SUMMARY
Mitochondria are dynamic organelles that constantly fuse and divide to maintain their proper morphology, which is essential for their normal functions. Energy production, a central role of mitochondria, demands highly folded structures of the mitochondrial inner membrane (MIM) called cristae and a dimeric phospholipid (PL) cardiolipin (CL). Previous studies identified a number of factors involved in mitochondrial dynamics, crista formation, and CL biosynthesis, yet it is still enigmatic how these events are interconnected and cooperated. Here, we first report that mitochondrial fusion/division dynamics is important to maintain CL abundance. Second, our genetic and biochemical analyses revealed that intra-mitochondrial PL transport plays an important role in crista formation. Finally, we show that simultaneous defects in MIM fusion and intra-mitochondrial PL transport cause a drastic decrease in crista structure, resulting in decreased CL levels. These results expand our understanding of the integrated functional network among the PL transport, crista formation, and CL biogenesis.
INTRODUCTION
Mitochondria, power plants in eukaryotic cells, produce most cellular ATP molecules by oxidative phosphorylation, as well as reactive oxygen species (ROS) as bi-products. To prevent accumulation of ROS-mediated oxidative damage of mitochondrial contents including mitochondrial DNA, mitochondria constantly exchange their contents through their fusion and division processes. For efficient ATP production, the mitochondrial inner membrane (MIM) is folded inwards to form crista structures for maximizing its surface area. Cardiolipin (CL), a mitochondria-specific phospholipid (PL), also plays critical roles in ATP production through stabilizing the respiratory chain supercomplexes (Böttinger et al., 2012; Claypool et al., 2008; Pfeiffer et al., 2003; Zhang et al., 2005). In addition, CL is also important for other mitochondrial functions including mitochondrial fusion/division dynamics (DeVay et al., 2009; Joshi et al., 2012; Kameoka et al., 2018), mitochondrial protein import (Gebert et al., 2009; Kutik et al., 2008; Malhotra et al., 2017; Sauerwald et al., 2015; Tamura et al., 2009) and regulation of mitophagy (Chu et al., 2013; Shen et al., 2017). In Saccharomyces cerevisiae, CL is synthesized via a series of modifications of phosphatidic acid (PA), the precursor PL generated in the ER. To synthesize CL, PA is first transported to the MIM, where CL synthetic enzymes are located. On the matrix side of the MIM, Tam41, mitochondrial CDP-diacylglycerol (DAG) synthase converts PA to CDP-DAG by using a nucleotide CTP (Tamura et al., 2013). Then, Pgs1, PGP synthase substitutes glycerol for the CDP moiety of CDP-DAG, producing phosphatidylglycerol phosphate (PGP) (Chang et al., 1998a). Gep4, PGP phosphatase dephosphorylates PGP to form phosphatidylglycerol (PG) (Osman et al., 2010). Crd1, CL synthase couples PG and CDP-DAG to generate CL (Tuller et al., 1998; Chang et al., 1998b). Finally, CL is matured through remodeling of its fatty acid chains by Cld1 CL-specific phospholipase and Taz1 acyltransferase (Beranek et al., 2009; Gu et al., 2004).
In addition to these CL synthetic enzymes, PL transfer proteins play essential roles in CL biosynthesis. A highly conserved protein complex, Ups1-Mdm35 is a lipid transfer machinery present in the mitochondrial intermembrane space (IMS) (Osman et al., 2009; Potting et al., 2010; Sesaki et al., 2006; Tamura et al., 2009, 2010). Biochemical and structural studies revealed that the Ups1-Mdm35 complex specifically recognizes the head group of PA and transfers PA from the MOM to MIM (Connerth et al., 2012; Watanabe et al., 2015; Yu et al., 2015). The Ups1-Mdm35 complex thus contributes to CL biosynthesis by delivering PA to the MIM, where the CL synthetic enzymes are located. Ups2, a Ups1 homologue, also forms a complex with Mdm35 and mediates phosphatidylserine (PS) transfer from the MOM to MIM, thereby contributing to phosphatidylethanolamine (PE) synthesis in mitochondria since PS synthesized outside mitochondria is converted to PE by Psd1 PE synthase in the MIM (Aaltonen et al., 2016; Miyata et al., 2016). In fact, loss of Ups1 or Ups2 leads to a decrease in the CL or PE level, respectively. CL and PE, both of which tend to form a nonbilayer membrane structure, are functionally redundant in part and the simultaneous loss of enzymes involved in the biosynthesis of CL and PE is synthetic lethal (Gohil et al., 2005). Therefore, one can naturally expect that simultaneous loss of Ups1 and Ups2 would cause a severe growth defect. However strangely, this is not the case; ups1Δups2Δ cells grow almost normally and contain a comparable amount of CL to that of wild-type cells (Tamura et al., 2009). These findings indicate that defective CL synthesis due to the absence of Ups1 is restored by the additional loss of Ups2. One explanation for this observation could be that the decreased PE level in mitochondria rather than loss of Ups2 would activate other pathway(s) to replenish CL in the absence of Ups1 (Miyata et al., 2017). Puzzlingly, CL synthetic enzymes such as Tam41 and Pgs1 are dispensable for CL synthesis when Ups1 is absent (Connerth et al., 2012). These findings collectively suggest that there are still unknown pathways and/or factors important for CL biosynthesis.
We reasoned that the unknown factors that can compensate the defects in PL transport for the CL and PE synthesis would become critical for cell growth in the absence of Mdm35, which is the functional binding partner for both Ups1 and Ups2. In this work, we focused on Mgm1, a dynamin-related GTPase that mediates the fusion of the MIM (Sesaki et al., 2003; Wong et al., 2000) and the formation of lamellar cristae, because it is reported to genetically interact with the MDM35 gene (Hoppins et al., 2011; Harner et al., 2016). Interestingly, simultaneous loss of Mdm35 and Mgm1, but not another mitochondrial protein for the MOM fusion Fzo1(Hermann et al., 1998), led to severe defects in growth and crista formation, suggesting that Mdm35 plays an overlapping function with Mgm1 in crista formation rather than mitochondrial fusion (Sesaki et al., 2003; Wong et al., 2000). Indeed, we confirmed that loss of Mdm35 alone causes a decrease in the crista structure. In addition, we show that the CL level is significantly decreased when crista formation is strongly impaired. These findings indicate that mitochondrial inner membrane fusion and intra-mitochondrial PL transport collectively contribute to the formation of normal MIM structures, which are prerequisite for the CL maintenance. These findings point to the importance of the previously overlooked relationship among the mitochondrial dynamics, PL transport, crista structure, and CL biogenesis.
