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Open Access

Peer-reviewed

Research Article

Calnexin Regulates Apoptosis Induced by Inositol Starvation in Fission Yeast

Abstract

Inositol is a precursor of numerous phospholipids and signalling molecules essential for the cell. Schizosaccharomyces pombe is naturally auxotroph for inositol as its genome does not have a homologue of the INO1 gene encoding inositol-1-phosphate synthase, the enzyme responsible for inositol biosynthesis. In this work, we demonstrate that inositol starvation in S. pombe causes cell death with apoptotic features. This apoptotic death is dependent on the metacaspase Pca1p and is affected by the UPR transducer Ire1p. Previously, we demonstrated that calnexin is involved in apoptosis induced by ER stress. Here, we show that cells expressing a lumenal version of calnexin exhibit a 2-fold increase in the levels of apoptosis provoked by inositol starvation. This increase is reversed by co-expression of a calnexin mutant spanning the transmembrane domain and C-terminal cytosolic tail. Coherently, calnexin is physiologically cleaved at the end of its lumenal domain, under normal growth conditions when cells approach stationary phase. This cleavage suggests that the two naturally produced calnexin fragments are needed to continue growth into stationary phase and to prevent cell death. Collectively, our observations indicate that calnexin takes part in at least two apoptotic pathways in S. pombe, and suggest that the cleavage of calnexin has regulatory roles in apoptotic processes involving calnexin.

Citation: Guérin R, Beauregard PB, Leroux A, Rokeach LA (2009) Calnexin Regulates Apoptosis Induced by Inositol Starvation in Fission Yeast. PLoS ONE 4(7): e6244. https://doi.org/10.1371/journal.pone.0006244

Editor: Andreas Bergmann, University of Texas MD Anderson Cancer Center, United States of America

Received: March 23, 2009; Accepted: June 8, 2009; Published: July 16, 2009

Copyright: © 2009 Guérin et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Funding: This work was supported by grants of the Canadian Institutes of Health Research (Grants MOP-62703 and MOP-89702) to LAR. RG and PBB, received scholarships from the Faculte des etudes superieures-Department of Biochemistry, Universite de Montreal. PBB received an NSERC PhD studentship. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Competing interests: The authors have declared that no competing interests exist.

Introduction

The endoplasmic reticulum (ER) is a central organelle playing crucial roles in several cellular processes. The ER is at the center of the synthesis and the folding of secreted, membrane-bound and some organelle-targeted proteins [1], [2]. To assist in the protein folding process, the ER contains molecular chaperones, several co-factors such as ATP and Ca2+, and an optimal oxidizing environment to facilitate disulphide-bond formation [1], [2], [3]. In addition to its protein folding roles in the secretory pathway, the ER is crucial for other fundamental cellular processes including lipid biosynthesis, membrane biogenesis, and Ca2+ storage [4].

Perturbations in the ER homeostasis by stresses such as lipid and glycolipid imbalances, alterations in the levels of Ca2+, or modification in redox state, negatively affect the protein folding capacity of this organelle [5], [6], [7]. These adverse conditions, referred to as ER stress, result in the accumulation and aggregation of unfolded or incompletely folded proteins. The functions of the ER are tightly regulated. To counteract ER stress and restore its full protein-folding capacity, the ER responds by inducing a stress-response pathway called UPR, for Unfolded Protein Response [8], [9]. The UPR mechanism is well conserved from yeast to mammals. The UPR stops general protein synthesis and stimulates the transcription of genes coding for ER-folding factors such as molecular chaperones and foldases [5], [6], [7], [10], [11]. Concomitantly, the cell improves the ERAD (ER-Associated Degradation) pathway in order to degrade the unfolded proteins present is the ER [12], [13], [14]. These actions allow the ER to stabilize its environment and ensure cell survival. A major regulator of the UPR is the ER transmembrane protein Ire1p (Inositol requiring kinase I) which is conserved from fungi to mammals [8], [9]. Ire1p is an endoribonuclease that is activated by homodimerization and autophosphorylation, and whose downstream effect is to stimulate the transcription of genes encoding ER chaperones and other factors involved in all stages of the secretory pathway. So far, Ire1p is the only factor identified in yeast to transduce the UPR, but mammalian cells contain additional transducers called PERK and ATF6 [9], [15]. If this initial suite of actions is not able to restore ER homeostasis, the UPR switches its downstream effects from pro-survival to pro-death. Prolonged ER stress induces via IRE1, PERK and ATF6 an apoptotic pathway involving the Bcl-2 family of proteins, which act at the level of the mitochondria and the ER [6], [7], [16].

Apoptosis is a central molecular process first identified in multicellular organisms for its crucial roles in development and in regulation of many diseases. Apoptosis is a tightly regulated form of programmed cell death that is characterized by specific biochemical and morphological features such as cell rounding and shrinkage, chromatin breakage, nuclear fragmentation and activation of caspases [17], [18]. Mounting evidence accumulated in the last ten years established that unicellular organisms such as yeasts undergo apoptotic cell death [19], [20], [21]. Apoptosis in yeast is induced by numerous conditions including DNA damage, aging, replication defects, deficiency in triacylglycerols, ER stress, and mating [19], [22]. The yeast genomes encode several homologues of proteins characterized for their involvement in apoptosis including Yca1/Pca1, AIF, EndoG, HtrA2/Omi and IAP [23], [24], [25], [26], [27], [28]. As is for mammalian cells, the involvement of these factors and their interaction in different apoptotic pathways are under current investigation.

Inositol is a precursor for numerous molecules including inositol-containing phospholipids, inositol esters and phosphorylated versions of inositol playing central roles in membrane integrity, cell signalling and vesicular traffic [29], [30], [31]. The pathway involving inositol 1,4,5-triphosphate (IP3) has been extensively characterized. IP3 derives from the hydrolysis of phosphatidylinositol 4,5-biphosphate (PIP2) by phospholipase C (PLC), thereby producing IP3 and DAG (diacylglycerol). IP3 acts a second messenger via its binding to the IP3R receptor on the ER membrane [32]. The binding of IP3 to its receptor provokes the release of the ER-stored calcium into the cytoplasm, which in turn elicits a range of cellular responses [33]. Calcium release from vacuoles following IP3-signalling was also observed in Saccharomyces cerevisiae [34]. Inositol can be synthesized by intracellular processes involving conversion of glucose-6-phosphate (G6P) into inositol monophosphate (IP1) through a set of complex reactions of oxidation/reduction which are mediated by a unique enzyme, the inositol-1-phosphate synthase (INO1) [35], [36]. The INO1 gene was identified in various species of yeasts, protozoa, plants and mammals [37], [38], [39]. Deletion of INO1 and mutation in other genes such as IRE1 cause inositol auxotrophy in S. cerevisiae [40], [41]. In this yeast, expression of INO1 is under the control of IRE1 via HAC1, two key players of the UPR pathway [42], [43].

Schizosaccharomyces pombe is considered a good model to study the involvement of inositol in cell pathways because this yeast is naturally auxotroph for inositol due to the absence of a gene coding for an inositol-1-phosphate synthase [44], [45]. Studies in S. pombe have shown that absence of inositol in the culture medium is lethal for this yeast, and that partial depletion provokes sexual sterility with no effects on growth [46], [47], [48], [49], [50], [51]. Although S. pombe cells die in the absence of inositol, they are able to survive longer than S. cerevisiae cells auxotroph for inositol as result of genetic manipulation (Δino1 cells) [49].

Calnexin is an ER transmembrane chaperone playing key roles in translocation, in protein folding, and in the quality control of newly synthesized polypeptides [1], [2], [52]. Structurally, calnexin is a type I transmembrane protein of the ER containing a large lumenal domain, a transmembrane domain (TM), and a short cytosolic tail. The lumenal domain folds into a globular structure formed by the C- and N-terminal extremities, and a hairpin structure formed by the highly conserved central domain (hcd), which is the most conserved calnexin domain across species. Calnexin interacts with client proteins via glycan-lectin or protein-protein interactions [52], [53], [54], [55], [56], [57], [58], [59], [60], [61].

The knockout of calnexin in mice causes early postnatal death and severe motor disorders and is lethal in S. pombe, thus pointing to the critical cellular roles of this protein [62], [63], [64]. We showed that certain calnexin chaperone-deficient mutants are viable. Interestingly, this demonstrates that the essentiality of calnexin is not its chaperone activity but another yet to be defined cellular role [58], [65], [66].

Several studies published in the recent years indicate that calnexin is involved in apoptotic processes induced by ER stresses. First indications came from a report showing that the cytosolic tail of S. pombe calnexin is required for cell death mediated by the heterologous expression of mammalian Bak, suggesting that calnexin can form a complex with lethal partners in apoptotic situations [67]. In mammalian cells, it was shown that calnexin-deficient cells are more resistant to apoptosis [68], [69]. It was suggested that calnexin could act as a scaffold for the cleavage of the ER transmembrane apoptotic protein Bap31 by caspase 8 in ER-stress conditions [70]. In addition, calnexin in mammalian cells was shown to be sensitive to caspase cleavage under stress conditions [71]. It was proposed that this cleavage could have a role in the transduction of an apoptotic signal. More recently, we showed that overexpression of calnexin in S. pombe causes apoptosis and that this induction requires the anchoring of calnexin to the ER membrane [72]. We also demonstrated that apoptosis induced by tunicamycin is less efficient in cells containing only a lumenal version of calnexin, thus pointing to the importance of both the membrane anchoring of calnexin and of its cytosolic tail.

Here, we demonstrate that inositol starvation induces cell death in S. pombe via an apoptotic pathway dependent on the metacaspase Pca1p and that is modulated by the UPR transducer Ire1p. We show that calnexin is cleaved under normal conditions when cells approach stationary phase. While a lumenal version of calnexin exacerbates the apoptosis provoked by inositol starvation, co-expression of a mutant spanning the TM and C-terminal tail significantly reduces the levels of apoptosis. Our work suggests that the TM and C-terminal tail of calnexin are involved in apoptotic signalling when inositol is depleted.

