Lipid-Induced ER Stress: Synergistic Effects of Sterols and Saturated Fatty Acids


  • Ludovic Pineau,

    1. Université de POITIERS, FRE CNRS 3091 “Physiologie moléculaire du Transport des Sucres ”, 40 avenue du Recteur Pineau, 86022 Poitiers, France
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  • Jenny Colas,

    1. Université de POITIERS, FRE CNRS 3091 “Physiologie moléculaire du Transport des Sucres ”, 40 avenue du Recteur Pineau, 86022 Poitiers, France
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  • Sébastien Dupont,

    1. Laboratoire de Génie des Procédés Microbiologiques et Alimentaires, ENSBANA, 1, Esplanade Érasme, Domaine Universitaire, 21000 Dijon, France
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  • Laurent Beney,

    1. Laboratoire de Génie des Procédés Microbiologiques et Alimentaires, ENSBANA, 1, Esplanade Érasme, Domaine Universitaire, 21000 Dijon, France
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  • Pierrette Fleurat-Lessard,

    1. Université de POITIERS, FRE CNRS 3091 “Physiologie moléculaire du Transport des Sucres ”, 40 avenue du Recteur Pineau, 86022 Poitiers, France
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  • Jean-Marc Berjeaud,

    1. UMR CNRS 6008 “Microbiologie fondamentale et appliquée”, 40 avenue du Recteur Pineau, 86022 Poitiers, France
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  • Thierry Bergès,

    1. Université de POITIERS, FRE CNRS 3091 “Physiologie moléculaire du Transport des Sucres ”, 40 avenue du Recteur Pineau, 86022 Poitiers, France
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  • Thierry Ferreira

    Corresponding author
    1. Université de POITIERS, FRE CNRS 3091 “Physiologie moléculaire du Transport des Sucres ”, 40 avenue du Recteur Pineau, 86022 Poitiers, France
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Thierry Ferreira,


Stress within the endoplasmic reticulum (ER) induces a coordinated response, namely the unfolded protein response (UPR), devoted to helping the ER cope with the accumulation of misfolded proteins. Failure of the UPR plays an important role in several human diseases. Recent studies report that intracellular accumulation of saturated fatty acids (SFAs) and cholesterol, seen in diseases of high incidence, such as obesity or atherosclerosis, results in ER stress. In the present study, we evaluated the effects of perturbations to lipid homeostasis on ER stress/UPR induction in the model eukaryote Saccharomyces cerevisiae. We show that SFA originating from either endogenous (preclusion of fatty acid desaturation) or exogenous (feeding with extracellular SFA) sources trigger ER stress and that ergosterol, the major sterol in yeast, acts synergistically with SFA in this process. This latter effect is connected to ergosterol accumulation within microsomal fractions from SFA-accumulating cells, which display highly saturated phospholipid content. Moreover, treating the cells with the molecular chaperone 4-phenyl butyrate abolishes UPR induction, suggesting that lipid-induced ER stress leads to an overload of misfolded protein that acts, in turn, as the molecular signal for induction of the UPR. The present data are discussed in the context of human diseases that involve lipid deregulation.

The endoplasmic reticulum (ER) consists of a network of intracellular membranes devoted to crucial cellular functions including lipid synthesis, storage and regulated release of calcium, and biosynthesis of proteins destined for either intracellular organelles or the cell surface. ER is also the site where secretory proteins are primarily assembled and folded. Accordingly, eukaryotic cells have developed quality control processes to monitor the proper folding of ER-borne proteins, a process referred to generally as ‘ER quality control’. When the ER fails to cope with misfolded or unfolded proteins, a complex cellular response, called the unfolded protein response (UPR), is triggered (for recent reviews, see refs (1–3)). UPR provides multiple strategies to avoid ER stress and to maintain ER integrity and secretory pathway function. In higher eukaryotes, the UPR is comprised of three distinct pathways controlled by transmembrane proteins that act as sensors, namely Inositol-Requiring Enzyme 1 (IRE1), Protein-kinase like Endoplasmic Reticulum Kinase (PERK) and Activating Transcription Factor 6 (ATF6). The IRE1 regulated pathway is conserved from yeast to humans. In the yeast Saccharomyces cerevisiae, accumulation of unfolded proteins results in the dimerization of Ire1p, a process that activates its cytosolic endoribonuclease function. Kar2p, an ER-resident member of the HSP70 family, is an important regulator of the UPR in both yeast and mammalian cells (the mammalian orthologue of Kar2p is known as Immunoglobulin-binding Protein (BiP)). In the original model, it was proposed that, in non-stressed cells, Kar2p associates with Ire1p to repress its activation. In response to accumulation of unfolded proteins in the ER, Kar2p dissociates from Ire1p, resulting in Ire1p dimerization and subsequent activation of its endoribonucelase function (3). The substrate for Ire1p endonuclease activity is the transcript of HAC1 (the yeast orthologue of mammalian XBP1), a transcription factor that binds to promoter unfolded protein response elements (UPREs) and regulates the transcription of more than 380 yeast genes, i.e. approximately 5% of all yeast genes (4). Recent studies suggest that the mechanisms of unfolded protein sensing in the ER and Ire1p activation may be more complex than previously expected. Crystal structure of the ER-conserved luminal domain (cLD) of Ire1p revealed that a cLD dimer can form a major histocompatibility complex (MHC)-like groove (5). By analogy with MHC, peptide fragments and, more speculatively, unfolded proteins could be directly captured by this groove. In the latest scenario, Ire1p activation would occur in two regulatory steps (6,7): in the context of unfolded protein accumulation, (i) Kar2p dissociation may lead Ire1p to dimerize and form cLD dimers and (ii) unfolded protein binding to Ire1p may tether dimerized cLD to result in oligomerized active Ire1p.

Ultimately, failure to handle ER stress can result in cell death (for reviews, see refs (8,9)). ER stress and subsequent UPR induction have been implicated in many human diseases including cancer, diabetes, atherosclerosis and late-onset neurological diseases, such as Alzheimer, Huntington, Parkinson and familial amyotrophic lateral sclerosis (1,10). While ER stress clearly contributes to the aetiology of these disorders, whether or not it is the direct cause of disease remains, in most cases, under debate.