Results and Discussion
Mgm1 is critical for cell growths when Mdm35 is absent
To gain insight into the “alternative” CL synthetic pathway that is activated upon inactivation of the Ups proteins, we investigated the genes that genetically interact with MDM35 using the genetic interactome information available at the Saccharomyces genome database (http://www.yeastgenome.org/). Among a number of the candidate genes, we focused on DNM1 and MGM1, which are involved in mitochondrial division and fusion, respectively, since these genetic interactions could point to uncharacterized roles, if any, of mitochondrial dynamics in the cellular phospholipid (PL) homeostasis.
We first tried to confirm if Dnm1 is indeed important for the growth of mdm35Δ cells as reported previously (Hoppins et al., 2011). To this end, we combined the mdm35Δ mutation with the dnm1Δ mutation and compared their cell growth with that of wild-type or the single-deletion mutant strains. As controls, we also created the dnm1Δ mutant strains with the ups1Δ or ups2Δ mutation. The ups1Δ strain showed defective cell growth whereas both the ups2Δ and mdm35Δ strains grew normally at 30°C and 37°C on a fermentable medium, YPD (Fig. 1A) (Tamura et al., 2009, 2010), although mdm35Δ cells grew more slowly than that of wild-type at low temperature 23°C (Potting et al., 2010). The dnm1Δ mutation hardly affected the growth of the single-deletion ups1Δ, ups2Δ and mdm35Δ strains on the fermentable medium, YPD, whereas it caused synthetic growth defects with the ups2Δ or mdm35Δ mutation on a nonfermentable medium, YPLac at 37°C. This suggests that mitochondrial division becomes important when Ups2 or Mdm35 is absent under non-fermentable conditions and at elevated temperature (Fig. 1B).
Figure 1. Mdm35 is critical for the growth of the mgm1Δdnm1Δ but not fzo1Δdnm1Δ strain.
Serial dilutions of the indicated yeast cells were spotted onto (A) (C) YPD or (B) (D) YPLac and cultivated at 23, 30, 37°C (YPD) or 23, 30, 35°C (YPLac) for the indicated time periods.
Next, we attempted to confirm if MGM1 genetically interacts with MDM35. The phenotypes caused by impaired mitochondrial fusion, such as fragmented mitochondria, growth defects and loss of mitochondrial DNA (mtDNA), can be restored by additional loss of a mitochondrial division factor like Dnm1 (Lee et al., 2012; Sesaki and Jensen, 2001; Sesaki et al., 2003). We thus deleted the MGM1 gene together with the DNM1 gene to eliminate possible secondary effects associated with such strong phenotypes seen in cells defective in the mitochondrial fusion. As a control, we tested the effects of lack of Fzo1, which is another mitochondrial fusion protein localized in the MOM (Hermann et al., 1998), although genetic interactions have not been reported for MDM35 with FZO1. The triple-deletion mgm1Δdnm1Δmdm35Δ cells showed severe growth defects under any conditions we tested, including both fermentable and non-fermentable media, even severer than those of mgm1Δdnm1Δups1Δ cells. In contrast, the fzo1Δ mutation only mildly affected the growths of cells lacking Mdm35 (Fig. 1C, D). Therefore, Mgm1, not Fzo1, is crucial for the restoration of the growth defects arising from the block of the Ups1/2 mediated PA and PS transports across the IMS by the MDM35 deletion.
Cardiolipin is significantly decreased when both Mgm1 and Mdm35 are lacking
The strong negative-genetic interactions between the MDM35 and MGM1 genes suggest that Mgm1 is required for the restoration of the CL level in mdm35Δ cells. To test this notion, we analyzed the effects of the ups1Δ, ups2Δ or mdm35Δ mutation on PL compositions of wild-type, dnm1Δ, fzo1Δdnm1Δ , and mgm1Δdnm1Δ cells. Total PLs were prepared from those yeast cells cultivated in YPD containing 32Pi and analyzed by thin-layer chromatography (TLC). As we reported previously, the ups1Δ strain showed decreased CL but normal PE levels whereas ups2Δ and mdm35Δ strains contained nearly normal CL but decreased PE levels (Fig. 2). The levels of other major PLs such as phosphatidylserine, phosphatidylinositol, and phosphatidylcholine were not drastically altered among the mutant cells although the levels of a minor PL, phosphatidic acid decreased when mitochondrial fusion/division dynamics was defective. The dnm1Δ mutation slightly decreased the CL levels for the strains with the ups1Δ or ups2Δ background, not for wild-type strain, and these decreases appeared additive in the dnm1Δmdm35Δ strain. Nevertheless, the CL level was significantly restored in dnm1Δmdm35Δ strain as compared with the dnm1Δups1Δ strain. The dnm1Δ mutation did not affect the levels of PE for the strains with or without ups1Δ, ups2Δ or mdm35Δ background at all. These data suggest that Dnm1 has a minor role in maintaining the normal CL abundance. Interestingly, we found that CL levels decreased to ~50% in the fzo1Δdnm1Δ and mgm1Δdnm1Δ strains, but not in the dnm1Δ strain. We further noticed that the CL and PE levels were significantly lower in fzo1Δ and mgm1Δ cells like those of ρ0 cells, suggesting that the presence of mtDNA is critical to maintain normal levels of CL and PE (Fig. S1). This may indicate an uncharacterized role of mitochondrial fusion/division dynamics in the maintenance of the normal CL levels.
Figure 2. CL level is significantly decreased in the mgm1Δdnm1Δmdm35Δ strain.
Indicated yeast cells were grown in YPD media containing 2 μCi/ml 32Pi. Total phospholipids were extracted from whole cells and analyzed by TLC (left panels). Phospholipids were migrated in solvent A (chloroform/ethanol/water/triethylamine, 30/35/7/35, v/v) (top left) and solvent B (chloroform/methanol/25% ammonia, 65/35/5, v/v) (top right). The amounts of CL, PE, PS, PC, PA, and PI were determined (right panels). The ratio of each phospholipid to total phospholipids in wild-type was set to 100%. Values are mean ± SEM (n = 3).