Results

Inositol starvation induces apoptotic cell death in S. pombe

Inositol starvation induces cell death in fission yeast [44], [45]. This effect is due to the inability of S. pombe to synthesize inositol because of the lack of a homologue the INO1 gene in its genome. Inositol starvation in S. cerevisiae also provokes cell death when INO1 is defective, indicating a crucial role of inositol in the physiology of the cell. To characterize the death resulting from inositol depletion in S. pombe, we first measured the capacity of cells to grow on media containing inositol after 48 hours of culture in media without inositol. As expected, cells cultured in media deprived of inositol show a dramatic reduction in the ability to form colonies compared to cells cultured in media containing inositol (Figure 1A). The death phenotype was confirmed by staining with the fluorescent vital dye Phloxin B, followed by quantification by fluorescence-activated cell sorting (FACS). As shown in Figure 1B, about 25% of cells are stained after 12 hours of inositol starvation and more than 50% after 24 hours, as compared to near 0% for the unstarved cells. The cell death observed following inositol starvation is specific since starvation in adenine or leucine for 12 h and 24 h do not promote cell death (Supplemental Figure S1). Next, to determine the type of cell death we measured apoptotic markers in cells starved for inositol. As we have previously demonstrated, S. pombe cells undergoing apoptotic death show specific phenotypes including metacaspase activation, DNA breakage and nuclear fragmentation [72]. Using the fluorescent probe FITC-VAD-fmk in FACS analyses, cells submitted to inositol starvation for 12 and 24 hours show respectively 15 and 50% of fluorescence, compared to near 0% for cells that were not starved (Figure 1C). Fluorescence microscopy with DAPI revealed nuclear fragmentation for cells cultured for 48 h without inositol (Figure 1D). By contrast, cells cultured in inositol-containing medium exhibited round and intact nuclei. Consistent with the DAPI staining results, a TUNEL assay shows that only cells starved for inositol display DNA breakage (Figure 1E). Taken together, these results confirm that depletion of inositol is lethal for S. pombe and indicate that this death is mediated via an apoptotic mechanism.

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Figure 1. Inositol starvation induces apoptotic cell death.

(A) Survival of cells cultured for 48 h in MM with or without inositol was assayed by serial dilution on media containing inositol. Samples of 10 µl of four 10-fold serial dilutions (10−1–10−4) of cells at OD595 1 were spotted on selective MM with inositol, and incubated at 30°C for 7 days (see Materials and Methods). (B) Percentage of dead cells measured by staining with the fluorescent vital dye Phloxin B. Cells cultured for 24 h in MM containing inositol or for 12 h and 24 h in MM without inositol, were stained with Phloxin B and fluorescent cells were quantified by FACS. Stained cells were considered as dead. (C) Metacaspase activation. The fluorescent probe FITC-VAD-FMK was used to measure metacaspase activation by FACS after 12 and 24 h of growth in MM without inositol, or after 24 h of growth in MM containing inositol, as described in Materials and Methods. (D) Nuclear fragmentation. Cells cultured for 48 h in MM with or without inositol were stained with DAPI and visualized under fluorescence microscopy; Nomarski fields are shown. White arrows indicate fragmented nuclei. (E) DNA breakage. TUNEL assay was carried out with cells cultured for 48 h in MM with or without inositol. The percentage of stained cells was measured by FACS. The significance of differences in the results was evaluated by a Student's t test pairwise calculated between the different conditions assayed. **p<0.01 and *p<0.05.

https://doi.org/10.1371/journal.pone.0006244.g001

The metacaspase Pca1p is required to mediate apoptosis induced by inositol starvation

The S. pombe genome encodes several characterized homologues of factors associated with apoptotic pathways in mammalian cells [23], [24], [25], [26], [27]. In S pombe, Pca1p is the only caspase-like protein identified so far, and the most studied factor for dependence in apoptosis [73], [74]. As this is the first time that death by inositol starvation is described as apoptotic in fission yeast, we investigated whether this process requires the metacaspase Pca1p. To this end, Δpca1 cells were cultured in inositol-less medium for 48 h and spotted on inositol-containing plates. A dramatic diminution in the death level induced by inositol starvation was observed in a Δpca1 strain compared to the wild-type control (Figure 2A). The reduction in the levels of cell death in the absence of pca1+ was confirmed by Phloxin B staining. Following 24 h of inositol starvation, the Δpca1 strain exhibited practically no Phloxin B staining compared to more than 50% for the wild-type control (Figure 2B). Metacaspase activation was measured with the fluorescent probe FITC-VAD-FMK. Congruently, a significant reduction in the level of metacaspase activation was observed in a Δpca1 background compared to the wild-type control (Figure 2C). These results demonstrate the importance of the metacaspase Pca1p in the apoptotic pathway induced by the absence of inositol.

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Figure 2. The metacaspase Pca1p is required for apoptosis induced by inositol starvation.

(A) Survival of pca1+ and Δpca1 cells cultured for 48 h in MM without inositol was assayed by serial dilution on solid media containing inositol. Samples of 10 µl of four 10-fold serial dilutions (10−1–10−4) of cells at OD595 1 were spotted on selective MM plates with inositol, and incubated at 30°C for 7 days (see Materials and Methods). (B) Percent of dead cells measured by staining with the fluorescent vital dye Phloxin B. Pca1+ and Δpca1 cells grown for 24 h in MM without inositol were stained with Phloxin B, and fluorescent cells were quantified by FACS. Stained cells were considered as dead. (C) Metacaspase activation. The fluorescent probe FITC-VAD-FMK was used to measure metacaspase activation by FACS of pca1+ and Δpca1 cells after 24 h of culture in MM without inositol, as described in Materials and Methods. The significance of differences in the results was evaluated by a Student's t test pairwise calculated between the different conditions assayed. **p<0.01 and *p<0.05.

https://doi.org/10.1371/journal.pone.0006244.g002

The UPR transducer Ire1p affects the apoptosis induced by inositol starvation

Inositol is essential for the survival of S. pombe because its genome does not have an INO1 homologue encoding for the inositol-1-phosphate synthase enzyme responsible for the biosynthesis of inositol from glucose-6-phosphate [35], [36], [44], [45]. In S. cerevisiae, INO1 is under the control of the UPR via IRE1 and HAC1, and the knockout of IRE1 leads to inositol auxotrophy [42], [43]. To investigate if ire1+ is involved in apoptosis induced by inositol starvation in S. pombe, Δire1 cells were cultured in the absence of inositol for 48 h and the death levels were measured by spotting on inositol containing plates. A significant improvement in the levels of survival was observed for the Δire1 strain in comparison to the wild-type control (Figure 3A). The induction of metacaspase was measured by FITC-VAD-FMK staining and FACS analysis. Here again, a dramatic reduction in the levels of metacaspase activation was observed for the Δire1 strain compared to wild-type cells. The levels of metacaspase activation dropped to about 15% in the absence of Ire1p compared to near 50% in its presence (Figure 3B). These results demonstrate that Ire1p significantly influences the apoptotic cell death induced by inositol starvation in S. pombe, albeit not totally as a certain level of death is measured in Δire1 cells.

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Figure 3. The UPR transducer Ire1p affects the apoptosis induced by inositol starvation.

(A) Survival of ire1+ and Δire1 cells cultured for 48 h in MM without inositol was assayed by serial dilution on solid media containing inositol. Samples of 10 µl of four 10-fold serial dilutions (10−1–10−4) of cells at OD595 1 were spotted on selective MM plates with inositol, and incubated at 30°C for 7 days (see Materials and Methods). (B) Metacaspase activation. The fluorescent probe FITC-VAD-FMK was used to measure metacaspase activation by FACS of ire1+ and Δire1 cells after 24 h of culture in MM without inositol, as described in Materials and Methods. The significance of differences in the results was evaluated by a Student's t test pairwise calculated between the different conditions assayed. **p<0.01 and *p<0.05.

https://doi.org/10.1371/journal.pone.0006244.g003

A lumenal version of calnexin is more sensitive to apoptosis induced by inositol starvation

We have previously demonstrated that calnexin is involved in apoptosis caused by ER stress in S. pombe [72]. To examine if calnexin is part of the death pathway induced by the absence of inositol, we compared WT cells to a strain expressing the lumenal_Cnx1p mutant which lacks the trasmembrane domain (TM) and the cytosolic tail of this ER protein. As shown in Figure 4B, in inositol starvation the cells expressing the lumenal_Cnx1p mutant as the only version of calnexin exhibited a dramatic increase in the death levels, as measured by the ability to form colonies (Figure 4A). Moreover, the lumenal_cnx1 cells showed a close to 2-fold increase in cell death measured by Phloxin B staining after 12 h of starvation (Figure 4B). Calcofluor-white is a fluorescent dye that stains a cell-wall polysaccharide, probably chitin, accumulating in the septa of S. pombe [65], [75]. Interestingly, inositol-starved lumenal_cnx1 cells accumulate large, round vesicles containing material that is highly stained with Calcofluor (Figure 4D). These vesicles were also observed when lumenal_cnx1 cells were cultured in inositol-containing media but at a much lower frequency, as compared to deprivation conditions (Figure 4D). This phenotype exacerbated by inositol starvation is also observed when calnexin mutants are submitted to heat stress in inositol-containing medium, conditions in which they exhibit cell-wall defects [65]. Thus, this Calcofluor-staining phenotype suggests a link between inositol and cell-wall biosynthesis.

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Figure 4. Increased apoptosis induced by inositol starvation in cells expressing a lumenal calnexin version.

(A) Survival of Cnx1p and lumenal_Cnx1p strains cells cultured for 48 h in MM depleted of inositol was assayed by serial dilution on media containing inositol. Samples of 10 µl of four 10-fold serial dilutions (10−1–10−4) of cells at OD595 1 were spotted on selective MM plates containing inositol, and incubated at 30°C for 7 days (see Materials and Methods). (B) Percent of dead cells measured by staining with the fluorescent vital dye Phloxin B. cnx1+ and lumenal_cnx1 strains cultured for 12 h in MM depleted in inositol were stained with Phloxin B and the fluorescent cells were quantified by FACS. Stained cells were considered as dead. (C) Metacaspase activation. The fluorescent probe FITC-VAD-FMK was used to measure metacaspase activation by FACS of cnx1+ and lumenal_cnx1 strains after 12 h of growth in MM without inositol, as described in Materials and Methods. (D) Cell morphology. Cells expressing only Cnx1p or lumenal_Cnx1p were cultured for 48 h in MM without inositol and stained with Calcofluor-white, and visualized under fluorescence microscopy; Nomarski fields are shown. White arrows indicate the vesicles containing Calcofluor-stainable material accumulating in the lumenal_cnx1 strain. The significance of differences in the results was evaluated by a Student's t test pairwise calculated between the different conditions assayed. **p<0.01 and *p<0.05.

https://doi.org/10.1371/journal.pone.0006244.g004

In agreement with the levels of cell death observed by Phloxin B, cells expressing only the lumenal version of calnexin exhibited a close to 2-fold increase in metacaspase activation following 12 h of growth in media without inositol as compared to the cnx1+ strain (Figure 4C). These results implicate calnexin in the death cascade triggered by inositol starvation, and point to a role of its TM and cytosolic tail in this apoptotic pathway.

