endomembrane system requires coregulation with ERMES. Indeed, our screen showed that loss of vCLAMP caused an increase in the number of ERMES mediated contact sites (Figure 1B and Figures S1A and S1B). Therefore, we wanted to explore the effect of deleting ERMES on the vCLAMP. Loss of ERMES (caused by deletion of mdm34 [Figures 3A and S3A] or any of the other three complex members [Figure 3C]) caused a dramatic expansion of the vCLAMP interface. In fact, in this genetic background, vCLAMP completely surrounded mitochondria, despite the fact that the cells retained normal morphology (Figure S3C). Electron microscopy demonstrated that the contact site expanded to the entire circumference of each mitochondrion (Figures 3B and S3B). This finding demonstrates that ERMES and vCLAMP are tightly coregulated, which would ensure that mitochondria does not detach from the endomembrane system and, as a consequence, no reduction in endomembrane/mitochondrial transport occurs in the absence of one MCS. The tight interplay between ERMES and vCLAMP would indeed ensure that, in the absence of one structure, the increase in the other would serve as a ‘‘backup’’ path for small-molecule transport. This predicts that disturbing both MCSs should, in fact, cause a dramatic cellular phenotype. To obtain double mutants, we crossed a Dvps39 strain with a strain harboring a deletion in each of the ERMES subunits. Diploid yeast were sporulated, and the haploid progeny from single meiosis events were dissected. We found that, in all cases, the combination of both mutations was lethal (Figure S4A), indicating that both MCSs cannot be simultaneously disturbed. Thus, as would be expected, mitochondria must be in contact with at least one organelle in the endomembrane system to maintain proper cellular functions. Loss of Both Mitochondrial Contact Sites Causes Defects in Phospholipid Transport Phospholipid transport between the endomembrane system and mitochondria is essential for the three-step enzymatic pathway generating aminoglycerophospholipids (see model in Figure 4H). In this pathway, phosphatidylserine (PtdSer) that is synthesized in the ER is transported to mitochondria to be converted to phosphatidylethanolamine (PtdEtn) by the inner mitochondrial membrane enzyme, PtdSer decarboxylase 1 (Psd1). Some of the PtdEtn is then transported back to the ER, where it can serve as a source for generation of phosphatidylcholine (PtdCho) (Birner and Daum, 2003; Daum et al., 1998). Although 5% of PtdEtn is created in the late endomembrane system by the Developmental Cell 30, 95–102, July 14, 2014 ª2014 Elsevier Inc. 97
Developmental Cell A Dynamic Mitochondria/Vacuole Interface
A
B
C
Figure 3. vCLAMP and ERMES Are in Dynamic Equilibrium (A) In a strain deleted for mdm34, mitochondria (visualized by MTS-RFP) become large and spherical, and the GFP-Vps39 marked vCLAMP expands to surround mitochondria. Scale bars, 5 mm.