The decreased CL levels could be a cause for the impaired cell growth of fzo1Δdnm1Δups2Δ and fzo1Δdnm1Δmdm35Δ under non-fermentable conditions (Fig. 1D) because concomitant decreases in levels of functionally redundant CL and PE are known to cause synthetic growth defects (Gohil et al., 2005). In addition, we noticed that the loss of Ups1 further decreases the CL levels in the fzo1Δdnm1Δ and mgm1Δdnm1Δ strains to ~10% as compared with that of the wild-type strain (Fig. 2). These findings indicate that impaired mitochondrial fusion/division events could lead to the decrease in the CL levels irrespective of the normal PA transport by Ups1. It should be emphasized that the mdm35Δ mutation did not restore the CL level at all in the strains with the mgm1Δdnm1Δ background whereas it did for the fzo1Δdnm1Δ background.
In summary, despite the overlapping function of Fzo1 and Mgm1 in mitochondrial fusion, our genetic and biochemical analyses showed that Mgm1 but not Fzo1 is critical for the cell growth and CL abundance of the strain lacking Mdm35. These data point out the crucial role of Mgm1 in the maintenance of CL abundance when Mdm35 is absent and suggest that Mgm1, but not Fzo1, has an overlapping function with Mdm35 in the CL biogenesis.
Mgm1 and Mdm35 play an overlapping role in crista formation
Electron tomographic analyses revealed the presence of two distinct types of crista structures, lamellar and tubular cristae, and lamellar crista was suggested to be generated by the fusion of two distinct MIMs by Mgm1 after fusion of MOMs (Harner et al., 2016). In contrast, tubular crista formation may instead require the import of proteins and PLs into the MIM, although this idea has not been experimentally tested yet. The MICOS complex at the crista junctions (CJs) appears to be involved in the formation of both the lamellar and tubular cristae (Harner et al., 2011; Hoppins et al., 2011; von der Malsburg et al., 2011; Rabl et al., 2009; Tarasenko et al., 2017). Taking this knowledge into consideration, we reasoned that simultaneous loss of Mgm1, involved in lamellar crista formation, and Mdm35, involved in tubular cristae formation through PL transport, may lead to impairment of the total crista formation and thereby result in severe growth defects and decreased CL levels. To test this idea, we analyzed the ultrastructure of the MIM in the presence or absence of Mdm35 and/or mitochondrial dynamics proteins Fzo1, Dnm1, and Mgm1 by electron microscopy (Fig. 3A). Consistent with previous studies, the electron micrographs showed that clear crista structures are present in fzo1Δdnm1Δ and in mgm1Δdnm1Δ cells (Fig. 3B) (Harner et al., 2016; Sesaki et al., 2003). We also confirmed the previous observation that septa structures were observed more frequently in the mitochondria without Fzo1 than in those without Mgm1 (Fig. 3A, B) (Harner et al., 2016). Interestingly, loss of Mdm35 led to a decrease in crista structures, suggesting the potential role of intramitochondrial PL transports in crista formation. Strikingly, the MIM structure was drastically altered when both MDM35 and MGM1 were deleted in the dnm1Δ background, and even empty mitochondria without internal MIM structures were frequently detected (Fig. 3A, B). Such severe alterations in the MIM structure were not seen in fzo1Δdnm1Δmdm35Δ cells, suggesting that Mdm35 and Mgm1 but not Fzo1 play an overlapping function in crista formation. In addition, these results clearly pointed out that Mgm1 can generate lamellar crista structure even without MOM-fusion events. Although how Mgm1 contributes to lamellar crista formation without MOM fusion is unclear at this point, the MIM may divide in a Dnm1-independent manner (Messerschmitt et al., 2003) and lamellar crista may be generated by Mgm1-dependent fusion of the two distinct MIMs. Identification of such factors that mediate the MIM division would be of important for understanding the mechanism of crista formation.
Figure 3. The crista formation is drastically impaired in mgm1Δdnm1Δmdm35Δ but not in fzo1Δdnm1mdm35Δ.
(A) Mitochondria in the indicated yeast cells were observed by TEM. Scale bar: 500 nm (B) About 100 mitochondria were observed for each strain and the ratios of mitochondria with the indicated internal membranes are shown. (C) A summary of relationships among Mgm1, MICOS and Mdm35 functions in crista formation. Δ means lack of MICOS component such as Mic10 or Mic60. “CJ ×” means that CJ formation is defective. 〇* means that the growth and CL level are almost normal.
(D) A model of two parallel mechanisms for crista formation.
Based on these observations, we conclude that the three key factors i.e. Mgm1, MICOS and Mdm35 (or intramitochondrial PL transport) are functionally interconnected to form normal crista structures (Fig. 3C and D). As summarized in Fig. 3C, defective intramitochondrial PL transport due to the Mdm35 loss negatively affects crista formation even though both Mgm1 and MICOS are functional (Fig. 3B and C, row 2). On the other hand, either the mgm1Δdnm1Δ or mic60Δ mutation does not affect crista formation to that extent although CJs are largely missing in the absence of Mic60 (Körner et al. 2009; Harner et al., 2011; Hoppins et al., 2011; von der Malsburg et al., 2011, Fig. 3 C lines 3 and 4). Therefore, functions of Mgm1 and MICOS are not sufficient to generate normal crista structures in the absence of Mdm35, or in other words, intramitochondrial PL transport contributes to Mgm1- and/or MICOS-dependent crista formations.
Interestingly, intramitochondrial PL transport seems to act on the MICOS-dependent crista formation rather than Mgm1-dependent one because simultaneous deletions of MICOS (MIC10 or MIC60) and MDM35 did not lead to strong synthetic defects in growths or CL abundance (Fig. S2A and B), which is in contrast to the case when the mgm1Δdnm1Δ and mdm35Δ mutations are combined (Figs. 1 and 2). In addition, it should be noted that either MICOS or Mdm35 becomes crucial for crista formation when Mgm1 is absent (mgm1ΔdnmΔ background) (Fig. 3C, rows 5 and 6). These results clearly indicate that Mgm1 can produce crista structures to some extent even in the absence of both MICOS and Mdm35, yet intramitochondrial PL transport or MICOS alone is not able to produce sufficient crista structures, especially lamellar cristae, in the mgm1Δdnm1Δ background. Taken together, we propose that the two distinct pathways that generate crista structures (Fig. 3D); one is the Mgm1-dependent pathway for lamellar crista formation, in which only CJ formation requires the MICOS function (Korner et al. 2009; Harner et al., 2011; Hoppins et al., 2011; von der Malsburg et al., 2011), and the other is the MICOS-dependent pathway for tubular crista formation, in which intramitochondrial PL transport is critical. When either Mgm1- or MICOS-dependent pathway is active, crista structures can be formed. However, simultaneous defects in both the pathways lead to depletion of crista structures, which appears to result in a drastic decrease in the CL levels and severe growth defects (Fig. 3D). The basis that the defective crista formation leads to decreased CL levels will be discussed below.