The diversity of ER-related human pathologies also suggests that the origins of ER stress may vary in kind. Recently, particular attention has been given to the impact of perturbations of lipid homeostasis on this process. Indeed, ER stress has been reported in mammalian cells in response to various lipotoxic conditions. For example, ER stress is induced by chronic non-esterified saturated fatty acid (SFA) exposure in pancreatic β-cells, a process that could account for insulin-resistant type 2 diabetes in obese individuals (11). Furthermore, ER stress in macrophages has been reported in response to free cholesterol accumulation in the ER (12). This phenomenon has been considered as a contributor to plaque destabilization in advanced atherosclerotic lesions. Interestingly, transient cell adaptation to these lipid-stress conditions requires a functional UPR (11,13). Altogether, these observations suggest that maintenance of lipid homeostasis in the ER is an absolute requisite for cell survival.

Unicellular organisms may experience massive modification of their lipid composition when confronted with changes in their environment. S. cerevisiae, for example, is a facultative anaerobe that can grow under low oxygen, but in so doing, the yeast loses its abilities to synthesize ergosterol, its major sterol, and to desaturate fatty acids. This situation arises because several enzymes of the ergosterol pathway as well as Ole1p, the fatty acid desaturase (homologous to mammalian SCD1), require haem as their prosthetic group. Haem synthesis is oxygen-dependent (for review, see ref. (14)); therefore, under oxygen or haem deprivation, yeast cells are confronted with intracellular SFA accumulation and ergosterol depletion, unless the medium is supplemented with exogenous UFAs and sterols (15).

In the present study, we took advantage of this physiological peculiarity to study the impacts of perturbations to lipid homeostasis on ER-stress induction in yeast, focussing mainly on the relative contributions of SFA and sterol to this process.


Low unsaturated fatty acid levels induce ER stress

To evaluate the consequences of perturbations in lipid homeostasis on UPR induction, we first used a knockout mutant of the δ-aminolevulinate (ALA) synthase (hem1Δ, (16)). This strain can synthesize haem only if the medium is supplemented with ALA because in yeast, haem is required as the prosthetic group of several enzymes of the ergosterol pathway and Ole1p, the fatty acid desaturase (for a review, see ref. (14)). As a consequence, when grown in the absence of ALA, the hem1Δ strain is confronted with double-lipid depletion, i.e. loss of both ergosterol and unsaturated fatty acid (UFA) (15,17). When transferred from ALA-supplemented to unsupplemented Yeast Peptone Dextrose medium (YPD), hem1Δ cells stop growing as early as 5 h after the shift (17). This arrest correlates with a progressive drop in ergosterol amounts (twofold decrease after 5 h and fourfold decrease after 7 h) and an accumulation of SFAs [mainly myristic (C14:0), palmitic (C16:0) and stearic (C18:0) acids] at the expense of unsaturated forms [palmitoleic (C16:1) and oleic (C18:1) acids] (15). After 8 h of haem limitation, hem1Δ cells will start growing again if ergosterol and oleic acid are added to the medium, suggesting that, among the various cellular consequences of haem depletion, ergosterol and UFA starvation are the most detrimental for growth. In the following text, hem1Δ cells grown in unsupplemented YPD medium will be referred to as ‘lipid-depleted’, whereas hem1Δ cells grown with ALA supplementation will be referred to as ‘lipid-competent’ cells.

To evaluate ER stress in response to UFA and ergosterol starvation, UPR activation in hem1Δ cells grown under lipid-depleted conditions (Figure 1A) was assayed using a UPRE-CYC-LacZ reporter gene (18) (see Materials and Methods). As shown, significant levels of UPRE induction were observed as soon as 5 h after shifting to lipid-depleted conditions. This induction increased gradually up to 8 h after the shift. For comparative purposes, UPRE expression was also determined in the presence of 2 mM dithiothreitol (DTT), a strong reducing agent that prevents disulfide bond formation, thereby disrupting protein folding in the ER and inducing ER stress (Figure 1A) (19). Both lipid depletion and DTT addition resulted in UPRE inductions of the same order of magnitude (9.5 ± 0.5 and 20.1 ± 2.9 β-galactosidase units, respectively), confirming the physiological relevancy of UPR induction by lipid depletion.

Figure 1.

Activation of the UPR under conditions of lipid depletion. A) hem1Δ cells transformed with pJC104 (UPRE-CYC-LacZ) (hem1Δ [pJC104]) were grown to stationary phase in YPD medium supplemented with 80 μg/mL ALA, before transfer to early exponential phase (2 × 106 cells/mL) in fresh YPD medium with (open bars) or without (black bars) ALA. The times following the shift are indicated. UPRE activation was monitored by measuring β-galactosidase activities, as described under Materials and Methods. The grey bar represents UPRE activation measured from cells grown in YPD + ALA in the presence of 2 mM DTT for 8 h. B) Cells were grown to stationary phase as in (A) before transfer to YPD + ALA (ALA), YPD (YPD), YPD + 1% Tween-80 (v/v) (Ole), YPD + Erg 80 μg/mL (Erg) or YPD + 1% Tween-80 (v/v) + Erg 80 μg/mL (Erg + Ole). β-galactosidase activity was defined as OD420 min/mL. Values are means ± SD of at least three independent determinations.

Next, to differentiate between the relative contributions of ergosterol and UFA depletion to UPR induction, the same experiments were conducted on hem1Δ cells grown in the presence of an exogenous source of either oleate (ergosterol depletion) or ergosterol (UFA depletion). In contrast to haem-competent cells, when hem1Δ cells were grown in the absence of ALA, i.e. when sterol biosynthesis was compromised, they became capable of importing these essential molecules, provided they were present in the medium. Figure 1B shows that UFA depletion alone could account for full-fledged induction of UPRE-CYC-LacZ expression. It should be noted that the addition of an exogenous ergosterol source under these conditions resulted in an even stronger UPR than that observed as a result of double-lipid depletion (YPD, Figure 1B), suggesting synergistic relationship between ergosterol supplementation and UFA starvation (see below).