98 Developmental Cell 30, 95–102, July 14, 2014 ª2014 Elsevier Inc.
paralogous enzyme Psd2 (Trotter and Voelker, 1995), the mitochondrial Psd1 is the major enzyme, and so a large lipid flux between the ER and mitochondria should exist. If indeed vCLAMP and ERMES are coregulated to sustain constant contact area between the endomembrane system and mitochondria, then this would explain the lack of phenotype on PtdEtn levels caused by losing ERMES alone (Kornmann et al., 2009; Nguyen et al., 2012; Voss et al., 2012). It would also predict that concomitant loss of both MCSs would result in a more dramatic decrease in the phospholipids that require such transport events. To measure this effect, we created a conditional strain that is deleted for vps39 and expresses MDM34 under the control of a repressible GalS promoter. We eliminated additional pathways for de novo PtdEtn synthesis by growing cells in synthetic media depleted for ethanolamine, so as to limit the Kennedy pathway (Daum et al., 1998), and by deleting the late endomembrane localized PtdSer decarboxylase, Psd2 (Trotter and Voelker, 1995). Under these conditions, the only existing pathway for de novo PtdEtn synthesis involves the mitochondrial Psd1 and the obligation of transporting ER synthesized PtdSer into mitochondria. The phospholipid composition of the triple mutant compared to control cells was examined after growth for 24 hr in glucose (to repress GalSp-MDM34 expression), at which point the cells are still alive (Figure S4B) but mitochondrial shape is already affected (Figure S4C), indicating a depletion in Mdm34 levels. Thin-layer chromatography (TLC) of phospholipids extracted from whole cells (Figure 4A, independent quadruplicate quantified in Figure 4B) or enriched mitochondrial samples (Figure 4C, independent triplicate quantified in Figure 4D) demonstrated that the mutant has up to 40% decrease in the amounts of PtdEtn and cardiolipin (CL) (CL observed only in the enriched mitochondrial samples) concomitantly with near doubling in phosphatidylinositol (PtdIns) levels. Specifically, the decrease in PtdEtn could only be seen in this condition, whereas in the single mutants (Dvps39 or GalSp-MDM34 in glucose; data not shown) or after short-term repression of GalSp-MDM34 in the triple mutant (Figures S4D and S4E), this reduction was not seen. This demonstrates that only the concomitant loss of both MCSs causes a halt in shuttling of PtdSer. More generally, PtdEtn, CL, and PtdIns are all synthesized from a common precursor, phosphatidic acid (PA); however, PtdEtn and CL are synthesized in mitochondria, whereas PtdIns is synthesized in the ER. Thus, accumulation of PtdIns and reduction of CL most probably also reflect a defect in PA transport from ER to mitochondria (Figure 4H). In order to monitor the direct conversion of PtdSer to PtdEtn and, consequently, to PtdCho, we metabolically labeled the cells using 3H-serine. The radioactive serine was incorporated in the ER to PtdSer, and, following phospholipid extraction and separation by TLC, we could follow its fate in control versus mutant cells. In the mutant cells, where vCLAMP is absent and ERMES is gradually depleted, there is a massive accumulation of PtdSer
(B) Electron microscopy analysis shows that, upon deletion of ERMES, the vCLAMP is enlarged to surround mitochondria (visualized by immunogold label against Por1). Cristae are marked by white arrows. (C) Deletion of each of the ERMES subunits yields a similar effect on vCLAMP dynamics. See also Figure S3.
Developmental Cell A Dynamic Mitochondria/Vacuole Interface
A
E
B
C
F
D
G
H
I
Figure 4. The vCLAMP Serves as a Route for Phospholipid Transport (A) TLC of phospholipids extracted from whole cells of control (WT) and a mutant disrupted for both ERMES and vCLAMP. PA, phosphatidic acid; PI, phosphatidylinositol; PS, phosphatidylserine; PC, phosphatidylcholine; PE, phosphatidylethanolamine. (B) Quantitation of four biologically independent repeats of whole cell extracts. Bars represent means ± SD. The disruption of both ERMES and vCLAMP causes a reduction in PE levels, although Psd1 is present in mitochondria. (C) TLC of phospholipids levels from mitochondrial (Mito) enriched fractions of control (WT) and a mutant disrupted for both ERMES and vCLAMP. Abbreviations for lipids are described in (A). (D) Quantitation of three biologically independent repeats of mitochondrial enriched fractions. Bars represent means ± SD. (E) TLC of phospholipids extracted from whole cells following a 2 hr pulse of 3H-serine to control (WT) and a mutant disrupted for both ERMES and vCLAMP. (F) Quantitation of four biologically independent repeats. Bars represent means + SD. Upon disruption of both vCLAMP and ERMES cells, phospholipid transport both into and out of mitochondria is impaired. As a result, high levels of de novo synthesized PtdSer accumulate in the ER, while very low levels of PtdCho are generated. (G) Autoradiograph of TLC of phospholipids extracted from whole cells of control (WT) and a mutant strain (Dvps39, GalSp-MDM34, Dpsd2) after 24 hr growth in glucose-containing media. Cells were labeled with 3H-serine for indicated time points, followed by 1 mM cold L-serine addition and phospholipid extraction. (H) Phospholipid biosynthesis pathways involve enzymes in both the ER and mitochondria, thus necessitating transport of substrates between both compartments. Etn, ethanolamine; Cho, choline; CDP-DAG, cytidine diphosphate diacylglycerol; DAG, diacylglycerol. (I) A schematic model representing the tight coregulation of ERMES and vCLAMP. Loss of one causes an enlargement of the other, and loss of both is lethal. See also Figure S4.