Co-expression of the cytosolic tail and TM with lumenal_Cnx1p reduces the sensitivity to inositol starvation

Since lumenal_Cnx1p is a calnexin mutant with full chaperone activity [58], we hypothesized that the increased effect on apoptosis induced by inositol starvation could be due to the absence of the cytosolic tail and the TM. To test this hypothesis, we co-expressed lumenal_Cnx1p with the mutant C-termTM_Cnx1p_cmyc, which spans the TM and cytosolic tail of calnexin (Figure 5A). Expression of the C-termTM_Cnx1p_cmyc mutant in conjunction with the lumenal_Cnx1p reduced the cell death to wild-type levels, as measured by Phloxin B staining (Figure 5B). The same reduction was observed for the levels of metacaspase activation when lumenal_Cnx1p was co-expressed with the C-termTM_Cnx1p_cmyc mutant (Figure 5C). This particular effect is not due to variations of lumenal_cnx1 expression because the level of luminal_Cnx1p remains unchanged in the presence of a plasmid expressing the C-termTM_Cnx1p mutant (Supplemental Figure S2). Interestingly, the Calcofluor-staining phenotype observed for the lumenal_cnx1 cells submitted to inositol starvation was not completely reverted in the presence of the C-termTM_Cnx1p_cmyc mutant (Figure 5D). The levels of large round structures stained with Calcofluor in media with or without inositol is the same whether the mutant C-termTM_Cnx1p_cmyc is co-expressed or not. These observations indicate that the anchoring of the C-terminal tail of calnexin to the ER membrane is important in the response to the apoptotic signal induced by inositol starvation. However, the cell-wall defects observed by Calcofluor staining appear to be more attributable to the requirement for an intact, i.e. wild-type calnexin. The importance of calnexin in cell-wall integrity was already shown in S. cerevisiae [76], and in S. pombe as we observed previously the same defect in calnexin mutants at 37°C [65], [66], [77].

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Figure 5. C-terminalTM_Cnx1pcmyc reverses the augmentation of apoptosis induced of the lumenal_cnx1 strain caused by inositol starvation.

(A) Schematic representation of WT calnexin and of calnexin mutants used in this study. The mutant lumenal_Cnx1p spans the lumenal domain of calnexin (415 aa) and is truncated of the transmembrane domain (TM) and the cytosolic tail. The mutant C-termTM_Cnx1p_cmyc spans the TM domain (23 aa) and the cytosolic tail (48 aa) fused to the cmyc tag. (B) Percent of dead cells measured by staining with the fluorescent vital dye Phloxin B. cnx1+, lumenal_cnx1 and lumenal_cnx1+C-termTM_cnx1_cmyc strains cultured for 12 h in MM without inositol were stained with Phloxin B, and fluorescent cells were quantified by FACS. Stained cells were considered as dead. (C) Metacaspase activation. The fluorescent probe FITC-VAD-FMK was used to measure metacaspase activation by FACS of cnx1+, lumenal_cnx1 and lumenal_cnx1+C-termTM_cnx1_cmyc strains after 12 h of cultured in MM depleted of inositol, as described in Material and Methods. (D) Cell morphology. Cells expressing only Cnx1p, lumenal_Cnx1p or lumenal_Cnx1p+C-termTM_Cnx1p_cmyc were cultured for 48 h in MM without inositol and stained with Calcofluor-White, and visualized under fluorescence microscopy; Nomarski fields are shown. White arrows indicate the accumulation of Calcofluor-stained material in the lumenal_cnx1 strain. The significance of differences in the results was evaluated by a Student's t test pairwise calculated between the different conditions assayed. **p<0.01 and *p<0.05.

https://doi.org/10.1371/journal.pone.0006244.g005

Calnexin is cleaved within the lumenal domain under normal culture conditions

The TM and the cytosolic tail of calnexin are important domains in at least two situations leading to apoptosis, ER stress and inositol starvation. Interestingly, in time-course experiments to measure the levels of calnexin expression by Western blotting with WT cells, we observed that calnexin undergoes cleavage in a time-dependent manner under normal culture conditions (Figure 6A). The size downshift of calnexin begins at 12 h of culture corresponding to OD595∼1, thus in early exponential phase (Figure 6A). Complete cleavage of calnexin of calnexin is observed after 48 h of culture when cells reach an OD595 of 7. It is important to note that this downshift in size is observed in medium containing inositol, and that the same pattern is observed whether the calnexin gene is in the genome or calnexin is expressed by a plasmid in a Δcnx1 background. The same downshift in the size of calnexin was observed in media without inositol (not shown). To determine whether the cleavage occurs closer to the N- or C-terminal end, we used a version of calnexin cmyc-tagged at its C-terminus. At time-point 0 corresponding to OD595 0.3, we detected the full-length calnexin by Western blotting against cmyc. However, after 48 h of culture, corresponding to OD595 7, we detected by anti-cmyc immunoblotting a small calnexin fragment migrating at Mr of 31 kDa, which apparently is not cleaved further. This indicates that the cleavage takes place at a site approximately within C-terminal third of the protein (Figure 6B). The band observed near 31 kDa migrated more slowly than the C-termTM_Cnx1p_cmyc construct [72], indicating that calnexin is probably cleaved within its lumenal domain. An approximation of the cleavage site was obtained by analysis of a collection of different calnexin mutants created in the laboratory (not shown), and by MS/MS analysis of the cmyc-tagged small fragment of calnexin isolated by immunoprecipitation from cell extracts after 48 h of growth. This confirmed that calnexin is cleaved in its lumenal domain near the TM domain within a sequence of residues that is framed in red in Figure 6C. To further investigate the cleavage of calnexin, protein extracts from cells cultured for 24 h were fractionated by gel-filtration. Full-length calnexin and the cleaved-version of calnexin did not elute in the same fractions, indicating that they do not associate in the same protein complexes (Figure 6D). Supporting this point, the BiP chaperone eluted in the same fractions with cleaved calnexin, but did not co-elute with full-length calnexin. Collectively, these results demonstrate that under normal growth conditions calnexin is cleaved at the end of its lumenal domain giving rise to two stable moieties, probably each having a different cellular role.

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Figure 6. Calnexin is cleaved in the lumenal domain under normal conditions.

(A) Time-course cleavage of calnexin. Samples corresponding to 10 µg of protein extracts from cells expressing endogenous calnexin at different time points of growth were loaded on a 10% (wt/vol) SDS-PAGE gel. Calnexin was detected by immunoblotting with an anti-Cnx1p rabbit polyclonal serum (at a 1∶30,000 dilution). The positions of the molecular mass markers (in kDa) are indicated on the left. The two forms of calnexin detected are identified with an asterisk (*). Concomitantly, the OD595 of cells expressing endogenous calnexin was measured at each time point and graphed in correlation with the cleavage of calnexin as assessed by Western blotting with anti-Cnx1p rabbit polyclonal serum. (B) Anti-cmyc immunoblot detection of c-myc-tagged calnexin before and after cleavage. Samples corresponding to 10 µg of protein extracts from cells expressing a cmyc-tagged version of calnexin (Cnx1p-cmyc) at OD595 = 0.3 or OD595 = 7 were loaded on a 15% (wt/vol) SDS-PAGE gel. Calnexin was detected by immunoblotting with the 9E10 anti-cmyc mouse mAb (at a 1∶500 dilution). The positions of the molecular mass markers (in kDa) are indicated on the left. (C) Region of calnexin spanning the site of cleavage. The region of cleavage of calnexin was determined by a series of truncation mutants and tandem mass spectrometry analysis of the small cmyc-tagged fragment of calnexin detected in Figure 6B. In this sequence, the amino acids in bold correspond to the highly conserved central domain (hcd) and the underlined residues correspond to the transmembrane domain (TM). The region delimited in red contains the cleavage site of calnexin. (D) Gel-filtration fractionation of uncleaved (full length) and cleaved calnexin. Samples from cells expressing endogenous calnexin cultured for 24 h were subjected to gel-filtration analysis. Equal volumes of fractions 2 to 10 were loaded on an 8.5% (wt/vol) SDS-PAGE gel and immunoblotted with anti-Cnx1p rabbit polyclonal serum (at a 1∶30,000 dilution) or anti-BiP rabbit polyclonal serum (at a1∶30,000 dilution). Arrows denote the migration of molecular mass standards (thyroglobulin, 669 kDa; β-amylase, 200 kDa; bovine serum albumin, 68 kDa; carbonic anhydrase, 29 kDa). The two forms of calnexin detected are identified with black arrows.

https://doi.org/10.1371/journal.pone.0006244.g006

Discussion

The evidence accumulated in the past ten years demonstrate the existence of apoptotic pathways in yeasts [19], [21], [78], [79], [80], [81], [82], [83], [84], [85], [86], [87], [88]. Although numerous homologues of mammalian apoptosis factors are found in yeast, the key players of specific apoptotic processes are for the most part unknown [21]. And like in mammalian [89], [90], [91], in yeasts the interactions between cells factors within the apoptotic mechanisms are not completely described. The use of yeast models should assist in charting the core interactions of apoptotic pathways conserved in higher eukaryotes.

Recent reports point to a role of calnexin in apoptosis triggered by ER stress in mammalian cells. Calnexin was reported to act as a scaffold for the cleavage by caspase 8 of the apoptotic protein Bap31 under conditions of tunicamycin stress [70]. These authors suggested that the action of calnexin is dependent on its localisation in a specific ER sub-compartment and on its expression level [92]. These studies raised the possibility that calnexin plays a role in the early steps in the transduction of an apoptotic signal initiated by ER stress. In fission yeast, our recent results demonstrate that calnexin is involved in ER-stress induced apoptosis elicited by tunicamycin [72]. We showed that the anchoring of calnexin into the ER membrane is crucial for its apoptotic action under ER stress. The levels of calnexin increase during ER stress triggered by tunicamycin [57]. Consistently, overexpression of calnexin also induces apoptotic death; an experimental intervention probably mimicking conditions of ER stress [72].