These results suggested that cellular UFA levels are a crucial parameter in the regulation of UPR induction. To confirm this hypothesis, UPRE-CYC-LacZ expression was monitored in an ole1Δ mutant grown with or without exogenous oleic acid (Figure 2A). Consistent with previous observations, ole1Δ cells grown in the absence of UFAs showed elevated UPRE activation, a response that was completely abolished by addition of UFA to the medium (Figure 2A). In a previous study, we reported that hem1Δ cells grown under UFA depletion resulted in the accumulation of short saturated fatty acyl chains within phospholipids (15). Therefore, it seemed of particular interest to determine if the same phenomenon could be observed under other UPR-activating conditions such as in ole1Δ cells grown in the absence of an exogenous UFA source. To this end, lipids were extracted from ole1Δ cells grown for 5 h in YPD medium, sufficient time to obtain maximal UPRE activation (Figure 2A), then subjected to phospholipid analysis by mass spectrometry (MS) (see Materials and Methods). As a control, the same experiments were performed on ole1Δ cells grown in oleic acid-supplemented medium. Figure 2B shows the relative distribution of phosphatidylcholine (PC) species under both culture conditions, obtained by comparison of positive ion mode spectra as described (15). Growth in unsupplemented YPD medium resulted in a decreased amount of PC species bearing long unsaturated chains (36:2, 36:1, 34:2, 34:1, 32:2, 32:1) in favor of species with shorter saturated acyl chains (32:0, 30:0, 28:0, 26:0, 24:0). Such rearrangements of the fatty acyl chain content were also observed in other phospholipid species (not shown). A general assessment of these compositional changes to PC species can be had by monitoring two parameters, the unsaturation ratio (Figure 2C) and the average number of carbons in the acyl chain (average chain length; Figure 2D). Thus, consistent with the detailed PC analysis, growth of the ole1Δ cells in the absence of exogenously supplied UFAs resulted in a dramatic decrease in unsaturation ratio and a small but significant decrease in the average chain length. As expected, these results were very similar to those previously reported for conditions of haem-induced UFA depletion (15).

Figure 2.

Impacts of desaturation preclusion on UPR activation and cellular PC content. ole1Δ[pJC104] cells were grown to stationary phase in YPD medium supplemented with 1% Tween-80 (v/v) (YPD + Ole), before shift to early exponential phase (2 × 106 cells/mL) in fresh YPD medium with (open bars; YPD + Ole) or without (black bars; YPD) 1% Tween-80 (v/v). A) UPRE activation was monitored by measuring β-galactosidase activities, as described under Materials and Methods. Times following the shift to fresh media are indicated. B) After 5 h growth in YPD (black bars) or YPD + Ole (open bars), lipids were extracted from ole1Δ [pJC104] cells and PC species were analyzed by MS in the positive ion mode, as described in Materials and Methods. The total carbon chain length (x) and the number of carbon–carbon double bonds (y) of PC molecular species are indicated (x:y). The unsaturation ratio (C) and the average chain length (D) were calculated from data displayed in B. Values are means ± SD of at least three independent determinations.

To evaluate the respective contributions of chain length and unsaturation level on UPRE activation, β-galactosidase activities were determined for hem1Δ cells grown in the presence of myristoleic acid (C14:1) instead of oleic acid (Figure 3). Cells grown under these conditions had phospholipids with short (15.5 carbon atoms versus 16.8 carbon atoms for oleate grown cells) but highly unsaturated (74 and 68% UFA for myristoleate and oleate grown cells, respectively) fatty acyl chains (15). As shown in Figure 3, myristoleic acid supplementation abolished β-galactosidase induction, showing that the unsaturation ratio of fatty acids within phospholipids rather than their length is the crucial parameter for UPRE activation.

Figure 3.

Supplementation with short unsaturated fatty acids prevents UPR induction in hem1Δ cells. hem1Δ[pJC104] cells were grown to stationary phase in YPD medium supplemented with 80 μg/mL ALA, before transfer to early exponential phase (2 × 106 cells/mL) in fresh YPD + Erg medium (black bars; Erg) or in YPD + Erg medium supplemented with 2 mM myristoleic acid (open bars; Erg + C14:1). The times following the shift are indicated. UPRE activation was monitored by measuring β-galactosidase activities, as described under Materials and Methods. Values are means ± SD of at least three independent determinations

UPR induction by lipids is not due to global secretory stress

It has been proposed that intracellular accumulation of SFA may induce disruption of ER function and structure both in lipotoxic Chinese Hamster Ovary (CHO) (11,20) and pancreatic β-cells (11), and that this disruption may be responsible for the triggering of ER stress and UPR response. Moreover, in yeast, secretory stress that results from disruption of the secretory pathway in sec mutants is also associated with UPRE activation (21).

To assess whether UPR induction could result from a global defect of the secretory pathway, the secretion of invertase was measured under conditions of high UPR induction (+Erg; Figure 4A). Internal and external invertase activities were determined using enzyme latency assays, as described elsewhere (Materials and Methods; (15)). As controls, invertase secretion was also measured in cells grown under conditions where no significant UPRE activation was detected (+ Ole; Figure 4A), and under conditions of low UPRE activation and normal growth (+ Erg + Ole; Figure 4A). As shown in this figure, the kinetics of invertase secretion was very similar under all conditions. More specifically, no intracellular accumulation of invertase could be detected under lipid stress, as would have been expected in the case of a general block of the secretory pathway, as observed with sec mutants (15).

Figure 4.