Developmental Cell 30, 95–102, July 14, 2014 ª2014 Elsevier Inc. 99
Developmental Cell A Dynamic Mitochondria/Vacuole Interface
(PtdSer content mutant/wild-type = 1.75), suggesting that as both contacts between mitochondria and the endomembrane system are gradually collapsing, more and more PtdSer, which cannot be transported to mitochondria, is accumulating in the ER (Figure 4E and independent quadruplicate quantified in Figure 4F; see also Figure 4G). Quantification of the various lipid species demonstrates a large difference in the amounts of de novo synthesized PtdCho. While PtdCho accumulates in control cells, it is hardly synthesized in the mutant. This difference probably reflects the inability of PtdEtn that was synthesized in the mitochondria to be transported back to the ER for further processing. Taken together, the results of steady-state and de novo phospholipid amounts demonstrate the role of both ERMES and vCLAMP in phospholipid transport. More importantly, they highlight how well coordinated these back-up pathways are, as only elimination of both yields a dramatic change in cellular phospholipid levels. DISCUSSION Functional cooperation between mitochondria and other organelles is essential. Since mitochondria is not connected to the endomembrane system through vesicular trafficking, the ERmitochondria MCS serves as an important interorganellar interaction, allowing for fast and direct transport of phospholipids and Ca+2. We show here that, in yeast, mitochondria maintain a second MCS, vCLAMP, linking it to the endomembrane system through the vacuole. Since vacuoles are linked to the ER—both directly, through the nuclear vacuolar junction (NVJ), and indirectly, through vesicular traffic—this MCS can serve as a bypass for flow of information and nutrients between the ER and mitochondria. Indeed, we show that ERMES and vCLAMP are coregulated and that vCLAMP serves as a backup for ERMES, as only in the case where both are missing do yeast suffer from dramatic alternations in phospholipid levels and death. Loss of both mitochondrial MCSs is lethal, and although a dramatic reduction in PtdEtn is observed when there is concomitant loss of ERMES and vCLAMP, it is possible that there exist additional essential functions for the two MCSs. In yeast, where the major ion stores (such as calcium, iron, and copper) reside in vacuoles, an additional essential function may be ion transport. Condition-specific direct traffic between endosomes and mitochondria in reticulocytes have been shown to facilitate iron transfer (Sheftel et al., 2007). Recently, it has been reported that mitochondria and melanosomes, pigment-synthesizing lysosome-like organelles, establish physical contacts. The proximities observed between both organelles correlated with melanosome biogenesis and maturation (Daniele et al., 2014). Hence, the need for sustaining a contact with later endomembrane organelles may be conserved from yeast to higher eukaryotes. More broadly, a better understanding of vCLAMP resident proteins must be obtained before the full extent of crosstalk between mitochondria and the endomembrane system can be determined. Previously described MCSs have been shown to be dynamic and respond to the needs of the cell. For example, the NVJ enlarges upon entry to stationary growth phase (Pan et al., 2000), the formation of new plasma membrane-ER MCSs is induced upon Ca2+ store depletion (Wu et al., 2006), and a late endo-