Here, we demonstrate that starvation in inositol induces apoptotic cell death in S. pombe as observed by an important reduction in the ability to form colonies, Phloxin B staining, metacaspase activation, DNA breakage and nuclear fragmentation. This apoptotic death is dependent on Pca1p, the only metacaspase so far identified in S. pombe, and is influenced by the extensively-characterized UPR transducer Ire1p.

Inositol is a precursor for numerous molecules playing central roles in membrane integrity, cell signalling and vesicular trafficking. Many organisms have the capacity to synthesize inositol from glucose-6-phosphate by the enzyme inositol-1-phosphate synthase, encoded by the INO1 gene in S. cerevisiae [35], [36]. The S. pombe genome does not encode an Ino1p homologue, thus sensitizing this yeast to inositol deficiency because inositol is an essential precursor [44], [45]. Moreover, S. cerevisiae strains deleted for INO1 die when inositol is absent in the culture media [46], [47], [48], [49], [50], [51]. Numerous screens performed in S. cerevisiae demonstrated that deficiencies in several pathways can lead to inositol auxotrophy, indicating that inositol biosynthesis is linked to diverse cellular processes [40], [41], [93], [94], [95], [96]. S. cerevisiae cells cultured in media depleted of inositol show also an activation of the UPR pathway, which shuts off when inositol is replenished in the media [42], [43], [97].

The apoptotic phenotypes observed following inositol depletion could be due to an imbalance of ER calcium, as in mammals it was shown that the IP3 receptor (IP3R) is a regulator of ER calcium, and calcium is pro-apoptotic in certain situations [98]. IP3R favours Ca2+ release from the ER after binding of its IP3 ligand. Although no homologue of IP3R was found in yeast to date, it was shown that IP3 could trigger the calcium release from vacuoles [34]. The lack of inositol in the culture medium of fission yeast could lower the IP3 concentration in the cytosol leading to inhibition of calcium release from ER or vacuoles. It is known that an increase in the calcium concentration in the cytosol is a condition triggering apoptosis [99]. On the other hand, some studies suggest that a high calcium concentration in the ER protects against apoptosis in particular situations [100], [101]. In this sense, an inhibition of regulated calcium release by low levels of IP3 could imbalance the Ca2+ concentration in the ER and/or the vacuoles eventually triggering apoptosis. Moreover, numerous studies also demonstrate that lowering the inositol concentration or inhibiting IP3R induce autophagy [102], [103], [104], [105]. Autophagy could be a precursor of apoptosis when the condition inducing autophagy is maintained [106]. Thus, apoptosis induced by the lack of inositol could be the endpoint of autophagy.

Low levels of both inositol and IP3, and defects in IP3R signalling have been associated with autophagy in mammals [103], [107], [108]. Moreover, alterations in phospholipid levels in S. cerevisiae correspond to the appearance of autophagy markers [109]. Inasmuch as the induction of autophagy can lead to apoptosis, a possibility is that inositol starvation triggers apoptotic cell death via an autophagic program [110], [111]. Inositol is an essential precursor for a large number of phospholipids, and inositol deficiency deregulates the levels of phospholipids in S. pombe [49]. Studies in fission yeast demonstrated that deregulation in the balance of lipids induces apoptotic death [112], [113], [114]. The same group showed that deregulation of lipid homeostasis could lead to cell death via several pathways dependent or not on different players such as Pca1p, Rad9p or Pkc1p. Moreover, it was shown that valproate, a short-chain fatty acid causing inositol depletion in budding yeast, induces a metacaspase-dependent apoptotic pathway with accumulation of neutral lipids in this yeast [88], [115], [116], [117]. Hence, these observations raise the possibility that inositol deficiency triggers apoptotic death in fission yeast via the loss of lipid homeostasis.

It was shown that INO1 is under the control of Ire1p in S. cerevisiae and that UPR in this yeast is tightly linked to inositol biosynthesis. Therefore, we expected that ire1+ could have a role in the apoptotic pathway induced by inositol starvation in S. pombe, although this fission yeast does not contain a homologue of INO1 [42], [43], [97]. We observed that deletion of ire1+ decreases significantly but not totally the levels of apoptotic death provoked by the absence of inositol. This observation demonstrates that in fission yeast, the consequences of inositol deficiency are partially dependent on the major UPR transducer. The fact that deletion of IRE1 leads to inositol auxotrophy in S. cerevisiae and that deletion of ire1+ in S. pombe reduces the apoptotic effect provoked by inositol starvation suggest that Ire1p could be involved not only in inositol biosynthesis but also in inositol signalling. It is possible that Ire1p acts as an inositol sensor in both yeasts, signalling for inositol biosynthesis in S. cerevisiae and for cell death in S. pombe when cells are depleted of inositol.

Absence of the TM and cytosolic tail of calnexin increases the apoptotic phenotype due the lack of inositol in the media. By co-expression of lumenal_Cnx1p with C-termTM_Cnx1p_cmyc, we observed that the cytosolic tail anchored to the ER membrane reduced to WT levels the apoptotic effect provoked by inositol deprivation. This observation demonstrates the involvement of calnexin in this apoptotic pathway and points to the importance of the cytosolic tail and TM of this ER protein. Remarkably, these results are in contrast with our previous observations regarding the involvement of calnexin in ER-stress mediated apoptotic cell death [72]. In our previous study, we demonstrated that apoptosis triggered by tunicamycin treatment is less efficient in cells expressing only lumenal_Cnx1p, a calnexin mutant not anchored to the ER membrane [72]. In the present work, apoptosis induced by inositol starvation is more efficient when only the lumenal version of calnexin is present in the cell, suggesting that calnexin is involved in at least two different apoptotic pathways: one induced by ER stress and the other by inositol starvation. Although inositol starvation and tunicamycin treatment are two inducers of apoptosis, their mechanisms of action leading to cell death are different. Tunicamycin treatment blocks N-glycosylation, which causes folding defects resulting in severe ER stress [118], [119]. In the case of inositol starvation, while the actual mechanism leading to apoptosis remains undefined, it is known that lack of inositol affects lipid homeostasis and signalling [46], [47], [48], [49], [50], [51], [103], [108]. These significant differences may underlie the two diverging effects observed with the mutant lumenal_Cnx1p in these two apoptotic situations. Another distinction between these two apoptotic situations resides in the dependence or independence on the Pca1p metacaspase. Here we demonstrated apoptosis induced by inositol starvation is dependant on the metacaspase Pca1p, whereas in the case of ER-stress apoptotic cell death is independent of Pca1p [72]. From our studies, it is clear that calnexin is implicated in apoptotic processes, and that the TM and cytosolic tail of calnexin play important regulatory roles in these death pathways. These observations suggest that the TM and cytosolic tail of calnexin form crucial interactions playing pro- or anti-death roles depending on the apoptosis pathway.

In the course of our experiments, we came to the unexpected observation that calnexin undergoes cleavage within its lumenal domain under normal growth conditions. At late exponential phase (OD595 4), calnexin is cleaved into two moieties: a large one spanning most of the lumenal domain and a small one containing a few residues of the C-terminal part of the lumenal domain attached to the TM and the cytosolic tail. This cleavage is consistently 100% efficient under normal culture conditions at the same growth stage, suggesting that this is a physiological processing of calnexin that is important for its cellular roles. In gel-filtration, full-length and processed calnexin elute in different fractions; hinting that they associate with different factors and that each calnexin fragment is involved in different cellular roles. It appears then possible that the small C-terminal moiety of calnexin interacts with other proteins in a complex that negatively regulates apoptosis induced by inositol starvation.

Based on our results obtained with the lumenal_Cnx1p and C-termTM_Cnx1p_cmyc mutants, in Figure 7 we propose a simple model explaining the role of the cleavage of calnexin in apoptosis induced by inositol starvation. Panel A depicts WT calnexin in cells growing in exponential and stationary phase. In the presence or absence of inositol, WT calnexin cells survive until they reach the early stationary phase. In early stationary phase, calnexin undergoes physiological cleavage into two moieties: a large lumenal portion, and a small one containing a few residues of the C-terminal part of the lumenal domain attached to the TM and the cytosolic tail. The small calnexin fragment associates with cell factors (that could be on either side or both sides of the ER membrane) into a complex that delays apoptosis under conditions of inositol starvation. This anti-apoptotic complex would allow autophagy to recycle inositol-containing molecules thus delaying apoptosis. However, under prolonged inositol starvation, cells finally undergo apoptotic death. In the case of cells expressing only the lumenal_Cnx1p mutant (Figure 7B), no such anti-apoptotic complex is formed and apoptosis ensues when they reach stationary phase. In panel C, co-expression of C-termTM_Cnx1p_cmyc with lumenal_Cnx1p mimics the cleavage of calnexin when cells attain stationary phase. C-termTM_Cnx1p_cmyc mediates the assembly of the anti-apoptotic complex. Interestingly, the homologue of calnexin in S. cerevisiae does not possess a cytosolic tail and Δino1 cells, which are unable to synthesize inositol, rapidly die in the absence of this molecule [49].

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Figure 7. Model for the involvement of calnexin domains in the regulation of apoptosis induced by inositol starvation.

(A) WT calnexin in cells growing in exponential and stationary phase. In the presence or absence of inositol, WT calnexin cells survive until they reach the early stationary phase. In early stationary phase, calnexin undergoes physiological cleavage into two moieties. The small calnexin fragment associates with cell factors into a complex that delays apoptosis under conditions of inositol starvation. (B) Cells expressing only the lumenal_Cnx1p mutant. The formation of an anti-apoptotic complex is not possible and apoptosis ensues when cells reach stationary phase. (C) Co-expression of C-termTM_Cnx1p_cmyc with lumenal_Cnx1p. The co-expression of C-termTM_Cnx1p_cmyc mimics the cleavage of calnexin when cells attain stationary phase. C-termTM_Cnx1p_cmyc mediates the assembly of the anti-apoptotic complex thus delaying apoptosis.

https://doi.org/10.1371/journal.pone.0006244.g007

Because the absence of cytosolic tail and TM reduced by 50% the apoptotic death induced by tunicamycin [72], in contrast to what we observed for inositol starvation, we propose that the cytosolic tail of calnexin and its TM have different regulatory roles in different apoptotic pathways, depending of the origin of the apoptotic signal. The apoptotic signal induced by tunicamycin treatment probably originates from the ER lumen, as this drug provokes folding defects within this organelle space. Whereas in the case of inositol starvation the apoptotic signal could originate from the cytosol or the ER membrane, as inositol is an essential precursor of molecules involved in signalling taking place in the cytosol and in the synthesis of membrane phospholipids. The difference observed for the dependency to the metacaspase in apoptosis induced by an N-glycosylation defect or an inositol depletion in S. cerevisiae and S. pombe is another indication of the differences in the two apoptotic pathways induced by these two particular conditions [72], [88], [118]. Taken together, these observations clearly indicate that calnexin is a partner in at least two apoptotic cell-death pathways, triggered by two different inducers. Further studies are needed to understand more precisely the mechanistic details of the involvement of calnexin in these apoptotic processes and to unveil whether calnexin takes part in other apoptotic pathways.