Effects of lipid depletion on invertase secretion and plasma membrane delivery of Pma1p. A) hem1Δ cells (upper panels) were grown as described in the legend of Figure 1 to stationary phase in YPD + ALA before shifting to fresh YPD + 1% Tween-80 (v/v) (Ole), YPD + Erg 80 μg/mL (Erg) or YPD + 1% Tween-80 (v/v) + Erg 80 μg/mL (Erg + Ole) for a further 7 h incubation. Invertase was induced by glucose limitation in cells as described in the Materials and Methods section. Samples were removed at the time-points indicated and the activity of internal and secreted invertase was determined as described in Materials and Methods. Closed symbols, extracellular invertase activities (e); open symbols, intracellular invertase activities (i). B) hem1Δ [Yiplac204-2HSEpr-HA-PMA1] cells were grown in the indicated media as described in (A) for 7 h, before incubation for 15 min at 39°C for induction of Pma1p-HA synthesis. Sixty minutes after the heat shock, the cells were fixed and processed for immunofluorescence with antibodies to HA. Bars: 5 μm

We also evaluated whether ER stress could be correlated with a defect in trafficking of integral plasma membrane proteins, by monitoring biogenesis of the proton ATPase Pma1p (Figure 4B). For this purpose, a Hemaglutinin-tagged (HA) version of Pma1p under control of a heat-shock promoter was used (22,23), that allows its synthesis by a transient shift to elevated temperature. The newly synthesized protein can be tracked by immunofluorescence using an anti-HA antibody (Materials and Methods). In the experiment displayed in Figure 4B, cells grown for 7 h under ER-stress conditions (+ Erg) and control conditions (+ Ole and + Erg + Ole) were shifted to 39°C for 15 min and Pma1p-HA was visualized after 2 h following the shift. Interestingly, Pma1p-HA could be detected at the cell periphery under all conditions, showing that ER stress does not prevent delivery of Pma1p to the plasma membrane.

Therefore, we conclude from these experiments that lipid-induced ER stress is not related to a global block of the secretory pathway.

Exogenous saturated fatty acids also induce ER stress

Previous studies reported that exogenously supplied SFAs can generate ER stress in various types of mammalian cells, ranging from Chinese Hamster Ovary cells (20) to human pancreatic β-cells (11) and liver cells (24,25). In the two human cell lines, it has been suggested that ER stress could be the central feature of peripheral insulin resistance and type 2 diabetes in obese patients (24,25). Given this possibility, we investigated whether exogenously supplied SFA could also induce ER stress in yeast cells in which endogenous fatty acid desaturation was precluded. In a previous study (26), the growth of hem1Δ cells was evaluated over a range of ratios of Tween-80 (oleic acid source) to Tween-40 (palmitate source; C16:0). To avoid working with high concentrations of Tween-40 that might have impaired growth by indirect detergent-related effects, Tween-80 amounts were decreased to 0.1‰ (v/v) from the 1% (v/v) used for our standard culture conditions. As shown in Figure 5A, decreasing Tween-80 to 0.1‰ (Erg + Ole 0.1‰) was sufficient to induce a significant UPRE induction compared to 1% supplementation (Erg + Ole 1%). This induction correlated with the accumulation of short saturated fatty acyl chains within PC (Figure 5B) and, as a corollary, a decrease in the average acyl chain length (Figure 5C) and unsaturation ratio (Figure 5D). Consistent with these findings, adding increasing amounts of palmitate, from 0 to 0.6‰, resulted in increased UPRs. Interestingly, omitting ergosterol from a medium supplemented with palmitate 0.6‰ (Ole 0.1‰ + Pal 0.6‰) caused a twofold decrease of UPR induction by comparison with the same culture conditions plus Erg (Erg + Ole 0.1‰ + Pal 0.6‰). This result demonstrated that ergosterol and exogenously supplied SFA act synergistically to induce the UPR, as previously observed when endogenous fatty acid desaturation was precluded (Figure 1B). When the profiles of PC species are compared (Figure 5B), the most significant effect of palmitate addition (Erg + Ole 0.1‰ + Pal 0.6‰) was an increase of 32:0 species corresponding to PC molecules bearing two C16:0 acyl chains, consistent with the incorporation of these exogenously supplied Fatty Acids (FAs) into phospholipids. From a global point of view, palmitate supplementation had little impact on the average fatty acyl chain length but did result in a significant decrease in the unsaturation ratio (Figure 5D).

Figure 5.

Effects of exogenously supplied saturated fatty acids on UPR activation and cellular PC content. hem1Δ[pJC104] cells were grown to stationary phase as in the legend of Figure 3 before transfer to YPD + 1% Tween-80 (v/v) + Erg 80 μg/mL (Erg + Ole 1%), YPD + 0.1‰ Tween-80 (v/v) + Erg 80 μg/mL (Erg + Ole 0.1‰), YPD + 0.1‰ Tween-80 (v/v) +0.2‰ Tween-40 (v/v) + Erg 80 μg/mL (Erg + Ole 0.1‰ + Pal 0.2‰), YPD + 0.1‰ Tween-80 (v/v) +0.6‰ Tween-40 (v/v) + Erg 80 μg/mL (Erg + Ole 0.1‰ + Pal 0.6‰) or YPD + 0.1‰ Tween-80 (v/v) +0.6‰ Tween-40 (v/v) (Ole 0.1‰ + Pal 0.6‰). A) UPRE activation was monitored by measuring β-galactosidase activities 8 h after shift to the indicated medium. B) After 8 h growth in Erg + Ole 1% (open bars), Erg + Ole 0.1‰ (black bars) or Erg + Ole 0.1‰ + Pal 0.6‰ (grey bars), lipids were extracted from cells and PC species were analyzed by MS in the positive ion mode, as described in Materials and Methods. The average chain length (C) and the unsaturation ratio (D) were calculated from data displayed in B. Values are means ± SD of at least three independent determinations.

In summary, these results show that exogenously supplied palmitate induces UPRE activation in yeast and modifies the composition of PC species, resulting in a decrease in their unsaturation ratio. It has been reported that increasing amounts of palmitate have a proportionally negative impact on yeast growth, showing a strict connection between the level of UPR induction and cell survival (26).

These results underscore the importance of the phospholipid unsaturation ratio as a crucial determinant of the UPR response, irrespective of the origin (endogenous or exogenous) of the perturbation.

UPR induction under lipotoxic conditions depends on the Ire1p/Hac1p pathway and is required for viability

As stated earlier, the main pathway for UPR induction in yeast originates at the ER-resident transmembrane protein kinase/endoribonuclease Ire1p that regulates the splicing of HAC1 mRNA. To evaluate the role of the UPR in yeast adaptation to lipid-induced ER stress, double mutant hem1Δ, ire1Δ and hem1Δ, hac1Δ strains were constructed. As expected, UPRE activation under lipid depletion was abolished in both double mutant strains, showing that the observed lipid-mediated UPR induction depends on the canonical Ire1p/Hac1p pathway (our unpublished data). Impacts of lipids on the growth of the double hem1Δ, ire1Δ mutant are displayed in Figure 6. Interestingly, although the growth curve of the hem1Δ, ire1Δ strain was indistinguishable from that of the wild-type under lipid-competent conditions (i.e. ALA and Erg + Ole 1%; Figure 6), it was suppressed significantly under conditions of SFA accumulation that resulted from either endogenous preclusion of fatty acid desaturation (Erg) or exogenous addition (Erg + Ole 0.1‰ + Pal 0.6‰). These results demonstrate that the UPR response induced by lipid stress is required for transient adaptation of yeast cells to SFA accumulation.