some/lysosome-ER MCS forms solely under conditions of increased cholesterol levels (Du et al., 2011). vCLAMP is dynamic too, in response to both loss of ERMES and changes in carbon source (see Ho¨nscher et al., 2014). It is this dynamic and condition-specific nature that can most probably account for the fact that the vCLAMP has remained elusive for so long. The dynamic nature of vCLAMP and its apparent regulation through metabolic cues (see Ho¨nscher et al., 2014) marks yet another link between intracellular organelle positioning, morphology, and degree of contact as determinants in the cell’s adaptation to changing metabolic requirements (Liesa and Shirihai, 2013) and cell fate (Csorda´s et al., 2006). To summarize, we demonstrate an MCS between vacuoles and mitochondria. This contact site, which we term vCLAMP, is in dynamic equilibrium with the ERMES mediated junction between mitochondria and the ER and works in parallel to enable the shuttling of small molecules (such as lipids) between the endomembrane system and mitochondria (model in Figure 4I). More generally, identification of the vCLAMP demonstrates the true complexity of interorganellar crosstalk: not only must an interface between two organelles be formed, but bypass systems must also be in place, and their size should be coregulated to serve the changing needs of the cell. Future efforts should be directed to elucidate the nature of the vCLAMP tether. Our proteomic analysis did not yield a definite answer as to how Vps39 is tethered to the mitochondria; whether through an adaptor protein or a yet-to-be-discovered mitochondrial partner. Future studies aimed at understanding the coregulation of the vCLAMP and ERMES should provide novel insights into what the cells sense to ensure optimal MCS surface area. Having visual markers and the genetic capacity to alter junction size should now give a platform for in depth investigations of this new cellular organization module. EXPERIMENTAL PROCEDURES Strains and Plasmids Strains created in this study are listed in a table in the Supplemental Experimental Procedures. All yeast strains in this study are based on the BY4741 laboratory strains (Brachmann et al., 1998). Genetic manipulations were performed using the Li-acetate, polyethylene glycol, single-stranded DNA method for transforming yeast strains (Gietz and Woods, 2006) using integration plasmids described elsewhere (Janke et al., 2004; Longtine et al., 1998). For mitochondria staining, we used either an MTS-red fluorescent protein (RFP) plasmid (kindly provided by Jodi Nunnari) or an MTS-BFP plasmid (kindly provided by Christian Ungermann and Benedikt Westermann). SGA and High-Content Screening The SGA technique was used to efficiently introduce an ERMES marker (GFPtagged Mdm34) into systematic yeast deletion and hypomorphic allele libraries (Breslow et al., 2008; Giaever et al., 2002). SGA was performed as described elsewhere (Cohen and Schuldiner, 2011; Tong and Boone, 2006; Tong et al., 2001). Microscopic screening was performed using an automated microscopy setup as described elsewhere (Cohen and Schuldiner, 2011). Further information is present in the Supplemental Experimental Procedures. Manual Fluorescence Microscopy Imaging was performed using an Olympus IX71 microscope controlled by the DeltaVision SoftWoRx 3.5.1 software with a 603 or 1003 oil lens. Images were captured by a Phoetometrics Coolsnap HQ camera with excitation at 490/ 20 nm and emission at 528/38 nm (for GFP); excitation at 555/28 nm and emission at 617/73 nm (for mCherry/RFP); or excitation at 402 nm and emission at