Calnexin was first identified as an ER phosphoprotein that binds calcium [120]. Soon after it was defined as glycoprotein-specific molecular chaperone binding its client proteins through lectin-glycan interactions [121], [122], [123], and later calnexin was shown to interact with non-glycosylated folding polypeptides via peptide-peptide contacts [58], [124], [125], [126]. Also from early reports, the role of calnexin in the quality control of protein folding was demonstrated [127]. Phosphorylation of the cytosolic tail of calnexin was shown to regulate its association with ribosomes and in this manner to affect translocation into the ER [128], [129]. More recently, calnexin was implicated in apoptosis induced by ER stress [70], [72], [92], and here we demonstrated its involvement in the regulation of another apoptotic pathway which is triggered by inositol deprivation. Clearly, calnexin is a multifunctional protein involved in several cellular processes. The use of the S. pombe model will continue to be instrumental in the elucidation of novel and unsuspected cellular roles of calnexin.

Materials and Methods

Yeast strains, media, and vectors

Experiments were carried out using the S. pombe strains described in Table 1. All strains were cultured at 30°C in Edinburgh minimal medium (here denoted as MM) supplemented with required supplements [130]. The nmt promoter in the vectors pREP41 and pREP42 is medium strength and expressed calnexin at levels comparable to the endogenous genomic copy of the gene [125]. pREP41 differs from pREP42 in that it contains the S. pombe LEU2 marker instead of ura4+. The wild-type calnexin and the viable mutant lumenal_cnx1 on pREP41 vector were used to study the effects of inositol starvation (strains SP3234-8 and SP3235-9). The unviable mutant C-termTM_cnx1_cmyc on pREP42 was also used to study the effects of inositol starvation, but in co-expression with the mutant lumenal_cnx1 on pREP41 (strain SP8244).

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Table 1. Yeast strains used for this study.

https://doi.org/10.1371/journal.pone.0006244.t001

Starvation Assay

Cells were cultured for 43 h until saturation in MM containing the required supplements. To obtain exponentially growing cells, the cells were diluted and cultured overnight until they reached OD595 = 1.0. A volume of 10 ml of culture was centrifuged and the cells washed once with MM without inositol or leucine or adenine and resuspended in MM without inositol or leucine or adenine. The tests were performed at the indicated times of starvation.

Time-course cleavage of calnexin

To study the physiological cleavage of calnexin, cells were taken at different time-points of the growth curve. Time 0 corresponds to an OD595 equivalent to 0.1 resulting from an original dilution from exponentially growing cells. Following time 0, a sample of cells was taken each 12 h to perform protein extraction and the OD corresponding to this time-point was measured. More cells were taken at time 0 in order to visualize proteins. Protein extracts were prepared as previously described in an immunoprecipitation buffer (50 mM HEPES pH 7, 50 mM NaCl, 1 mM CaCl2, 1% Nonidet P40) containing 10 mM iodoacetamide, 1 mM PMSF and 1× protease inhibitors (peptadine 1 µg/ml, leupepdine 1 µg/ml, phenathroline 1 mg/ml) [65]. Protein extracts were separated on 10% (w/v) SDS-PAGE gels. Proteins were transferred onto nitrocellulose membrane according to the manufacturer's instructions. Immunoblotting to detect Cnx1p was carried out with an anti-Cnx1p rabbit polyclonal antibody (LAR223), at a dilution of 1∶30,000.

Quantification of luminal_Cnx1p levels

An equivalent of 10 ml of cells at OD595 = 1 from strains SP3235-9 and SP8244 were taken. Protein extracts were prepared as previously described in an immunoprecipitation buffer (50 mM HEPES pH 7, 50 mM NaCl, 1 mM CaCl2, 1% Nonidet P40) containing 10 mM iodoacetamide, 1 mM PMSF and 1× protease inhibitors (peptadine 1 µg/ml, leupepdine 1 µg/ml, phenathroline 1 mg/ml) [65]. Protein extracts were separated on 10% (w/v) SDS-PAGE gels. Proteins were transferred onto nitrocellulose membrane according to the manufacturer's instructions and colored with Ponceau red (Sigma). Immunoblotting of Cnx1p was carried out with an anti-Cnx1p rabbit polyclonal antibody (LAR223, dilution 1∶20,000) and immunoblotting of tubulin was carried out with rabbit polyclonal anti-human tubulin antibodies (Santa Cruz Biotechnologies, dilution 1∶1000). Band quantification was performed with the Quantity One software (Biorad).

Gel-filtration chromatography

Protein extractions were performed as previously described in an immunoprecipitation buffer (50 mM HEPES pH 7, 50 mM NaCl, 1 mM CaCl2, 1% Nonidet P40) containing 10 mM iodoacetamide, 1 mM PMSF and 1× protease inhibitors (peptadine 1 µg/ml, leupepdine 1 µg/ml, phenathroline 1 mg/ml) [65]. A Sephacryl S-300 (Pharmacia Biotech Inc.) column (0.8 by 60 cm) was equilibrated in gel-filtration buffer (50 mM Tris-HCl, 200 mM NaCl, 10 mM MgCl2, 1 mM EDTA pH 7.5, 0.1% Triton X-100, 5% glycerol) at room temperature and calibrated with molecular weight standards (thyroglobulin, 669 kDa; β-amylase, 200 kDa; bovine serum albumin, 68 kDa; carbonic anhydrase, 29 kDa). The void volume was calculated as 12.5–13 ml. The sample was loaded on the column and eluted with the gel-filtration buffer. After 12.5–13 ml, 30 fractions of 1 ml were collected and an equal volume of each fraction was loaded on an 8.5% SDS-PAGE gel. Cnx1p and BiP were detected in each fraction by immunoblotting with the corresponding antibodies.

Mass spectrometry analysis

Immunoprecipitation was performed as previously described using cells expressing a C-terminal cmyc-tagged version of Cnx1p cultured for 48 hours [125]. Anti-cmyc mouse mAb 9E10 (1∶100 dilution) was used to perform immunoprecipitations. Immunoprecipitates were loaded and fractionated on a 15% SDS-PAGE gel, and the gel was stained with Coomassie blue. The band corresponding to the cmyc-tagged Cnx1p fragment was cut-out of the gel and analyzed by MS/MS by the Proteomics Core facility of the Institute for Research in Immunology and Cancer (IRIC), at Université de Montréal. The fragment was subjected to tryptic digestion and analyzed by nanoliquid chromatography/tandem mass spectrometry.

Viability Assays

The survival of cells was measured by two different techniques: 1) the ability to form colonies by serial 10-fold dilutions spotted on appropriate plates; and 2) by cytometry with the vital fluorescent dye Phloxin B. For serial dilutions spotting experiments, an equivalent of OD595 = 1.0 was taken from cells starved in inositol for 48 h. The cells were serially diluted (10−1–10−4), spotted on solid media and incubated for 7 days at 30°C. Viability assays with the Phloxin B fluorescent vital dye was carried out as previously described after 18 h of starvation [22].

Calcofluor staining

Samples containing 1.4×107 cells were taken after 48 h of inositol starvation. Cells were washed once in 1× PBS pH 7.4, fixed for 10 min in a solution of 3.7% formaldehyde and washed once in 1× PBS pH 7.4. The cells were resuspended in 100 µl 1× PBS pH 7.4 containing 20 µg/ml Fluorescent Brightener 28 (Sigma Inc.) for 5 min. and washed once in 1× PBS pH 7.4. Finally the cells were resuspended in 1× PBS pH 7.4 to a final concentration of 5×107–1×108 cells/ml. Suitable quantities of cells were applied to a poly-lysine coated coverslips, washed and let dry. The slides were mounted with a mounting media (1 mg/mL p-phenylenediamine, 90% glycerol). Microscopy analysis was performed using a fluorescence inverted microscope Nikon TE2000U. Images were acquired using a motion-picture camera CCD coolSnapFX M® 12 bit and treated with the UIC Metamorph® software.

Detection of Apoptotic Markers

Metacaspase Activation.

Culture samples were taken after 18 hours of inositol starvation. Aliquots containing 1×107 cells were washed once in 1 ml of 1× PBS pH 7.4 (136 mM NaCl, 25 mM KCl, 12 mM NaHPO4, 18 mM KH2PO4) and resuspended in 150 µl of 1× PBS pH 7.4 containing 10 µM FITC-VAD-fmk (CasPACE, Promega, Madison, WI, USA). After incubation for 20 min at 30°C, cells were washed once in 1× PBS pH 7.4 and resuspended in 100 µl 1× PBS pH 7.4 to be analysed by flow cytometry.

DAPI Staining.

For DAPI (4′,6-diamidino-2-phenylindole) staining, samples containing 1.4×107 cells were taken after 48 h of inositol starvation. Cells were fixed for 10 min in a solution of 3.7% formaldehyde, washed once in 1× PBS pH 7.4 containing 1% Nonidet P-40, and twice in 1× PBS pH 7.4. The cells were resuspended in 100 µl 1× PBS pH 7.4 to a final concentration of 5×107–1×108 cells/ml. Suitable quantities of cells were applied to a poly-lysine coated coverslips, washed and let dry. The slides were mounted with a DAPI-containing mounting media (1 µg/mL DAPI, 1 mg/mL p-phenylenediamine, 90% glycerol). Microscopy analysis was performed using a fluorescence inverted microscope Nikon TE2000U. Images were acquired using a motion-picture camera CCD coolSnapFX M® 12 bit and treated with the UIC Metamorph® software.

TUNEL assay.