Figure 6.

The UPR pathway is required for optimal growth under lipid-induced ER stress. hem1Δ[pJC104] and hem1Δ, ire1Δ[pJC104] cells were grown in YPD + ALA to stationary phase before transfer to YPD, YPD + ALA (ALA), YPD + Erg 80 μg/mL (Erg), YPD + 1% Tween-80 (v/v) + Erg 80 μg/mL (Erg + Ole 1%) or YPD + 0.1‰ Tween-80 (v/v) +0.6‰ Tween-40 (v/v) + Erg 80 μg/mL (Erg + Ole 0.1‰ + Pal 0.6‰) as in the legend of Figure 5. Attenuance of the cultures was determined at the indicated time-points after shift to the corresponding media.

Composition and organization of microsomal fractions from SFA-accumulating cells

As the UPR originates in the ER, it was important to assess whether the perturbations to lipid composition observed in total cells could be replicated using ER-enriched fractions. For this purpose, microsomal fractions were prepared as described by Zinser et al. (27) (see also Materials and Methods) from hem1Δ cells grown under lipid-competent conditions or under accumulation of SFA from endogenous (lipid depleted; YPD) or exogenous (Ole 0.1‰ + Pal 0.6‰) origins. When these microsomal fractions were compared with total cell extracts by western blotting, the ER-resident protein Sec 61p was enriched 10.7-fold relative to the plasma membrane marker Pma1p (unpublished data). These results are similar to the relative enrichment obtained by Zinser and coworkers (27).

As shown in Figure 7A, the patterns of PC species from microsomal fractions obtained from cells grown under lipid-competent and lipid-depleted conditions were very similar to that already reported for total cells (15): lipid depletion (YPD) resulted in the accumulation of PC species with short fatty acyl chains (24:0, 26:0, 28:0 and 30:0) at the expense of species bearing longer unsaturated chains (most significantly 32:2 and 34:2). This behavior resulted in decreases of both the average fatty acyl chain length (Figure 7B) and unsaturation ratio (Figure 7C).

Figure 7.

PC composition and generalized polarization of Laurdan in microsomes from SFA-accumulating cells. Microsomes were prepared, as described in Materials and Methods, from hem1Δ cells grown for 8 h in YPD, YPD + ALA (ALA) or YPD + 0.1‰ Tween-80 (v/v) +0.6‰ Tween-40 (v/v) (Ole 0.1‰ + Pal 0.6‰) medium. Lipids were extracted from 1 mg of microsomal fractions before analysis of PC species by MS, in the positive ion mode (A). The average chain length (B) and the unsaturation ratio (C) were calculated from data displayed in A. Values are means ± SD of at least three independent determinations. D) Normalized emission spectra of Laurdan at 7°C (left panel) and 27°C (right panel), using excitation at 350 nm. Generalized polarization (GP) values from representative experiments are indicated. See text for details.

Interestingly, microsomes isolated from cells grown in the presence of an exogenous source of palmitate displayed noticeable differences as compared to lipid-depleted microsomes (Figure 7A), i.e., lower amounts of short chain containing species and higher levels of 32:0 (C16:0 + C16:0; a hallmark of exogenous palmitate incorporation into PC) and, more surprisingly, of 34:1 PC species (C16:0 + C18:1 and/or C16:1 + C18:0). Overall, it appeared that palmitate accumulation into PC was not counteracted by short chain species as the average fatty chain length remained very equivalent to what observed with lipid-competent cells (Figure 7B).

It is well known that the length and unsaturation level of the acyl chains of phospholipids can influence the general organization of biological membranes. Specifically, at physiological temperatures, phospholipids can be in either the fluid state (liquid disordered, Ld) or the solid–liquid gel state (G), in which there is a preference for phospholipids with long saturated chains (for review, see ref. (28)). Laurdan (6-Lauryl-2-dimethylaminophtalene) fluorescence is unique in that it displays different and well resolvable spectral parameters between the two phases and can therefore be used for the detection and relative quantification of Ld and G phases (29,30). More specifically, in G-phase membranes with reduced molecular mobility in the hydrophobic region, the dipolar relaxation is too slow to change the properties of fluorescent emission; thus, the emission spectrum displays only one major peak at 440 nm. By contrast, in Ld membranes where the interior is fluid and allows the movement of lipid molecules, dipolar relaxation of water molecules occurs. This event leads to a red shift of about 50 nm (490 nm) in the Laurdan emission spectra (29).Parasassi et al. (29) have developed the concept of general polarization (GP) of Laurdan fluorescence. The definition of GP is:


where Ig and If are the fluorescence intensities at the emission maxima of Laurdan in gel (440 nm) and fluid phase (490 nm), respectively. Higher GP values therefore indicate a membrane in the G phase or a more tightly packed structure, with a low rate of solvent relaxation.

Representative emission spectra of Laurdan obtained upon excitation at 350 nm on microsomal fractions from lipid-competent (ALA) and SFA-accumulating (YPD and Ole 0.1‰ + Pal 0.6‰) cells are presented in Figure 7D. The spectra were obtained at low (7°C) and physiological (27°C) temperatures and were normalized to the emission at 440 nm. In each case, the corresponding GP values are indicated. Interestingly, spectra obtained with SFA-accumulating microsomes were not significantly different from those obtained with ALA microsomes, at both temperatures. In each case, a shift to the lower temperature resulted in a characteristic increase in GP that corresponded to a temperature-related increase of membrane order (31).