100 Developmental Cell 30, 95–102, July 14, 2014 ª2014 Elsevier Inc.
Developmental Cell A Dynamic Mitochondria/Vacuole Interface
457 nm (for BFP). Images were transferred to Adobe Photoshop CS3 for slight contrast and brightness adjustments. Electron Microscopy For immunoelectron microscopy, cells were fixed in 4% paraformaldehyde with 0.1% glutaraldehyde in 0.1 M cacodylate buffer (pH, 7.4) for 1 hr at room temperature and kept at 4 C during 1–2 days. The samples were soaked overnight in 2.3 M sucrose and rapidly frozen in liquid nitrogen. Frozen ultrathin (70–90 nm) sections were cut with a diamond knife at 120 C on a Leica EM UC6 ultramicrotome. The sections were collected on 200-mesh formvar-coated nickel grids. Sections were blocked by a blocking solution containing 1% BSA, 0.1% glycine, 0.1% gelatin, and 1% Tween20. Immunolabeling was performed using rabbit polyclonal anti-GFP antibody (ab6556, 1:100; Abcam) during 1.5–2 hr at room temperature followed by goat anti-rabbit immunoglobulin G coupled to 10 nm (or 15 nm) gold particles (1:20 dilution) for 30 min at room temperature. Contrasting and embedding were performed as described elsewhere (Tokuyasu, 1986). The embedded sections were scanned and digitally viewed on a Tecnai Spirit transmission electron microscope (FEI) at 120 kV using a CCD Eagle camera with TIA software (FEI). Phospholipid Analysis Phospholipids were extracted (Bligh and Dyer, 1959) from mitochondria-enriched fractions (Daum et al., 1982) or from whole-cell homogenates (Folch et al., 1957) and analyzed by TLC using chloroform:acetone:methanol:acetic acid:water (50:20:10:15:5, v:v:v:v:v) as the developing solvent. Lipids were visualized using copper sulfate (6.25 mM in 9.4 ml phosphoric acid) and heated at 110 C for 30–60 min, identified using lipid standards (Sigma, Avanti Polar Lipids), and quantified using ImageGauge Ver 4.0 software (FUJIFILM). For measuring de novo phospholipid synthesis, we metabolically labeled cells with 3H-serine as follows: cells were grown to early logarithmic phase and pelleted, and 15 optical density units were resuspended in 25 ml synthetic media (S-dextrose) supplemented with 0.5 mM ethanolamine and 1 mg/ml myriocin. 3H-serine (1 mCi/ml) was then added to a final concentration of 2 mCi/ml. Cells continued growth at 30 C until pulse was stopped by the addition of 1 mM cold L-serine at various time points (as indicated in the figures). Phospholipids were extracted (Folch et al., 1957) and separated by TLC using chloroform:methanol:acetic acid:water (25:15:4:2, v/v/v) as the developing solvent. [3H]-labeled lipids were visualized using a phosphorimaging screen (Fuji) and quantified using ImageGauge Ver 4.0 software (FUJIFILM). Interaction Proteomics Pull-down of GFP-Vps39 was performed from mitochondria-enriched preparations followed by liquid chromatography-tandem mass spectrometry analysis. Detailed information is present in the Supplemental Experimental Procedures. SUPPLEMENTAL INFORMATION
with the Israel Ministry of Science, and a recipient of Marie Curie International Reintegration Grant 239224 of the European Union. A.H.F. is the Joseph Meyerhoff Professor of Biochemistry at the Weizmann Institute of Science. T.G. is supported by the Israel Science Foundation. Received: October 15, 2013 Revised: April 14, 2014 Accepted: June 9, 2014 Published: July 14, 2014 REFERENCES Achleitner, G., Gaigg, B., Krasser, A., Kainersdorfer, E., Kohlwein, S.D., Perktold, A., Zellnig, G., and Daum, G. (1999). Association between the endoplasmic reticulum and mitochondria of yeast facilitates interorganelle transport of phospholipids through membrane contact. Eur. J. Biochem. 264, 545–553. Angers, C.G., and Merz, A.J. (2009). HOPS interacts with Apl5 at the vacuole membrane and is required for consumption of AP-3 transport vesicles. Mol. Biol. Cell 20, 4563–4574. Birner, R., and Daum, G. (2003). Biogenesis and cellular dynamics of aminoglycerophospholipids. Int. Rev. Cytol. 