Compromised mitochondrial fusion/division partly affects the levels of CL synthetic enzymes
PL analyses revealed that the CL level is decreased to about 50% in fzo1Δdnm1Δ and mgm1Δdnm1Δ cells (Fig. 2). Besides, loss of Mdm35 drastically decreased the CL level in mgm1Δdnm1Δ cells whereas it only mildly affected the CL abundance in fzo1Δdnm1Δ cells (Fig. 2). A simple explanation for the decreased CL level is that compromised mitochondrial dynamics, as well as abnormal crista morphology, had a negative impact on the stabilities of CL synthetic enzymes. We thus examined the steady-state levels of CL synthetic enzymes in these deletion mutant cells. To detect endogenous levels of Gep4 and Crd1, we chromosomally expressed C-terminally 3xHemagglutinin (HA)-tagged Gep4 and N-terminally 3xFLAG-tagged Crd1, respectively (Longtine et al., 1998). Immunoblotting of membrane fractions showed that FLAG-Crd1 levels decreased in fzo1Δdnm1Δ, mgm1Δdnm1Δ, fzo1Δdnm1Δmdm35Δ and mgm1Δdnm1Δmdm35Δ cells as compared with those in wild-type cells (Fig. 4A). In addition, the levels of Cho2 and Psd1 also decreased in these mutant cells (Fig. 4A), but on the other hand, Tam41 and Gep4-HA levels were comparable or even increased in fzo1Δdnm1Δ, mgm1Δdnm1Δ, and mgm1Δdnm1Δmdm35Δ membranes, as compared with wild-type membranes (Fig. 4A).
Figure 4. Abnormal crista formation leads to decreased CL levels.
(A) Membrane fractions were extracted from the indicated yeast cells expressing Gep4-HA and FLAG-Crd1 and analyzed by immunoblotting using the indicated antibodies. (B, C) The indicated yeast cells were observed by TEM. Scale bar, 500 nm (D) Total phospholipids prepared from the indicated yeast cells were analyzed by TLC in solvent A (left) or solvent B (right). The CL levels were determined. The ratio of CL to total phospholipids in wild-type cells was set to 100%. Values are mean ± SEM (n = 3). (E) Serial dilutions of the indicated yeast cells were spotted onto YPD or YPLac plates cultivated at 23, 30, 37°C (YPD) or 23, 30, 35°C (YPLac) for the indicated time periods. (F) Membrane fractions were extracted from the indicated yeast cells expressing Gep4-HA and FLAG-Crd1 and analyzed by immunoblotting using the indicated antibodies.
The decreased levels of FLAG-Crd1 could explain the decreased CL levels in fzo1Δdnm1Δ and mgm1Δdnm1Δ cells (Fig. 2). Although fzo1Δdnm1Δ and mgm1Δdnm1Δ cells are capable of maintaining mtDNA as compared with the single-deletion mutant (fzo1Δ and mgm1Δ) cells, they (fzo1Δdnm1Δ and mgm1dnm1Δ cells) still tend to show the ρ− phenotype more frequently than wild-type cells (Osman et al., 2015). Therefore, compromised respiratory activities arising from the possible accumulation of nonfunctional mtDNA could decrease the membrane potential (ΔΨ) across the MIM, which would result in defects in the ΔΨ-requiring MIM and matrix protein import. This is consistent with the previous observation as well as our results that ρ° cells without mtDNA possessed decreased amounts of CL (Chen et al., 2010, Fig. S1). We indeed observed that mitochondria isolated from fzo1Δdnm1Δ and mgm1Δdnm1Δ cells showed the decreased ΔΨ as compared with that of wild-type mitochondria, especially under the fermentable condition (Fig. S3A). In addition, our immunoblotting using mitochondria isolated from cells grown in the nonfermentable (YPLac) medium showed that steady-state levels of FLAG-Crd1 are restored in fzo1Δdnm1Δ and mgm1Δdnm1Δ mitochondria as compared with those under the fermentable condition (Fig. S3B vs Fig. 4A and F). Consistently, our in vitro protein import assay showed that mitochondrial proteins including Crd1 and Psd1 were almost normally imported into fzo1Δdnm1Δ and mgm1Δdnm1Δ mitochondria isolated under the nonfermentable condition (Fig. S3C). These results suggest that mitochondrial protein import is impaired in fzo1Δdnm1Δ and mgm1Δdnm1Δ cells due to the decreased ΔΨ, resulting in the decreased Crd1 levels under the fermentable condition. To further confirm this idea, we assessed the mitochondrial protein import under the fermentable condition in vivo. To this end, we forcibly induced the expression of FLAG-Crd1, which is controlled by the GAL1 promoter, by shifting the growth medium from YPD to YPGal and analyzed the protein levels by immunoblotting. Interestingly, the levels of FLAG-Crd1 were significantly lower in fzo1Δdnm1Δ and mgm1Δdnm1Δ cells as compared with that in wild-type cells after inducing FLAG-Crd1 expressions (Fig. S3D). We observed similar delayed accumulations of other mitochondrial proteins such as Om45, Cyt c1, and Kgd1, upon expression in the galactose-containing medium. These observations indeed confirmed our idea that mitochondrial protein import is generally defective in fzo1Δdnm1Δ and mgm1Δdnm1Δ cells under the fermentable condition. We performed cycloheximide chase experiments using wild-type, fzo1Δdnm1Δ and mgm1Δdnm1Δ cells overexpressing FLAG-Crd1 from a multi-copy plasmid and found that FLAG-Crd1 and Psd1 were stably present over time in all yeast strains we tested whereas a short-lived protein Fzo1 was rapidly degraded (Escobar-Henriques et al., 2006, Fig. S3E). Therefore, we propose that the general import defect of mitochondrial proteins was the cause of the decreased levels of several mitochondrial proteins including Crd1 in fzo1Δdnm1Δ and mgm1Δdnm1Δ cells under the fermentable condition. Although it is still not clear why the level of an ER protein Cho2 decreased and Tam41 and Gep4 levels increased in fzo1Δdnm1Δ and mgm1Δdnm1Δ cells, transcriptional regulation may contribute to the maintenance of these protein levels.