Fluorescence polarization of diphenyl-1,3,5-hexatriene (DPH) in microsomal membranes was used to confirm the results obtained with Laurdan. DPH is known to label the core of the membrane phospholipid bilayer, where its quantum yield is greatly enhanced and thus, it allows the degree of fluidity in the membrane lipid core to be monitored. DPH anisotropy values for microsomal fractions at 27°C were 0.191 ± 0.04, 0.195 ± 0.015, and 0.200 ± 0.04 for ALA, YPD and Ole 0.1‰ + Pal 0.6‰ cells, respectively. The similarity in fluidity between these three fractions provided confirmation of the Laurdan results.

From these experiments, we conclude that UPR induction by SFA accumulation cannot be simply correlated with a global change of ER membrane organization at physiological temperature.

Exogenous SFA alter ER morphology

In a next step, electron microscopy was carried out to explore whether SFA accumulation may alter ER morphology. Figure 8A,B reveals that ER morphology is very similar in lipid-competent and lipid-depleted cells. More specifically, in both conditions, peripheral ER appears as flat cisternae that characteristically associate with the plasma membrane (Figure 8C). This shows that endogenous SFA accumulation has low impact on ER morphology. By contrast, treatment of the cells with palmitate markedly altered cell morphology (Figure 8D–F). Palmitate-treated cells displayed two distinct morphologies. In the first population of cells, the major effects were the apparition of indentations at the plasma membrane level, detachment of the ER from the cell periphery and a swelling of this organelle (Figure 8D). In the second population, which may correspond to more severely affected cells, the peripheral ER could not be observed in association with the plasma membrane anymore, but appeared as electron-lucent clefts extending throughout the cytoplasm (Figure 8F).

Figure 8.

Effects of SFA accumulation on cell morphology. hem1Δ cells were grown in YPD + ALA to stationary phase, transferred for 8 h to YPD + ALA (lipid-competent), YPD (lipid-depleted), or YPD + 0.1‰ Tween-80 (v/v) +0.6‰ Tween-40 (v/v) (palmitate-treated) and examined by electron microscopy. Lipid-competent (A and C) and lipid-depleted cells (B) displayed very similar morphologies, with the familiar appearance of peripheral ER. The arrows in the boxed area (A) enlarged in C show characteristic ER cisternae. By contrast, in palmitate-treated cells (D–F) aberrant ER structures could be observed, ranking from slightly dilated cisternae, detached from the cell periphery (D and E), to electron-lucent clefts expanding throughout the cytoplasm (F). In many palmitate-treated cells, the plasma membrane appeared indented [boxed area (E) enlarged in D]. N, nucleus; V, vacuole and m, mitochondria. Bars: 500 nm (A, B, E and F) or 150 nm (C and D

Ergosterol accumulates in microsomal fractions from UFA-depleted cells

Another intriguing observation was the overinduction of UPR after addition of ergosterol to lipid-depleted cells (see above; Figure 1B). Could this effect be related to ergosterol accumulation within the microsomal fractions? Indeed, in human macrophages, cholesterol overload in microsomes has been shown to induce ER stress, a phenomenon that is likely to promote progression of atherosclerosis (32).

Ergosterol levels in microsomal fractions obtained from cells grown under the various culture conditions are displayed in Figure 9. Microsomal fractions from cells grown under lipid-competent conditions (ALA), lipid-depleted (YPD) and UFA-supplemented (+ Ole) conditions displayed similar amounts of ergosterol, i.e. 1.5 ± 0.8 μg ergosterol/mg protein. These amounts are very close to those reported by Zinser et al. (27). This result also confirmed the purity of our microsomal fractions because even though ergosterol is synthesized in the ER, it accumulates mainly at the plasma membrane (27). Interestingly, ergosterol addition to UFA-depleted cells resulted in a 100-fold accumulation of ergosterol in microsomal fractions (140 ± 22 μg ergosterol/mg protein; Erg, Figure 9). Addition of exogenous UFA led to decreased ergosterol accumulation but amounts were still significantly higher than those in microsomes obtained from cells grown without exogenous ergosterol (7.2 ± 0.8 μg ergosterol/mg protein; Erg + Ole, Figure 9). Note that these increased amounts of ergosterol correlated very well with the amplitude of UPRE activation (Figure 1B).

Figure 9.

Ergosterol content in microsomal fractions from. hem1Δcells grown on different lipid supplemented media Microsomes were prepared from hem1Δ cells grown for 8 h in YPD + ALA (ALA), YPD, YPD + 1% Tween-80 (v/v) (Ole), YPD + Erg 80 μg/mL (Erg) and YPD + 1% Tween-80 (v/v) + Erg 80 μg/mL (Erg + Ole). After lipid extraction, ergosterol amounts were determined by gas chromatography using cholesterol as a standard, as described in Materials and Methods

A molecular chaperone, 4-PBA, prevents UPR induction in cells subjected to lipid-induced ER stress

Next, we asked how the observed perturbations of lipid homeostasis could induce ER stress and UPR induction. A reasonable hypothesis was that lipid perturbations in the ER may reduce the overall ER-protein folding capacity, resulting in a misfolded protein overload in this organelle.

4-phenyl butyrate (4-PBA) is a chemical chaperone that is known to stabilize protein conformation, improve ER folding capacity and facilitate the trafficking of mutant proteins (33). Relevantly, 4-PBA can restore the growth of yeast ire1Δ cells pretreated with tunicamycin, a cytotoxic agent that induces accumulation of misfolded proteins by inhibiting N-linked glycosylation (34). Recently, 4-PBA has been shown to prevent aggregation of misfolded proteins both in vivo(34,35) and in vitro(34). We therefore evaluated whether 4-PBA addition could prevent UPR induction under our lipid-stress conditions.

As shown in Figure 10A, in the hem1Δ background, 4-PBA completely suppressed UPRE activation because of intracellular accumulation of SFA, and either in the absence (YPD) or in the presence of added ergosterol (Erg). The exact same effect of 4-PBA was seen in response to SFA accumulation in the ole1Δstrain (Figure 10B) and in palmitate-treated cells (not shown). As a control, the experiments were repeated in the presence of tunicamycin. As previously reported (36), addition of 1μg/mL tunicamycin triggered strong UPRE activation (Figure 10C) but this response was decreased significantly (fivefold reduction) when cells were incubated concomitantly with 4-PBA. The same results were obtained with DTT, another agent that induces ER stress (data not shown). Taken together, the results suggest that UPRs induced by the various conditions described earlier are indeed related to ER overload with misfolded proteins and this process can be attenuated by 4-PBA treatment.