225, 273–323. Bleazard, W., McCaffery, J.M., King, E.J., Bale, S., Mozdy, A., Tieu, Q., Nunnari, J., and Shaw, J.M. (1999). The dynamin-related GTPase Dnm1 regulates mitochondrial fission in yeast. Nat. Cell Biol. 1, 298–304. Bligh, E.G., and Dyer, W.J. (1959). A rapid method of total lipid extraction and purification. Can. J. Biochem. Physiol. 37, 911–917. Brachmann, C.B., Davies, A., Cost, G.J., Caputo, E., Li, J., Hieter, P., and Boeke, J.D. (1998). Designer deletion strains derived from Saccharomyces cerevisiae S288C: a useful set of strains and plasmids for PCR-mediated gene disruption and other applications. Yeast 14, 115–132. Bratic, A., and Larsson, N.G. (2013). The role of mitochondria in aging. J. Clin. Invest. 123, 951–957. Breker, M., Gymrek, M., and Schuldiner, M. (2013). A novel single-cell screening platform reveals proteome plasticity during yeast stress responses. J. Cell Biol. 200, 839–850. Breslow, D.K., Cameron, D.M., Collins, S.R., Schuldiner, M., Stewart-Ornstein, J., Newman, H.W., Braun, S., Madhani, H.D., Krogan, N.J., and Weissman, J.S. (2008). A comprehensive strategy enabling high-resolution functional analysis of the yeast genome. Nat. Methods 5, 711–718. Cohen, Y., and Schuldiner, M. (2011). Advanced methods for high-throughput microscopy screening of genetically modified yeast libraries. Methods Mol. Biol. 781, 127–159. Costa, V., and Scorrano, L. (2012). Shaping the role of mitochondria in the pathogenesis of Huntington’s disease. EMBO J. 31, 1853–1864. Csorda´s, G., Renken, C., Va´rnai, P., Walter, L., Weaver, D., Buttle, K.F., Balla, T., Mannella, C.A., and Hajno´czky, G. (2006). Structural and functional features and significance of the physical linkage between ER and mitochondria. J. Cell Biol. 174, 915–921.
Supplemental Information includes Supplemental Experimental Procedures, four figures, and three tables and can be found with this article online at http://dx.doi.org/10.1016/j.devcel.2014.06.007.
Daniele, T., Hurbain, I., Vago, R., Casari, G., Raposo, G., Tacchetti, C., and Schiaffino, M.V. (2014). Mitochondria and melanosomes establish physical contacts modulated by Mfn2 and involved in organelle biogenesis. Curr. Biol. 24, 393–403.
ACKNOWLEDGMENTS
Daum, G., Bo¨hni, P.C., and Schatz, G. (1982). Import of proteins into mitochondria. Cytochrome b2 and cytochrome c peroxidase are located in the intermembrane space of yeast mitochondria. J. Biol. Chem. 257, 13028–13033.
We thank Jodi Nunnari for sharing the MTS-RFP plasmid and Christian Ungermann and Benedikt Westermann for the mBFP plasmid. We thank Oren Elbaz for immense help with the graphic design, Michal Breker and Silvia Chuartzman for technical assistance with the screening system, and Shai Fuchs for technical assistance with the flow cytometer. We thank Oren Schuldiner, Tslil Ast, and Shai Fuchs for critical reading of the manuscript. The electron microscopy studies were conducted at the Irving and Cherna Moskowitz Center for Nano and Bio-Nano Imaging at the Weizmann Institute of Science. This study was funded by a European Research Council Starting Grant (260395). Y.E.-A. was the recipient of the Weizmann Institute Dean’s postdoctoral fellowship. M.S. is an awardee of the EMBO Young Investigator Program, in conjunction
Daum, G., Lees, N.D., Bard, M., and Dickson, R. (1998). Biochemistry, cell biology and molecular biology of lipids of Saccharomyces cerevisiae. Yeast 14, 1471–1510. Du, X., Kumar, J., Ferguson, C., Schulz, T.A., Ong, Y.S., Hong, W., Prinz, W.A., Parton, R.G., Brown, A.J., and Yang, H. (2011). A role for oxysterol-binding protein-related protein 5 in endosomal cholesterol trafficking. J. Cell Biol. 192, 121–135. Elbaz, Y., and Schuldiner, M. (2011). Staying in touch: the molecular era of organelle contact sites. Trends Biochem. Sci. 36, 616–623.