Similar to Crd1, Psd1, which is anchored to the MIM with its N-terminal transmembrane segment (Horvath et al., 2012), also showed a reduced protein level in fzo1Δdnm1Δ and mgm1Δdnm1Δ cells, yet without a drastic change in the PE levels (Fig. 2). The only mildly affected level of PE is probably due to the presence of multiple PE synthetic pathways mediated by the endosome-resident PE synthase Psd2, the Kennedy pathway using soluble substrate CDP-ethanolamine and DAG. The amounts of PE methyltransferase Cho2 was also decreased likely as a compensation effect to maintain the normal PE level (Fig. 4A).
Mitochondrial crista formation is prerequisite for normal CL accumulation
Importantly, the observation that CL was hardly present in mgm1Δdnm1Δmdm35Δ cells as compared with the mgm1Δdnm1Δ and fzo1Δdnm1Δmdm35Δ cells (Fig. 2) cannot be ascribed to the decrease in the levels of CL synthetic enzymes because the steady-state levels of CL synthetic enzymes in mgm1Δdnm1Δmdm35Δ cells were comparable with those in mgm1Δdnm1Δ and fzo1Δdnm1Δmdm35Δ cells (Fig. 4A). Rather, these results could be related to significant loss of crista structures in mgm1Δdnm1Δmdm35Δ cells, but not in mgm1Δdnm1Δ or fzo1Δdnm1Δmdm35Δ cells (Fig. 3A). The possibility that decreased CL levels, in turn, result in abnormal crista morphology can be ruled out because the crista structures were not so much affected in crd1Δ cells, in which CL was absent (Baile et al., 2014). Therefore, the loss of crista structures appears to negatively affect the CL synthesis and/or maintenance.
To test whether abnormal crista morphology negatively affects the mitochondrial CL abundance, we analyzed the growth and PL compositions of mgm1Δdnm1Δmic60Δ cells, in which crista formation would be drastically impaired (Harner et al., 2016). We confirmed that crista structures largely disappeared in mgm1Δdnm1Δmic60Δ mitochondria while fzo1Δdnm1Δmic60Δ possessed crista tubules without CJs like mic60Δ cells (Fig. 4B, C). Like the mdm35Δ mutation, the mic60Δ mutation also led to severe growth defects and 80% reduction in the CL level for mgm1Δdnm1Δ cells as compared with those in wild-type cells whereas it only mildly affected the CL abundance of fzo1Δdnm1Δ cells (Fig. 4D, E). In addition, we confirmed that the loss of Mic60 did not affect steady-state levels of CL synthetic enzymes such as Tam41, Gep4, and Crd1 (Fig. 4F). These findings strongly suggest that the abnormality of crista structures causes decreases in the CL levels.
How does the impaired crista formation negatively impact on the CL biogenesis? A simple explanation is that CL is destabilized due to a shortage of the MIM volume as a reservoir of CL if a considerable amount of CL can be synthesized. Another possibility is that CL synthesis itself is impaired when normal crista structures are disrupted. The latter scenario may indicate the presence of a specialized MIM region for CL synthesis, where CL synthetic enzymes, as well as precursor PLs such as PA, CDP-DAG, and PG, are concentrated for efficient PL synthesis. Alternatively, the abnormal MIM structures may disrupt the MOM-MIM contact sites, which would hamper the alternative MOM-to-MIM transfer route for the CL precursor PA that is independent of the Ups1-Mdm35 complex.
STAR★METHODS
CONTACT FOR REAGENT AND RESOURCE SHARING
Further information and requests for resources and reagents should be directed to and will be fulfilled by the Lead Contact, Yasushi Tamura (tamura@sci.kj.yamagata-u.ac.jp).
EXPERIMENTAL MODEL AND SUBJECT DETAILS
In this study, we used a Saccharomyces cerevisiae strain, FY833 (MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63) as a background strain. All the yeast cells used in this study are listed in the Key Resources Table.
KEY RESOURCES TABLE
REAGENT or RESOURCE | SOURCE | IDENTIFIER |
---|---|---|
Antibodies | ||
Rabibit polyclonal anti-Dnm1 | Endo Lab | N/A |
Rabibit polyclonal anti-Fzo1 | Endo Lab | N/A |
Rabibit polyclonal anti-Mgm1 | Endo Lab | N/A |
Rabibit polyclonal anti-Tam41 | Endo Lab | N/A |
Rabibit polyclonal anti-Cho1 | Endo Lab | N/A |
Rabibit polyclonal anti-Cho2 | Tamura Lab | N/A |
Rabibit polyclonal anti-Opi3 | Sesaki Lab | N/A |
Rabibit polyclonal anti-Psd1 | Tamura Lab | N/A |
Rabibit polyclonal anti-Tom70 | Sesaki Lab | N/A |
Rabibit polyclonal anti-Tom40 | Endo Lab | N/A |
Rabibit polyclonal anti-Tom20 | Endo Lab | N/A |
Rabibit polyclonal anti-Tim23 | Endo Lab | N/A |
Rabibit polyclonal anti-Tim44 | Endo Lab | N/A |
Rabibit polyclonal anti-Pam16 | Endo Lab | N/A |
Rabibit polyclonal anti-Pam18 | Endo Lab | N/A |
Rabibit polyclonal anti-Kgd1 | Endo Lab | N/A |
Rabibit polyclonal anti-Ssc1 | Endo Lab | N/A |
Rabibit polyclonal anti-Mdj1 | Endo Lab | N/A |
Rabibit polyclonal anti-Om45 | Endo Lab | N/A |
Rabibit polyclonal anti-Cytb2 | Endo Lab | N/A |