TUNEL (Terminal uridine deoxynucleotidyl transferase dUTP nick end labeling) assay was performed with the APO-BRDU TUNEL Kit (PHOENIX flow systems, San Diego, CA), essentially following the manufacturer's recommendations. Following 48 hours of inositol starvation by culturing cells in media without inositol, 1.4×107 cells were taken and fixed with 1 ml of 3.7% formaldehyde. After fixation, the cell wall was digested by resuspending the cell pellet in 200 µl of sorbitol buffer (1.2 M sorbitol, 0.5 mM MgCl2, potassium phosphate, pH 6.8) containing 5 mg/ml of lysing enzymes (SIGMA), and incubating for 90 min at room temperature, followed by 30 minutes incubation at 37°C. The cell pellet was resuspended in 500 µl of permeabilization solution (0.1% Triton in 0.1% sodium citrate) and let on ice for 2 minutes, washed twice with 400 µl of WASH solution and incubated in 50 µl of TUNEL solution for 30 min at 30°C. After incubation, the cells were washed twice in WASH solution and incubated 30 min at room temperature in the dark with 100 µl antibodies solution (anti-BrdU antibodies). Staining of the cells was analyzed by flow cytometry (FACS).

Flow Cytometry Analyses

Cells were stained with Phloxin B or with FITC-VAD-fmk as described above. Flow cytometry analyses were performed using a FACS Calibur (Becton Dickinson Biosciences) device, on 10,000 cells. Emission from the argon LASER was at 488 nm; emission settings were 515–545 nm (filter FL-1) for FITC-VAD-fmk or 560–600 nm (filter FL-2) for Phloxin B staining. The percentage of positive stained cells was determined as the population of fluorescent cells with a higher fluorescent intensity than a stained negative control. Parameters of the stained negative control were adjusted with an unstained negative control. Each experiment was repeated three times.

Statistical Analysis

The significance of the variations of results among strains was determined by a Student's t test.

Supporting Information

Figure S1.

Starvation in adenine or leucine does not induce apoptotic cell death (A) Survival of cells cultured for 12 h and 24 h in MM with or without leucine was assayed by serial dilution on media containing leucine. Samples of 10 µl of four 10-fold serial dilutions (10−1–10−4) of cells at OD595 = 1 were spotted on selective MM with leucine, and incubated at 30°C for 7 days (see Materials and Methods). (B) Survival of cells cultured for 12 h and 24 h in MM with or without adenine was assayed by serial dilution on media containing adenine. Samples of 10 µl of four 10-fold serial dilutions (10−1–10−4) of cells at OD595 = 1 were spotted on selective MM with adenine, and incubated at 30°C for 7 days (see Materials and Methods).

https://doi.org/10.1371/journal.pone.0006244.s001

(0.14 MB TIF)

Figure S2.

Quantification of lumenal_Cnx1p Anti-Cnx1p immunoblot of lumenal_Cnx1p alone (SP3235-9) or co-expressed with C-termTM_Cnx1p_cmyc (SP8244). Samples corresponding to 20 µg of protein extracts at OD595 = 0.5 were loaded onto a 10% (wt/vol) SDS-PAGE gel. Calnexin was detected by immunoblotting with anti-Cnx1p antibodies. Anti-tubulin immnublot and Ponceau-red staining are shown as loading controls. Band quantification was performed with the Quantity One software (Biorad).

https://doi.org/10.1371/journal.pone.0006244.s002

(0.42 MB TIF)

Acknowledgments

We thank all members of the Rokeach lab for fruitful discussions and critical reading of this manuscript.

Author Contributions

Conceived and designed the experiments: RG LAR. Performed the experiments: RG PBB AL. Analyzed the data: RG AL LAR. Contributed reagents/materials/analysis tools: RG. Wrote the paper: RG AL LAR.