Figure 10.

4-PBA decreases UPRE activation under lipid-induced ER stress. A) hem1Δ[pJC104] cells were grown for 8 h in YPD + ALA (ALA), YPD or YPD + Erg 80 μg/mL (Erg) for 8 h before determination of β-galactosidase activity, as in the legend of Figure 1. When indicated, 4-PBA was added to the culture at a final concentration of 5 mM. B) ole1Δ[pJC104] cells were grown to stationary phase in YPD + 1% Tween-80 (v/v) and shifted to YPD (–Ole), YPD + 1% Tween-80 (v/v) (+ Ole) or YPD + 5 mM 4-PBA (–Ole + 4-PBA) for 5 h before determination of β-galactosidase activity. C) hem1Δ[pJC104] cells were grown in YPD + ALA (ALA) as in A, in the presence of 1 μg/mL Tunicamycin and 5 mM 4-PBA, as indicated. D) KMY81[pJC104] cells bearing the kar2-1 thermosensitive mutation were grown at 23°C in YPD+50 μg/mL adenine before being shifted to the non-permissive temperature (37°C) for 8 h. UPRE activation was monitored by measuring β-galactosidase activity. Values are means ± SD of at least three independent determinations

The suppression of UPRE activation by 4-PBA could be explained by non-specific effects on the UPR signalling pathway rather than by direct action as a molecular chaperone. More specifically, one may argue that 4-PBA impairs Ire1p oligomerization/activation or interferes with the UPR reporter gene assay. To test this possibility, we used a yeast strain with a mutated allele of KAR2 (kar2-1; (37)). The kar2-1 mutant belongs to a subclass of kar2 mutants that display constitutive induction of the UPR cascade (type S mutants; (37)). It has been proposed that kar2-1p mutant protein can no longer bind Ire1p, resulting in constitutive activation of the latter (37). As Figure 10D shows, 4-PBA did not significantly decrease constitutive UPRE induction in the kar2-1 strain, ruling out indirect effects of 4-PBA on this process.


An increasing number of reports link cellular lipid homeostasis and its significance for organelle function and/or integrity, with the aetiology of several human pathologies. The finding, for example, that elevated SFAs (namely palmitate (11,20) and cholesterol levels (32)) can trigger a so-called ER stress that might, in turn, promote the progression of diseases of high incidence and morbidity (e.g. obesity-related insulin resistance and atherosclerosis), has shed new light on these complex, intricate processes.

The first objective of the present study was to evaluate whether a simple unicellular eukaryote such as S. cerevisiae, which is not routinely exposed to high-fat diets, could also experience ER stress under lipotoxic conditions. If so, would the relevant signalling pathways be conserved from yeast to humans, independent of the specific function associated with the particular cell type? One way to measure this phenomenon is to monitor the induction of the UPR, an ER-borne signal cascade that is known to be stimulated in mammalian cells by lipotoxic conditions (25).

Interestingly, we showed that yeast do indeed experience ER stress when faced with intracellular accumulation of SFAs. These stress conditions were generated either by feeding cells with extracellular SFA (Figure 5A), or by preventing endogenous fatty acid desaturation via downregulation of the activity of Ole1p, the unique yeast fatty acid desaturase. The latter effect was achieved either by haem depletion (Figure 1) or by using an ole1Δmutant (Figure 2). As suggested for mammalian cells (25), the UPR appears to help yeast cells cope with lipid-mediated ER stress triggered by SFA of either endogenous or exogenous origin, since cells bearing deletions in the IRE1 or HAC1 genes, both essential for the UPR cascade, arrested growth prematurely (Figure 6).

While such an effect caused by exogenous SFA feeding has been described in mammalian cells (11,20,38), we believe this is the first time that ER stress triggering by disruption of endogenous fatty acid desaturation has been reported. These observations should be considered in light of recent data concerning stearoyl-CoA desaturase 1, SCD1, the rate-limiting enzyme in monounsaturated fatty acid synthesis in humans, that displays a high degree of similarity with Ole1p (for review, see ref. (39)). Indeed, it has been shown that SCD1 is upregulated in obesity and may provide partial protection from obesity-induced insulin resistance (40,41). In addition, expression of SCD1 protects pancreatic β-cell from lipoapoptosis (42,43). As pancreatic β-cell death in obesity has been directly related to ER stress (44), these recently published findings, together with our own data, suggest that by preventing intracellular SFA accumulation, Ole1p/SCD1 activity may be a crucial factor in avoiding ER stress and consequent cell death.

Lipid composition of the ER is different from that of other organelles. First, it has the highest proportion of PC, a phospholipid that is highly unsaturated under normal conditions ((15); Figure 7), relative to other phospholipid species (45). Second, ergosterol amounts are very low in this organelle ((27); Figure 9). Collectively, these two features suggest that ER membranes have relatively low lipid chain order, a parameter that may be essential for specialized ER-based processes, such as protein translocation (46).

An important finding from the present study is that excess endogenous, or exogenous, SFA has a profound impact on ER phospholipid species, albeit in different ways. Preclusion of endogenous fatty acid desaturation led to both a decrease in the unsaturation ratio of PC species because of accumulation of SFA at the expense of unsaturated forms (as monitored by mass spectrometry), and also an unexpected decrease in the average length of fatty acyl chains (Figure 7A–C and (15)). By contrast, exogenously added palmitate did not significantly impact the average acyl chain length but resulted in a dramatic decrease in the unsaturation ratio of PC species (Figure 7A–C). Initially, we postulated that a decrease in the phospholipid unsaturation ratio might affect the overall fluidity in microsomal fractions, a phenomenon that could account for the UPR induction observed after preclusion of fatty acid desaturation or palmitate treatment. This hypothesis was tested directly by measuring the general fluorescence (GP) of Laurdan in microsomal fractions obtained from lipid-competent and SFA-accumulating cells (Figure 7D), but no significant differences in GP were observed between the three samples. Therefore, ER-stress induction cannot be simply related to a global change in lipid order at the ER level.