Developmental Cell 30, 95–102, July 14, 2014 ª2014 Elsevier Inc. 101
Developmental Cell A Dynamic Mitochondria/Vacuole Interface
Folch, J., Lees, M., and Sloane Stanley, G.H. (1957). A simple method for the isolation and purification of total lipides from animal tissues. J. Biol. Chem. 226, 497–509.
Murley, A., Lackner, L.L., Osman, C., West, M., Voeltz, G.K., Walter, P., and Nunnari, J. (2013). ER-associated mitochondrial division links the distribution of mitochondria and mitochondrial DNA in yeast. eLife 2, e00422.
Friedman, J.R., Lackner, L.L., West, M., DiBenedetto, J.R., Nunnari, J., and Voeltz, G.K. (2011). ER tubules mark sites of mitochondrial division. Science 334, 358–362.
Nguyen, T.T., Lewandowska, A., Choi, J.Y., Markgraf, D.F., Junker, M., Bilgin, M., Ejsing, C.S., Voelker, D.R., Rapoport, T.A., and Shaw, J.M. (2012). Gem1 and ERMES do not directly affect phosphatidylserine transport from ER to mitochondria or mitochondrial inheritance. Traffic 13, 880–890.
Giaever, G., Chu, A.M., Ni, L., Connelly, C., Riles, L., Ve´ronneau, S., Dow, S., Lucau-Danila, A., Anderson, K., Andre´, B., et al. (2002). Functional profiling of the Saccharomyces cerevisiae genome. Nature 418, 387–391. Gietz, R.D., and Woods, R.A. (2006). Yeast transformation by the LiAc/SS Carrier DNA/PEG method. Methods Mol. Biol. 313, 107–120. Griffiths, E.J. (2012). Mitochondria and heart disease. Adv. Exp. Med. Biol. 942, 249–267. Ho¨nscher, C., Mari, M., Auffarth, K., Bohnert, M., Griffith, J., Geerts, W., van der Laan, M., Cabrera, M., Reggiori, F., and Ungermann, C. (2014). Cellular metabolism regulates contact sites between vacuoles and mitochondria. Dev. Cell 30, this issue, 86–94. Janke, C., Magiera, M.M., Rathfelder, N., Taxis, C., Reber, S., Maekawa, H., Moreno-Borchart, A., Doenges, G., Schwob, E., Schiebel, E., and Knop, M. (2004). A versatile toolbox for PCR-based tagging of yeast genes: new fluorescent proteins, more markers and promoter substitution cassettes. Yeast 21, 947–962. Kopec, K.O., Alva, V., and Lupas, A.N. (2010). Homology of SMP domains to the TULIP superfamily of lipid-binding proteins provides a structural basis for lipid exchange between ER and mitochondria. Bioinformatics 26, 1927– 1931. Kornmann, B., and Walter, P. (2010). ERMES-mediated ER-mitochondria contacts: molecular hubs for the regulation of mitochondrial biology. J. Cell Sci. 123, 1389–1393. Kornmann, B., Currie, E., Collins, S.R., Schuldiner, M., Nunnari, J., Weissman, J.S., and Walter, P. (2009). An ER-mitochondria tethering complex revealed by a synthetic biology screen. Science 325, 477–481.