References

  1. 1. Bukau B, Deuerling E, Pfund C, Craig EA (2000) Getting newly synthesized proteins into shape. Cell 101: 119–122.
  2. 2. Fewell SW, Travers KJ, Weissman JS, Brodsky JL (2001) The action of molecular chaperones in the early secretory pathway. Annu Rev Genet 35: 149–191.
  3. 3. Trombetta ES, Parodi AJ (2003) Quality control and protein folding in the secretory pathway. Annu Rev Cell Dev Biol 19: 649–676.
  4. 4. Lai E, Teodoro T, Volchuk A (2007) Endoplasmic reticulum stress: signaling the unfolded protein response. Physiology (Bethesda) 22: 193–201.
  5. 5. Boyce M, Yuan J (2006) Cellular response to endoplasmic reticulum stress: a matter of life or death. Cell Death Differ 13: 363–373.
  6. 6. Breckenridge DG, Germain M, Mathai JP, Nguyen M, Shore GC (2003) Regulation of apoptosis by endoplasmic reticulum pathways. Oncogene 22: 8608–8618.
  7. 7. Szegezdi E, Logue SE, Gorman AM, Samali A (2006) Mediators of endoplasmic reticulum stress-induced apoptosis. EMBO Rep 7: 880–885.
  8. 8. Ron D, Walter P (2007) Signal integration in the endoplasmic reticulum unfolded protein response. Nat Rev Mol Cell Biol 8: 519–529.
  9. 9. Patil C, Walter P (2001) Intracellular signaling from the endoplasmic reticulum to the nucleus: the unfolded protein response in yeast and mammals. Curr Opin Cell Biol 13: 349–355.
  10. 10. Schroder M, Kaufman RJ (2005) ER stress and the unfolded protein response. Mutat Res 569: 29–63.
  11. 11. Wu J, Kaufman RJ (2006) From acute ER stress to physiological roles of the Unfolded Protein Response. Cell Death Differ 13: 374–384.
  12. 12. Ahner A, Brodsky JL (2004) Checkpoints in ER-associated degradation: excuse me, which way to the proteasome? Trends Cell Biol 14: 474–478.
  13. 13. Meusser B, Hirsch C, Jarosch E, Sommer T (2005) ERAD: the long road to destruction. Nat Cell Biol 7: 766–772.
  14. 14. Marciniak SJ, Ron D (2006) Endoplasmic reticulum stress signaling in disease. Physiol Rev 86: 1133–1149.
  15. 15. Schroder M, Kaufman RJ (2005) The mammalian unfolded protein response. Annu Rev Biochem 74: 739–789.
  16. 16. Xu C, Bailly-Maitre B, Reed JC (2005) Endoplasmic reticulum stress: cell life and death decisions. J Clin Invest 115: 2656–2664.
  17. 17. Kerr JF (2002) History of the events leading to the formulation of the apoptosis concept. Toxicology 181–182: 471–474.
  18. 18. Kerr JF, Wyllie AH, Currie AR (1972) Apoptosis: a basic biological phenomenon with wide-ranging implications in tissue kinetics. Br J Cancer 26: 239–257.
  19. 19. Severin FF, Meer MV, Smirnova EA, Knorre DA, Skulachev VP (2008) Natural causes of programmed death of yeast Saccharomyces cerevisiae. Biochim Biophys Acta 1783: 1350–1353.
  20. 20. Hamann A, Brust D, Osiewacz HD (2008) Apoptosis pathways in fungal growth, development and ageing. Trends Microbiol 16: 276–283.
  21. 21. Frohlich KU, Fussi H, Ruckenstuhl C (2007) Yeast apoptosis–from genes to pathways. Semin Cancer Biol 17: 112–121.
  22. 22. Roux AE, Quissac A, Chartrand P, Ferbeyre G, Rokeach LA (2006) Regulation of chronological aging in Schizosaccharomyces pombe by the protein kinases Pka1 and Sck2. Aging Cell 5: 345–357.
  23. 23. Wissing S, Ludovico P, Herker E, Buttner S, Engelhardt SM, et al. (2004) An AIF orthologue regulates apoptosis in yeast. J Cell Biol 166: 969–974.
  24. 24. Walter D, Wissing S, Madeo F, Fahrenkrog B (2006) The inhibitor-of-apoptosis protein Bir1p protects against apoptosis in S. cerevisiae and is a substrate for the yeast homologue of Omi/HtrA2. J Cell Sci 119: 1843–1851.
  25. 25. Jin C, Reed JC (2002) Yeast and apoptosis. Nat Rev Mol Cell Biol 3: 453–459.
  26. 26. Fahrenkrog B, Sauder U, Aebi U (2004) The S. cerevisiae HtrA-like protein Nma111p is a nuclear serine protease that mediates yeast apoptosis. J Cell Sci 117: 115–126.
  27. 27. Burhans WC, Weinberger M (2007) Yeast endonuclease G: complex matters of death, and of life. Mol Cell 25: 323–325.
  28. 28. Madeo F, Herker E, Maldener C, Wissing S, Lachelt S, et al. (2002) A caspase-related protease regulates apoptosis in yeast. Mol Cell 9: 911–917.
  29. 29. De Camilli P, Emr SD, McPherson PS, Novick P (1996) Phosphoinositides as regulators in membrane traffic. Science 271: 1533–1539.
  30. 30. Majerus PW, Connolly TM, Deckmyn H, Ross TS, Bross TE, et al. (1986) The metabolism of phosphoinositide-derived messenger molecules. Science 234: 1519–1526.
  31. 31. Steiner S, Lester RL (1972) Studies on the diversity of inositol-containing yeast phospholipids: incorporation of 2-deoxyglucose into lipid. J Bacteriol 109: 81–88.
  32. 32. Berridge MJ (1987) Inositol trisphosphate and diacylglycerol: two interacting second messengers. Annu Rev Biochem 56: 159–193.
  33. 33. Berridge MJ, Bootman MD, Roderick HL (2003) Calcium signalling: dynamics, homeostasis and remodelling. Nat Rev Mol Cell Biol 4: 517–529.
  34. 34. Belde PJ, Vossen JH, Borst-Pauwels GW, Theuvenet AP (1993) Inositol 1,4,5-trisphosphate releases Ca2+ from vacuolar membrane vesicles of Saccharomyces cerevisiae. FEBS Lett 323: 113–118.
  35. 35. Donahue TF, Henry SA (1981) myo-Inositol-1-phosphate synthase. Characteristics of the enzyme and identification of its structural gene in yeast. J Biol Chem 256: 7077–7085.
  36. 36. Eisenberg F, Bolden AH, Loewus FA (1964) Inositol formation by cyclization of glucose chain in rat testis. Biochem Biophys Res Commun 14: 419–424.
  37. 37. Klig LS, Antonsson B, Schmid E, Friedli L (1991) Inositol biosynthesis: Candida albicans and Saccharomyces cerevisiae genes share common regulation. Yeast 7: 325–336.
  38. 38. Klig LS, Henry SA (1984) Isolation of the yeast INO1 gene: located on an autonomously replicating plasmid, the gene is fully regulated. Proc Natl Acad Sci U S A 81: 3816–3820.
  39. 39. Majumder AL, Johnson MD, Henry SA (1997) 1L-myo-inositol-1-phosphate synthase. Biochim Biophys Acta 1348: 245–256.
  40. 40. Dean-Johnson M, Henry SA (1989) Biosynthesis of inositol in yeast. Primary structure of myo-inositol-1-phosphate synthase (EC 5.5.1.4) and functional analysis of its structural gene, the INO1 locus. J Biol Chem 264: 1274–1283.
  41. 41. Greenberg ML, Lopes JM (1996) Genetic regulation of phospholipid biosynthesis in Saccharomyces cerevisiae. Microbiol Rev 60: 1–20.
  42. 42. Chang HJ, Jones EW, Henry SA (2002) Role of the unfolded protein response pathway in regulation of INO1 and in the sec14 bypass mechanism in Saccharomyces cerevisiae. Genetics 162: 29–43.
  43. 43. Cox JS, Chapman RE, Walter P (1997) The unfolded protein response coordinates the production of endoplasmic reticulum protein and endoplasmic reticulum membrane. Mol Biol Cell 8: 1805–1814.
  44. 44. MacNeill SA (2002) Genome sequencing: and then there were six. Curr Biol 12: R294–296.
  45. 45. Yarbrough HF Jr, Clark FM (1957) Utilization of inositol, an essential metabolite for Schizosaccharomyces pombe. J Bacteriol 73: 318–323.
  46. 46. Niederberger C, Graub R, Schweingruber AM, Fankhauser H, Rusu M, et al. (1998) Exogenous inositol and genes responsible for inositol transport are required for mating and sporulation in Shizosaccharomyces pombe. Curr Genet 33: 255–261.
  47. 47. Poitelea M, Rusu M (1997) [Inositol is necessary to sexual differentiation in Schizosaccharomyces pombe]. Rev Med Chir Soc Med Nat Iasi 101: 87–91.
  48. 48. Voicu PM, Poitelea M, Schweingruber E, Rusu M (2002) Inositol is specifically involved in the sexual program of the fission yeast Schizosaccharomyces pombe. Arch Microbiol 177: 251–258.
  49. 49. Fernandez S, Homann MJ, Henry SA, Carman GM (1986) Metabolism of the phospholipid precursor inositol and its relationship to growth and viability in the natural auxotroph Schizosaccharomyces pombe. J Bacteriol 166: 779–786.
  50. 50. Minskoff SA, Gaynor PM, Greenberg ML (1992) Mutant enrichment of Schizosaccharomyces pombe by inositol-less death. J Bacteriol 174: 4078–4085.
  51. 51. Ingavale SS, Bachhawat AK (1999) Restoration of inositol prototrophy in the fission yeast Schizosaccharomyces pombe. Microbiology 145 ( Pt 8): 1903–1910.
  52. 52. Williams DB (2006) Beyond lectins: the calnexin/calreticulin chaperone system of the endoplasmic reticulum. J Cell Sci 119: 615–623.
  53. 53. Arunachalam B, Cresswell P (1995) Molecular requirements for the interaction of class II major histocompatibility complex molecules and invariant chain with calnexin. J Biol Chem 270: 2784–2790.
  54. 54. Beaulieu H, Elagoz A, Crine P, Rokeach LA (1999) Interaction of mammalian neprilysin with binding protein and calnexin in Schizosaccharomyces pombe. Biochem J 340 ( Pt 3): 813–819.
  55. 55. Fernandez F, Jannatipour M, Hellman U, Rokeach LA, Parodi AJ (1996) A new stress protein: synthesis of Schizosaccharomyces pombe UDP–Glc:glycoprotein glucosyltransferase mRNA is induced by stress conditions but the enzyme is not essential for cell viability. Embo J 15: 705–713.
  56. 56. Hebert DN, Garman SC, Molinari M (2005) The glycan code of the endoplasmic reticulum: asparagine-linked carbohydrates as protein maturation and quality-control tags. Trends Cell Biol 15: 364–370.
  57. 57. Jannatipour M, Rokeach LA (1998) A Schizosaccharomyces pombe gene encoding a novel polypeptide with a predicted alpha-helical rod structure found in the myosin and intermediate-filament families of proteins. Biochim Biophys Acta 1399: 67–72.
  58. 58. Marechal A, Tanguay PL, Callejo M, Guerin R, Boileau G, et al. (2004) Cell viability and secretion of active proteins in Schizosaccharomyces pombe do not require the chaperone function of calnexin. Biochem J 380: 441–448.
  59. 59. Parodi AJ (2000) Protein glucosylation and its role in protein folding. Annu Rev Biochem 69: 69–93.
  60. 60. Saito Y, Ihara Y, Leach MR, Cohen-Doyle MF, Williams DB (1999) Calreticulin functions in vitro as a molecular chaperone for both glycosylated and non-glycosylated proteins. Embo J 18: 6718–6729.
  61. 61. Thammavongsa V, Mancino L, Raghavan M (2005) Polypeptide substrate recognition by calnexin requires specific conformations of the calnexin protein. J Biol Chem 280: 33497–33505.
  62. 62. Denzel A, Molinari M, Trigueros C, Martin JE, Velmurgan S, et al. (2002) Early postnatal death and motor disorders in mice congenitally deficient in calnexin expression. Mol Cell Biol 22: 7398–7404.
  63. 63. Jannatipour M, Rokeach LA (1995) The Schizosaccharomyces pombe homologue of the chaperone calnexin is essential for viability. J Biol Chem 270: 4845–4853.
  64. 64. Parlati F, Dignard D, Bergeron JJ, Thomas DY (1995) The calnexin homologue cnx1+ in Schizosaccharomyces pombe, is an essential gene which can be complemented by its soluble ER domain. EMBO J 14: 3064–3072.
  65. 65. Elagoz A, Callejo M, Armstrong J, Rokeach LA (1999) Although calnexin is essential in S. pombe, its highly conserved central domain is dispensable for viability. J Cell Sci 112 ( Pt 23): 4449–4460.
  66. 66. Hajjar F, Beauregard PB, Rokeach LA (2007) The 160 N-terminal residues of calnexin define a novel region supporting viability in Schizosaccharomyces pombe. Yeast 24: 89–103.
  67. 67. Torgler CN, de Tiani M, Raven T, Aubry JP, Brown R, et al. (1997) Expression of bak in S. pombe results in a lethality mediated through interaction with the calnexin homologue Cnx1. Cell Death Differ 4: 263–271.