Rabibit polyclonal anti-Cytc1 | Endo Lab | N/A |
Rabibit polyclonal anti-F1β | Endo Lab | N/A |
Rabibit polyclonal anti-Porin | Endo Lab | N/A |
Rabibit polyclonal anti-Mic60 | Endo Lab | N/A |
Rabibit polyclonal anti-HA | MBL Lifescience | Cat# M180-3; |
RRID:AB_10951811 | ||
ANTI-FLAG M2 Monoclonal Antibody | Sigma-Aldrich | Cat# F3165; RRID:AB_259529 |
Cy5 AffiniPure Goat Anti-Rabbit IgG (H+L) | Jackson ImmunoResearch Labs | Cat# 111-175-144; RRID:AB_2338013 |
Goat anti-Mouse IgG1 Secondary Antibody, Alexa Fluor 488 conjugate | Thermo Fisher Scientific | Cat# A-21121, RRID:AB_2535764 |
Goat anti-Rabbit IgG (H+L) Cross-Adsorbed Secondary Antibody, HRP | Thermo Fisher Scientific | Cat# A16104, RRID:AB_2534776) |
Goat anti-Mouse IgG (H+L) Cross-Adsorbed Secondary Antibody, HRP | Thermo Fisher Scientific | Cat# A16072; RRID:AB_2534745 |
Bacterial and Virus Strains | ||
XL2-blue | N/A | N/A |
Chemicals, Peptides, and Recombinant Proteins | ||
Valinomycin | Wako | Cat# 228-01121 |
JC-1 | Thermo Fisher Scientific | Cat# T3168 |
Zymolyase-20T | Nakalai tesque | Cat# 07663-91 |
TnT® Quick Coupled Transcription/Translation System | Promega | Cat# L1170 |
Met-35S-Label, [L-Methionine 35S+L-Cysteine 35S] | American Radiolabeled Chemicals Inc. | Cat# ARS0110 |
NADH | Sigma-Aldrich | Cat# N8129 |
ATP | Sigma-Aldrich | Cat# A7699 |
BSA fatty acid free | Nakalai tesque | Cat# 08587-26 |
Cycloheximide | Wako | Cat# 037-20991 |
TLC plates (SILGUR-25-C UV254) | MACHEREY-NAGEL | Cat# 810123 |
PVDF membrane (Immobilon-P) | Merck-Millipore | Cat# IPVH00010 |
PVDF membrane (Immobilon-FL) | Merck-Millipore | Cat# PFL00010 |
Experimental Models: Organisms/Strains | ||
All listed below are Saccharomyces cerevisiae strains | ||
FY833- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 | Winston et al., 1995 | N/A |
FY833 ρ°- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 ρ° | This study | N/A |
dnm1Δ- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 dnm1Δ∷URA3 | This study | N/A |
ups1Δ- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 ups1Δ∷kanMX4 | Sesaki et al., 2006 | N/A |
dnm1Δups1Δ- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 dnm1Δ∷URA3 ups1Δ∷kanMX4 | This study | N/A |
ups2Δ- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 ups2A∷HIS3 | Tamura et al., 2009 | N/A |
dnm1Δups2Δ- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 dnm1Δ∷URA3 ups2A∷HIS3 | This study | N/A |
mdm35Δ- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 mdm35Δ∷URA3 | Tamura et al., 2010 | N/A |
dnm1Δmdm35Δ- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 dnm1Δ∷kanMX4 mdm35Δ∷HIS3 | This study | N/A |
fzo1Δ- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 fzo1Δ∷kanMX4 | This study | N/A |
fzo1Δdnm1Δ- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 fzo1Δ∷kanMX4 dnm1Δ∷kanMX4 | Sesaki et al., 2006 | N/A |
fzo1Δdnm1Δups1Δ- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 fzo1Δ∷kanMX4 dnm1Δ∷kanMX4 ups1Δ∷HIS3 | This study | N/A |
fzo1Δdnm1Δups2Δ- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 fzo1Δ∷kanMX4 dnm1Δ∷kanMX4 ups2Δ∷HIS3 | This study | N/A |
fzo1Δdnm1Δmdm35Δ- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 fzo1Δ∷kanMX4 dnm1Δ∷kanMX4 mdm35Δ∷HIS3 | This study | N/A |
mgm1Δ- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 mgm1Δ∷kanMX4 | This study | N/A |
mgm1Δdnm1Δ- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 mgm1Δ∷kanMX4 dnm1Δ∷kanMX4 | Sesaki et al., 2006 | N/A |
mgm1Δdnm1Δups1Δ- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 mgm1Δ∷kanMX4 dnm1Δ∷kanMX4 ups1Δ∷HIS3 | This study | N/A |
mgm1Δdnm1Δups2Δ- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 mgm1Δ∷kanMX4 dnm1Δ∷kanMX4 ups2Δ∷HIS3 | This study | N/A |
mgm1Δdnm1Δmdm35Δ- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 mgm1Δ∷kanMX4 dnm1Δ∷kanMX4 mdm35Δ∷HIS3 | This study | N/A |
mic60Δ- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1- Δ63 mic60Δ∷hphMX | This study | N/A |
mic60Δmdm35Δ- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 mic60Δ∷hphMX mdm35Δ∷kanMX4 | This study | N/A |
mic10Δ- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 mic10Δ∷hphMX | This study | N/A |
mic10Δmdm35Δ- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 mic10Δ∷hphMX mdm35Δ∷kanMX4 | This study | N/A |
dnm1Δmic60Δ- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 dnm1Δ∷kanMX4 mic60Δ∷hphMX | This study | N/A |
crd1Δ- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 dnm1Δ∷kanMX4 crd1Δ∷kanMX4 | This study | N/A |
fzo1Δdnm1Δmic60Δ- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 fzo1Δ∷kanMX4 dnm1Δ∷kanMX4 mic60Δ∷hphMX | This study | N/A |
mgm1Δdnm1Δmic60Δ- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 mgm1Δ∷kanMX4 dnm1Δ∷kanMX4 mic60Δ∷hphMX | This study | N/A |
Gep4-HA FLAG-Crd1- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 GEP4-3HA∷TRP1 3FLAG-CRD1 | This study | N/A |