Pan, X., Roberts, P., Chen, Y., Kvam, E., Shulga, N., Huang, K., Lemmon, S., and Goldfarb, D.S. (2000). Nucleus-vacuole junctions in Saccharomyces cerevisiae are formed through the direct interaction of Vac8p with Nvj1p. Mol. Biol. Cell 11, 2445–2457. Price, A., Wickner, W., and Ungermann, C. (2000). Proteins needed for vesicle budding from the Golgi complex are also required for the docking step of homotypic vacuole fusion. J. Cell Biol. 148, 1223–1229. Sheftel, A.D., Zhang, A.S., Brown, C., Shirihai, O., and Ponka, P. (2007). Direct interorganellar transfer of iron from endsome to mitochondrion. Blood 110, 125–132. Stroupe, C., Collins, K.M., Fratti, R.A., and Wickner, W. (2006). Purification of active HOPS complex reveals its affinities for phosphoinositides and the SNARE Vam7p. EMBO J. 25, 1579–1589. Tatsuta, T., Scharwey, M., and Langer, T. (2014). Mitochondrial lipid trafficking. Trends Cell Biol. 24, 44–52. Tokuyasu, K.T. (1986). Application of cryoultramicrotomy to immunocytochemistry. J. Microsc. 143, 139–149. Tong, A.H., and Boone, C. (2006). Synthetic genetic array analysis in Saccharomyces cerevisiae. Methods Mol. Biol. 313, 171–192. Tong, A.H., Evangelista, M., Parsons, A.B., Xu, H., Bader, G.D., Page´, N., Robinson, M., Raghibizadeh, S., Hogue, C.W., Bussey, H., et al. (2001). Systematic genetic analysis with ordered arrays of yeast deletion mutants. Science 294, 2364–2368.
Levine, T., and Loewen, C. (2006). Inter-organelle membrane contact sites: through a glass, darkly. Curr. Opin. Cell Biol. 18, 371–378.
Trotter, P.J., and Voelker, D.R. (1995). Identification of a non-mitochondrial phosphatidylserine decarboxylase activity (PSD2) in the yeast Saccharomyces cerevisiae. J. Biol. Chem. 270, 6062–6070.
Liesa, M., and Shirihai, O.S. (2013). Mitochondrial dynamics in the regulation of nutrient utilization and energy expenditure. Cell Metab. 17, 491–506.
Vance, J.E. (1990). Phospholipid synthesis in a membrane fraction associated with mitochondria. J. Biol. Chem. 265, 7248–7256.
Longtine, M.S., McKenzie, A., 3rd, Demarini, D.J., Shah, N.G., Wach, A., Brachat, A., Philippsen, P., and Pringle, J.R. (1998). Additional modules for versatile and economical PCR-based gene deletion and modification in Saccharomyces cerevisiae. Yeast 14, 953–961. Merz, S., and Westermann, B. (2009). Genome-wide deletion mutant analysis reveals genes required for respiratory growth, mitochondrial genome maintenance and mitochondrial protein synthesis in Saccharomyces cerevisiae. Genome Biol. 10, R95. Mozdy, A.D., McCaffery, J.M., and Shaw, J.M. (2000). Dnm1p GTPase-mediated mitochondrial fission is a multi-step process requiring the novel integral membrane component Fis1p. J. Cell Biol. 151, 367–380.
Vives-Bauza, C., and Przedborski, S. (2011). Mitophagy: the latest problem for Parkinson’s disease. Trends Mol. Med. 17, 158–165. Voss, C., Lahiri, S., Young, B.P., Loewen, C.J., and Prinz, W.A. (2012). ER-shaping proteins facilitate lipid exchange between the ER and mitochondria in S. cerevisiae. J. Cell Sci. 125, 4791–4799. Wu, M.M., Buchanan, J., Luik, R.M., and Lewis, R.S. (2006). Ca2+ store depletion causes STIM1 to accumulate in ER regions closely associated with the plasma membrane. J. Cell Biol. 174, 803–813. Yu, E., Mercer, J., and Bennett, M. (2012). Mitochondria in vascular disease. Cardiovasc. Res. 95, 173–182.
102 Developmental Cell 30, 95–102, July 14, 2014 ª2014 Elsevier Inc.