  68. 68. Groenendyk J, Zuppini A, Shore G, Opas M, Bleackley RC, et al. (2006) Caspase 12 in calnexin-deficient cells. Biochemistry 45: 13219–13226.
  69. 69. Zuppini A, Groenendyk J, Cormack LA, Shore G, Opas M, et al. (2002) Calnexin deficiency and endoplasmic reticulum stress-induced apoptosis. Biochemistry 41: 2850–2858.
  70. 70. Delom F, Emadali A, Cocolakis E, Lebrun JJ, Nantel A, et al. (2006) Calnexin-dependent regulation of tunicamycin-induced apoptosis in breast carcinoma MCF-7 cells. Cell Death Differ.
  71. 71. Takizawa T, Tatematsu C, Watanabe K, Kato K, Nakanishi Y (2004) Cleavage of calnexin caused by apoptotic stimuli: implication for the regulation of apoptosis. J Biochem (Tokyo) 136: 399–405.
  72. 72. Guerin R, Arseneault G, Dumont S, Rokeach LA (2008) Calnexin is involved in apoptosis induced by endoplasmic reticulum stress in the fission yeast. Mol Biol Cell 19: 4404–4420.
  73. 73. Lim HW, Kim SJ, Park EH, Lim CJ (2007) Overexpression of a metacaspase gene stimulates cell growth and stress response in Schizosaccharomyces pombe. Can J Microbiol 53: 1016–1023.
  74. 74. Uren AG, O'Rourke K, Aravind LA, Pisabarro MT, Seshagiri S, et al. (2000) Identification of paracaspases and metacaspases: two ancient families of caspase-like proteins, one of which plays a key role in MALT lymphoma. Mol Cell 6: 961–967.
  75. 75. Elorza MV, Rico H, Sentandreu R (1983) Calcofluor white alters the assembly of chitin fibrils in Saccharomyces cerevisiae and Candida albicans cells. J Gen Microbiol 129: 1577–1582.
  76. 76. Shahinian S, Dijkgraaf GJ, Sdicu AM, Thomas DY, Jakob CA, et al. (1998) Involvement of protein N-glycosyl chain glucosylation and processing in the biosynthesis of cell wall beta-1,6-glucan of Saccharomyces cerevisiae. Genetics 149: 843–856.
  77. 77. Turcotte C, Roux A, Beauregard PB, Guerin R, Senechal P, et al. (2007) The calnexin-independent state does not compensate for all calnexin functions in Schizosaccharomyces pombe. FEMS Yeast Res 7: 196–208.
  78. 78. Buttner S, Eisenberg T, Herker E, Carmona-Gutierrez D, Kroemer G, et al. (2006) Why yeast cells can undergo apoptosis: death in times of peace, love, and war. J Cell Biol 175: 521–525.
  79. 79. Knorre DA, Smirnova EA, Severin FF (2005) Natural conditions inducing programmed cell death in the yeast Saccharomyces cerevisiae. Biochemistry (Mosc) 70: 264–266.
  80. 80. Balzan R, Sapienza K, Galea DR, Vassallo N, Frey H, et al. (2004) Aspirin commits yeast cells to apoptosis depending on carbon source. Microbiology 150: 109–115.
  81. 81. King DA, Hannum DM, Qi JS, Hurst JK (2004) HOCl-mediated cell death and metabolic dysfunction in the yeast Saccharomyces cerevisiae. Arch Biochem Biophys 423: 170–181.
  82. 82. Silva RD, Sotoca R, Johansson B, Ludovico P, Sansonetty F, et al. (2005) Hyperosmotic stress induces metacaspase- and mitochondria-dependent apoptosis in Saccharomyces cerevisiae. Mol Microbiol 58: 824–834.
  83. 83. Madeo F, Frohlich E, Ligr M, Grey M, Sigrist SJ, et al. (1999) Oxygen stress: a regulator of apoptosis in yeast. J Cell Biol 145: 757–767.
  84. 84. Pozniakovsky AI, Knorre DA, Markova OV, Hyman AA, Skulachev VP, et al. (2005) Role of mitochondria in the pheromone- and amiodarone-induced programmed death of yeast. J Cell Biol 168: 257–269.
  85. 85. Ludovico P, Sousa MJ, Silva MT, Leao C, Corte-Real M (2001) Saccharomyces cerevisiae commits to a programmed cell death process in response to acetic acid. Microbiology 147: 2409–2415.
  86. 86. Huh GH, Damsz B, Matsumoto TK, Reddy MP, Rus AM, et al. (2002) Salt causes ion disequilibrium-induced programmed cell death in yeast and plants. Plant J 29: 649–659.
  87. 87. Granot D, Levine A, Dor-Hefetz E (2003) Sugar-induced apoptosis in yeast cells. FEMS Yeast Res 4: 7–13.
  88. 88. Mitsui K, Nakagawa D, Nakamura M, Okamoto T, Tsurugi K (2005) Valproic acid induces apoptosis dependent of Yca1p at concentrations that mildly affect the proliferation of yeast. FEBS Lett 579: 723–727.
  89. 89. Harada H, Grant S (2003) Apoptosis regulators. Rev Clin Exp Hematol 7: 117–138.
  90. 90. Chowdhury I, Tharakan B, Bhat GK (2006) Current concepts in apoptosis: the physiological suicide program revisited. Cell Mol Biol Lett 11: 506–525.
  91. 91. Carmody RJ, Cotter TG (2001) Signalling apoptosis: a radical approach. Redox Rep 6: 77–90.
  92. 92. Delom F, Fessart D, Chevet E (2007) Regulation of calnexin sub-cellular localization modulates endoplasmic reticulum stress-induced apoptosis in MCF-7 cells. Apoptosis.
  93. 93. Chang Q, Petrash JM (2008) Disruption of aldo-keto reductase genes leads to elevated markers of oxidative stress and inositol auxotrophy in Saccharomyces cerevisiae. Biochim Biophys Acta 1783: 237–245.
  94. 94. Hirsch JP, Henry SA (1986) Expression of the Saccharomyces cerevisiae inositol-1-phosphate synthase (INO1) gene is regulated by factors that affect phospholipid synthesis. Mol Cell Biol 6: 3320–3328.
  95. 95. Jesch SA, Zhao X, Wells MT, Henry SA (2005) Genome-wide analysis reveals inositol, not choline, as the major effector of Ino2p-Ino4p and unfolded protein response target gene expression in yeast. J Biol Chem 280: 9106–9118.
  96. 96. Nunez LR, Jesch SA, Gaspar ML, Almaguer C, Villa-Garcia M, et al. (2008) Cell wall integrity MAPK pathway is essential for lipid homeostasis. J Biol Chem 283: 34204–34217.
  97. 97. Nikawa J, Yamashita S (1992) IRE1 encodes a putative protein kinase containing a membrane-spanning domain and is required for inositol phototrophy in Saccharomyces cerevisiae. Mol Microbiol 6: 1441–1446.
  98. 98. Joseph SK, Hajnoczky G (2007) IP3 receptors in cell survival and apoptosis: Ca2+ release and beyond. Apoptosis 12: 951–968.
  99. 99. Pinton P, Giorgi C, Siviero R, Zecchini E, Rizzuto R (2008) Calcium and apoptosis: ER-mitochondria Ca2+ transfer in the control of apoptosis. Oncogene 27: 6407–6418.
  100. 100. Rong Y, Distelhorst CW (2008) Bcl-2 protein family members: versatile regulators of calcium signaling in cell survival and apoptosis. Annu Rev Physiol 70: 73–91.
  101. 101. White C, Li C, Yang J, Petrenko NB, Madesh M, et al. (2005) The endoplasmic reticulum gateway to apoptosis by Bcl-X(L) modulation of the InsP3R. Nat Cell Biol 7: 1021–1028.
  102. 102. Criollo A, Vicencio JM, Tasdemir E, Maiuri MC, Lavandero S, et al. (2007) The inositol trisphosphate receptor in the control of autophagy. Autophagy 3: 350–353.
  103. 103. Sarkar S, Floto RA, Berger Z, Imarisio S, Cordenier A, et al. (2005) Lithium induces autophagy by inhibiting inositol monophosphatase. J Cell Biol 170: 1101–1111.
  104. 104. Sarkar S, Rubinsztein DC (2008) Small molecule enhancers of autophagy for neurodegenerative diseases. Mol Biosyst 4: 895–901.
  105. 105. Criollo A, Maiuri MC, Tasdemir E, Vitale I, Fiebig AA, et al. (2007) Regulation of autophagy by the inositol trisphosphate receptor. Cell Death Differ 14: 1029–1039.
  106. 106. Scarlatti F, Granata R, Meijer AJ, Codogno P (2009) Does autophagy have a license to kill mammalian cells? Cell Death Differ 16: 12–20.
  107. 107. Lam D, Golstein P (2008) A specific pathway inducing autophagic cell death is marked by an IP3R mutation. Autophagy 4: 349–350.
  108. 108. Sarkar S, Rubinsztein DC (2006) Inositol and IP3 levels regulate autophagy: biology and therapeutic speculations. Autophagy 2: 132–134.
  109. 109. Iwanyshyn WM, Han GS, Carman GM (2004) Regulation of phospholipid synthesis in Saccharomyces cerevisiae by zinc. J Biol Chem 279: 21976–21983.
  110. 110. Galluzzi L, Joza N, Tasdemir E, Maiuri MC, Hengartner M, et al. (2008) No death without life: vital functions of apoptotic effectors. Cell Death Differ 15: 1113–1123.
  111. 111. Sinha S, Colbert CL, Becker N, Wei Y, Levine B (2008) Molecular basis of the regulation of Beclin 1-dependent autophagy by the gamma-herpesvirus 68 Bcl-2 homolog M11. Autophagy 4: 989–997.
  112. 112. Zhang Q, Chieu HK, Low CP, Zhang S, Heng CK, et al. (2003) Schizosaccharomyces pombe cells deficient in triacylglycerols synthesis undergo apoptosis upon entry into the stationary phase. J Biol Chem 278: 47145–47155.
  113. 113. Low CP, Liew LP, Pervaiz S, Yang H (2005) Apoptosis and lipoapoptosis in the fission yeast Schizosaccharomyces pombe. FEMS Yeast Res 5: 1199–1206.
  114. 114. Low CP, Shui G, Liew LP, Buttner S, Madeo F, et al. (2008) Caspase-dependent and -independent lipotoxic cell-death pathways in fission yeast. J Cell Sci 121: 2671–2684.
  115. 115. Shaltiel G, Shamir A, Shapiro J, Ding D, Dalton E, et al. (2004) Valproate decreases inositol biosynthesis. Biol Psychiatry 56: 868–874.
  116. 116. Sun Q, Bi L, Su X, Tsurugi K, Mitsui K (2007) Valproate induces apoptosis by inducing accumulation of neutral lipids which was prevented by disruption of the SIR2 gene in Saccharomyces cerevisiae. FEBS Lett 581: 3991–3995.
  117. 117. Vaden DL, Ding D, Peterson B, Greenberg ML (2001) Lithium and valproate decrease inositol mass and increase expression of the yeast INO1 and INO2 genes for inositol biosynthesis. J Biol Chem 276: 15466–15471.
  118. 118. Hauptmann P, Riel C, Kunz-Schughart LA, Frohlich KU, Madeo F, et al. (2006) Defects in N-glycosylation induce apoptosis in yeast. Mol Microbiol 59: 765–778.
  119. 119. Perez-Sala D, Mollinedo F (1995) Inhibition of N-linked glycosylation induces early apoptosis in human promyelocytic HL-60 cells. J Cell Physiol 163: 523–531.
  120. 120. Wada I, Rindress D, Cameron PH, Ou WJ, Doherty JJ 2nd, et al. (1991) SSR alpha and associated calnexin are major calcium binding proteins of the endoplasmic reticulum membrane. J Biol Chem 266: 19599–19610.
  121. 121. Hammond C, Braakman I, Helenius A (1994) Role of N-linked oligosaccharide recognition, glucose trimming, and calnexin in glycoprotein folding and quality control. Proc Natl Acad Sci U S A 91: 913–917.
  122. 122. Hebert DN, Foellmer B, Helenius A (1995) Glucose trimming and reglucosylation determine glycoprotein association with calnexin in the endoplasmic reticulum. Cell 81: 425–433.
  123. 123. Kapoor M, Srinivas H, Kandiah E, Gemma E, Ellgaard L, et al. (2003) Interactions of substrate with calreticulin, an endoplasmic reticulum chaperone. J Biol Chem 278: 6194–6200.
  124. 124. Brockmeier A, Williams DB (2006) Potent lectin-independent chaperone function of calnexin under conditions prevalent within the lumen of the endoplasmic reticulum. Biochemistry 45: 12906–12916.
  125. 125. Jannatipour M, Callejo M, Parodi AJ, Armstrong J, Rokeach LA (1998) Calnexin and BiP interact with acid phosphatase independently of glucose trimming and reglucosylation in Schizosaccharomyces pombe. Biochemistry 37: 17253–17261.
  126. 126. Leach MR, Williams DB (2004) Lectin-deficient calnexin is capable of binding class I histocompatibility molecules in vivo and preventing their degradation. J Biol Chem 279: 9072–9079.
  127. 127. Ellgaard L, Helenius A (2003) Quality control in the endoplasmic reticulum. Nat Rev Mol Cell Biol 4: 181–191.
  128. 128. Chevet E, Wong HN, Gerber D, Cochet C, Fazel A, et al. (1999) Phosphorylation by CK2 and MAPK enhances calnexin association with ribosomes. EMBO J 18: 3655–3666.
  129. 129. Delom F, Chevet E (2006) In vitro mapping of calnexin interaction with ribosomes. Biochem Biophys Res Commun 341: 39–44.
  130. 130. Moreno S, Klar A, Nurse P (1991) Molecular genetic analysis of fission yeast Schizosaccharomyces pombe. Methods Enzymol 194: 795–823.