Gep4-HA FLAG-Crd1 fzo1Δdnm1Δ- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 GEP4-3HA∷TRP1 3FLAG-CRD1 fzotΔ∷kanMX4 dnm1Δ∷kanMX4 | This study | N/A |
Gep4-HA FLAG-Crd1 mgm1Δdnm1Δ- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 GEP4-3HA∷TRP1 3FLAG-CRD1 mgm1Δ∷kanMX4 dnm1Δ∷kanMX4 | This study | N/A |
Gep4-HA FLAG-Crd1 mgm1Δdnm1Δmdm35Δ- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 GEP4-3HA∷TRP1 3FLAG-CRD1 mgm1Δ∷kanMX4 dnm1Δ∷kanMX4 mdm35∷HIS3 | This study | N/A |
Gep4-HA FLAG-Crd1 fzo1Δdnm1Δmic60Δ- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 GEP4-3HA∷TRP1 3FLAG-CRD1 fzo1Δ∷kanMX4 dnm1Δ∷kanMX4 mic60Δ∷hphMX | This study | N/A |
Gep4-HA FLAG-Crd1 mgm1Δdnm1Δmic60Δ- MATa ura3-52 his3-Δ200 leu2-Δ1 lys2-Δ202 trp1-Δ63 GEP4-3HA∷TRP1 3FLAG-CRD1 mgm1Δ∷kanMX4 dnm1Δ∷kanMX4 mic60Δ∷hphMX | This study | N/A |
Oligonucleotides | ||
AGTATTGTCTGGCTTCTGAGACGGCGGTAAGATATCCTTAAGAGTTGCAAGATTGTACTGAGAGTGCAC | This study | PNU80 |
CAATTTCGCTCGCCCATGGTGATATCTTTAAAGATCTTTAAATGGGAACACTGTGCGGTATTTCACACCG | This study | PNU81 |
AATAGATTTCAGACTAAGATAAAATAATCGAGAATAATTAAAAGACGATAAGATTGTACTGAGAGTGCAC | This study | PNU82 |
AGGAATATAAGTAGTATGCAGTGCCATGCGGGATCAAGGAATTTGTATCTCTGTGCGGTATTTCACACCG | This study | PNU83 |
GAGTTTATCATTAAGTAGCTACCAGCGAATCTAAATACGACGGATAAAGAGTTGTAAAACGACGGCCAGT | This study | PNU1076 |
ATCACGCCCGCAATGTTGAAGTAAGATCAAAAATGAGATGAATTATGCAACACAGGAAACAGCTATGACC | This study | PNU1077 |
ATCTGTTTGTGTTTTAACTTGAATTACAATAACAATAATACCAGTTTTATGTTGTAAAACGACGGCCAGT | This study | PYU674 |
ATTAGTATTTACATGTTGAATAATGCACATTCTGTGCTAAAATATATACTCACAGGAAACAGCTATGACC | This study | PYU675 |
ATTAACTTTGCTACGAGAGGGAATAAACACGGAAAAAGACAAAATATACCGTTGTAAAACGACGGCCAGT | This study | PYU736 |
ATTTTTATTATTTTTTTTTTTGAATATATATAAAGCATCGTCGCTTAAGACACAGGAAACAGCTATGACC | This study | PYU737 |
ACACAAAAGGCATAAGAACGCATTGAAAAGTCTAAAAAACTAATATTCGTGTTGTAAAACGACGGCCAGT | This study | PYU740 |
GGAAATTGAGGTGTAATGACGTACATCTCTTTTCTCTTTGTATTATTCTTCACAGGAAACAGCTATGACC | This study | PYU741 |
CTCCAAGTTCGAAAAAAAATTATACAACTTTTTGGGATTTCGGATCCCCGGGTTAATTAA | This study | PYU972 |
AAAAAATTAAAATGTTTTACTTTTTATTAAAGTTGCCTAAGAATTCGAGCTCGTTTAAAC | This study | PYU973 |
AATTGGATCCAATTTAGTCAATGATTCAAATGG | This study | PYU1550 |
CAGTAAGCTTCTATTTTAAAAGTTTAAAAGCG | This study | PYU1551 |
CCCCGAGCTCCGCTCAAATCCTTCTTGGTCGTTATT | This study | PJH189 |
CCCCGGATCCTCATTTTAAATCATTCTTTCCAATTATG | This study | PJH190 |
Recombinant DNA | ||
pBS-kanMX4 | Endo Lab | N/A |
pBS-hphMX | Endo Lab | N/A |
pBS-CgHIS3 | Endo Lab | N/A |
pGEM-4Z | Promega | Cat#: P2161 |
pGEM-4Z-Crd1 | This study | N/A |
pGEM-4Z-Psd1 | This study | N/A |
pGEM-4Z-Hsp60 | Tamura et al., 2009 | N/A |
pGEM-4Z-Tim23 | Tamura et al., 2009 | N/A |
pGEM-4Z-AAC | Tamura et al., 2009 | N/A |
pFA6a-3xHA-TRP1 | Longtine et al., 1998 | N/A |
pAUR135 | Takara Bio Inc. | N/A |
pAUR135-FLAG-Crd1 | This study | N/A |
pRS314-GAL1-FLAG-Crd1 | This study | N/A |
METHOD DETAILS
Strains, plasmids, primers, and growth conditions
Disruption or chromosomal C-terminal tagging of the yeast gene was accomplished by PCR-mediated gene replacement (Longtine et al., 1998) with a pair of primers listed in the KEY RESOURCES TABLE. The kanMX4 (pBS-kanMX4), hphMX (pBS-hphMX), HIS3 (pBS-CgHIS3), URA3 (pRS306) genes were used as disruption markers. For C-terminal HA tagging of GEP4 gene, the 3xHA-TRP1 gene was amplified by PCR using pFA6a-3xHA-TRP1 as a template (Longtine et al., 1998). For N-terminal FLAG tagging into CRD1 gene, we used a shuttle vector pAUR135, in which the Aureobasidin A-resistance gene (AUR1-C) and GIN11M86 under the control of GAL10 promoter were cloned. Since overexpression of GIN11M86 causes growth inhibition, cultivation on galactose-containing media allowed the transformant cells to remove the marker genes from the genomic DNA. (Takara Bio Inc.). Synthetic gene encoding Crd1(1-60)-3xFLAG tag-Crd1(61-283) purchased from Eurofins Genomics was cloned into pAUR135, resulting in pAUR135-FLAG-Crd1. After a single cutting of the plasmid at BglII site present in the N-terminal side of CRD1, the resulting linear DNA was introduced into yeast cells and Aureobasidin A-resistance transformants were selected. The transformants were streaked on YPGa1 plate to select marker-removed clones to obtain yeast cells expressing FLAG-Crd1.
A pair of primers, PNU80/81, PNU82/83, PYU674/675, PYU736/737, PYU740/741, PNU1076/1077, or PYU972/973 was used to amplify the gene cassette for ups1Δ, nps2Δ, mdm35Δ, mic10Δ, mic60Δ, dnm1Δ mutation or HA-tagging to the GEP4 gene, respectively.
To construct pGEM-4Z-Crd1 and pGEM-4Z-Psd1, the CRD1 and PSD1 genes were amplified by PCR using the yeast genomic DNA as a template and a pair of primers PYU1550/1551 and PJH189/190, respectively. The CRD1 and PSD1 fragments were digested with BamHI/HindIII and SacI/BamHI, and cloned to the multi-cloning site of pGEM-4Z.
Yeast cells were grown in YPD (1% yeast extract, 2% polypeptone, and 2% glucose), YPLac (1% yeast extract, 2% polypeptone, and 3% lactate), YPGal (1% yeast extract, 2% polypeptone, and 3% galactose), SCD (0.67% yeast nitrogen base without amino acids, 0.5% casamino acid, and 2% glucose), SGalSuc (0.67% yeast nitrogen base without amino acids, 2% galactose, and 2% sucrose) supplemented with 20 μg/ml each of adenine, uracil, L-tryptophan and L-histidine and 30 μg/ml each of L-leucine and L-lysine. YPD supplemented with 200 μg/ml G418, 200 μg/ml Hygromycin B or 100 μg/ml Aureobasidin were used for selection of kanMX4, hphMX6 or AURa-C-containing transformants, respectively.
Phospholipid analysis
Yeast cells were diluted to an A600 = 0.02 in 1 ml of YPD in the presence of 2 μ Ci/ml 32Pi and cultivated to stationary phase. Phospholipids were extracted from total cells and separated by TLC with developing solvent A (chloroform/ethanol/water/triethylamine, 30/35/7/35, v/v) or solvent B (chloroform/methanol/25% ammonia, 65/35/5, v/v) (Tamura et al., 2013). After 2 to 3 hours developing, the TLC plates were dried and sandwiched with storage phosphor screens using an exposure cassette (GE healthcare). The RI-labeled phospholipids were detected by radioimaing with Typhoon FLA-7000 image analyzer (GE Healthcare).
Electron microscopy
For the EM analyses in Fig. 3, yeast cells grown in SGalSuc until early log phase (A600 = 0.3 – 0.5) were fixed and embedded in Epon resin, as described (Itoh et al., 2013). Briefly, yeast cells were fixed by perfusing 2% glutaraldehyde, 1 mM CaCl2, and 0.1 M sodium cacodylate, pH 7.2 for 1 hr. After digestion of cell wall by Zymolyase 20T, cells were further fixed in 0.5% osmium teroxide, and 0.8% potassium ferrocyanide. After washing in 0.1 M sodium cacodylate, pH 7.2, cells were embedded in 2.5% low-melting agarose and post-fixed in 1% uranyl acetate for 1.5 hrs. After dehydration using 50, 70, 90, and 100% ethanol and 100% propylene oxide, samples were embedded in Epon resin (Ted Pella). Ultrathin sections were obtained using and Ultracut E (Reichert-Jung, Germany). Sections were stained with 2% uranyl acetate and 3 mg/ml lead citrate, and observed on a Hitachi H-7600 transmission electron microscope (Hitachi, Japan).
For the EM analyses in Fig. 4, yeast cells were fixed by perfusing 2% paraformaldehyde 2% glutaraldehyde, and 0.1 M sodium cacodylate, pH 7.4 at room temperature for 1 h, then transferred to 4°C. The cells were collected and further fixed with 2% glutaraldehyde, and 0.1 M Sodium cacodylate, pH 7.4 at 4°C overnight. After fixation, the samples were washed 3 times with 0.1 M sodium cacodylate, pH 7.4 for 10 min each, and were post-fixed with 1% KMnO4 at room temperature for 2 h. The samples were dehydrated in graded 50% and 70% ethanol solutions for 30 min each at 4°C, 90% for 30 min at room temperature, and then 100% for 30 min at room temperature. The samples were infiltrated with propylene oxide (PO) twice for 30 min each and were put into a 70:30 mixture of PO and resin (Quetol-812; Nisshin EM Co.) for 1 h. PO was volatilized by keeping the cap open overnight. The samples were transferred to a fresh 100% resin, and were polymerized at 60°C for 48 h. The polymerized resins were ultrathin sectioned at 70 nm with a diamond knife using a ultramicrotome (Ultracut UCT, Leica) and the sections were mounted on copper grids. They were stained with 2% uranyl acetate at room temperature for 15 min, and then washed with distilled water followed by being secondary-stained with Lead stain solution (Sigma) at room temperature for 3 min. The grid was observed by a transmission electron microscope (JEM-1400Plus, JEOL) wtih an acceleration voltage of 100 kV. Digital images were taken with a CCD camera (EM014830RUBY2, JEOL).
Isolation of membrane fractions
Yeast cells were cultivated to stationary phase in 50 ml YPD at 30°C. Cells were harvested and incubated in alkaline buffer (0.1 M Tris-HCl, pH 9.5, 10 mM dithiothreitol (DTT)) for 15 min at 30°C. After washing with spheroplast buffer (20 mM Tris-HCl, pH 7.5, 1.2 M sorbitol), the cells were treated with Zymolyase 20T (100 μg/ml final) in spheroplast buffer for 40 min at 30°C. The resulting spheroplasts were re-suspended in ice-cold lysis buffer (20 mM Tris-HCl, pH 7.5, 0.6 M sorbitol, 1 mM phenylmethylsulfonyl fluoride (PMSF)) and frozen in liquid nitrogen. Spheroplasts were then disrupted by rotational vibration at 4,260 rpm for 90 seconds using a cell destroyer PS 1000 (BMS, Japan). After removing unbroken cells by centrifugation at 2,000 × g for 5 min, membrane fractions containing mitochondria were precipitated by centrifugation at 15,000 × g for 5 min.
Immunoblotting
For immunoblotting, proteins separated on SDS gels were transferred to PVDF membranes (Immobilon-FL or Immobilon-P, Millipore). Then, the membranes were incubated with primary antibodies as stated below and specific proteins were detected by Cy5, Alexa Fluor 488 or HRP-conjugated secondary antibodies, goat anti-rabbit or mouse IgG (H+L) (Thermo Fisher Scientific) and analyzed with Typhoon imager (GE Healthcare) or LAS-4000mini (FUJIFILM Corporation).
Supplementary Material
Acknowledgments
We thank Kayoko Shishido, Tomomi Sasaki and Michiko Hashimoto for technical assistance and Dr. Yasunori Watanabe for pRS314-GAL-FLAG-Crd1 plasmid. We are grateful to the members of the Tamura, Endo and Sesaki laboratories for valuable discussions. This work was supported by JSPS KAKENHI (Grant Numbers 15H05595 and 17H06414 to YT, and 15H05705 and 22227003 to TE), National Institutes of Health Grants to H.S. (GM123266), AMED-PRIME from Japan Agency for Medical Research and Development, AMED (Grant Number 18gm5910026h0002) (YT), and a CREST Grant from JST (TE).
Footnotes
DECLARATION OF INTERESTS
The authors declare no competing interests.
SUPPLEMENTAL INFORMATION
Supplemental Information includes three figures can be found with this article online.
Reference
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