By contrast, striking differences on ER morphology between cells grown under the two different conditions of SFA accumulation were revealed by electron microscopy. Whereas cells grown under preclusion of endogenous desaturation were undistinguishable from lipid-competent cells (Figure 8A–C), palmitate-treated cells displayed marked alterations of ER morphology (Figure 8D–F). In the most affected cells, ER is dilated and forms clefts that extend throughout the cytoplasm (Figure 8F). This pattern is very similar to what has already been reported for palmitate-treated INS-1 (Insulin-producing) β-pancreatic cells (11,47). We proposed previously that the appearance of short chain fatty acids under desaturation preclusion may compensate for the accumulation of longer saturated chains (15). As short chain compensation does not occur as efficiently in palmitate-treated cells than under endogenous SFA accumulation (Figure 7A–C), data from the present study tend to confirm this hypothesis: shortening of the acyl chain may help to maintain ER morphology and enable cell survival, at least transiently. These compensatory effects probably account for efficient secretory capacity of the cells under preclusion of endogenous desaturation, as the secretion of invertase and the cell surface delivery of the plasma membrane proton ATPase PMA1p were largely unaffected under desaturation preclusion (Figure 4).

On the other hand, if the global order of microsomal fractions is not significantly modified under SFA accumulation, there may be some local impacts on lipid chain packing in response to the decrease in phospholipid unsaturation level, which could result in protein misfolding. Evidence to strengthen this hypothesis comes from experiments using 4-PBA. 4-PBA acts as a chemical chaperone by preventing protein aggregation in vivo(34,35). Kubota et al. (34) demonstrated that 4-PBA possesses chaperone activity in vitro, as it can prevent the aggregation between chemically denatured α-lactalbumine and bovine serum albumin. In our hands, 4-PBA almost completely abolished UPR induction triggered by SFA accumulation (Figure 10A,B). From these results, we postulate that lipid-related UPR induction is probably connected to the accumulation of misfolded proteins in the ER. It is known that integral proteins in artificial systems tend to partition preferentially to lipid bilayers in the Ld phase rather than the G phase (for review, see ref. (48)). Interestingly, in the G phase, polytopic proteins tend to form aggregates (48). Moreover, protein translocation across the ER bilayer has been shown to be highly sensitive to lipid composition (46,49). Thus, it is possible that local increases in membrane order may interfere with the folding and/or translocation of selective client proteins, a phenomenon that could account directly for the observed UPR induction. According to this hypothesis, whereas induction of the UPR may be sufficient to meet minimal endogenous protein folding demands under desaturation preclusion (thus allowing normal invertase secretion and Pma1p cell surface delivery), it may be limiting in palmitate-treated cells. As a result, misfolded protein overload could account for the alterations in ER morphology observed under these latter conditions.

Another important finding from the present study is that ergosterol has a synergistic effect with SFA on UPR induction (Figure 1B and Figure 5A). Cholesterol accumulation in the ER of human macrophages induces ER stress, a process that has been proposed to account for cholesterol-induced apoptosis of macrophages (12,50). We found that ergosterol also induced ER stress in yeast and that it was mediated via the same pathway as SFA-induced ER stress because: (i) the process involved the IRE1/HAC1 pathway (Figure 6) and (ii) UPR induction was abolished by 4-PBA whether excess SFA came from an exogenous or endogenous source (Figure 10A, B). This latter observation also suggests that induction of UPR by ergosterol probably relies on the accumulation of misfolded proteins in the ER, as proposed for SFA. Interestingly, the level of UPRE activation is proportional to the amounts of ergosterol that accumulate in the microsomal fractions; this level increases dramatically in cells in which desaturation is precluded compared with the same cells grown with exogenous UFA (140 ± 21 μg ergosterol/mg protein and 7.2 ±μg ergosterol/mg protein, respectively; Erg and Erg + Ole, Figure 9). Moreover, this ergosterol accumulation in microsomes (100-fold) is much higher than that observed in total cells, since we showed previously that ergosterol amounts are very similar in cells grown in hem1Δ cells grown under the same range of conditions (15). This suggests that ergosterol is somehow ‘glued’ in the ER in SFA-accumulating cells. Surprisingly, it appears that yeast cells can accommodate these huge amounts of ergosterol, at least transiently, as their secretory capacity is not impaired (Figure 4) and lipid-deficient cells grown in the presence of ergosterol can complete an additional cell division as compared to the same cells grown in the absence of ergosterol (unpublished data). This sterol tolerance does rely, however, on a functional UPR, as deletion of the IRE1 gene confers an increased sensitivity to exogenous ergosterol on the cells (Figure 6). To account for the high ergosterol accumulation in microsomes from UFA-depleted cells as compared with the same cells grown in oleate-supplemented medium, we first hypothesized that this may be due to different relative capacities of the two cell populations to store excess ergosterol in lipid bodies as ergosterylesters. This process is known to help mammalian cells cope with harmful accumulation of cholesterol (50,51) and sterols are preferentially esterified as sterylesters with UFA (17). However, it is also known that exogenous ergosterol is not esterified in yeast, even in the presence of UFA (52). Indeed, we confirmed that no detectable amounts of ergosterylesters were synthesized, even in the presence of an extracellular source of UFA (data not shown), conditions where ergosterol accumulation in microsomes is lower (Erg + Ole; Figure 9). Our current hypothesis is that ergosterol trapping in the ER is probably related to the phospholipid composition, as it has been shown in artificial membranes that cholesterol has a much higher affinity for saturated phospholipids than for unsaturated ones (53,54). This observation may have general significance, since genetic links between obesity and atherosclerosis have been reported (for recent review, see ref. (55)). We propose that the high saturation levels of phospholipids that result from decreased desaturation capacity (i.e. decreased SCD1/Ole1p activity) may render the cell more susceptible to toxic sterol effects by favouring their sequestration in the ER.

Finally, the present study points out that molecular chaperones such as 4-PBA, for which potential therapeutic applications in the treatment of obesity-related insulin resistance have been proposed (24), may act as regulators of sterol-induced ER stress. Testing macrophage protection from free cholesterol-induced apoptosis (32) by 4-PBA should provide additional information as to the possible use of such molecules to delay or, ideally, prevent progression of atherosclerotic lesions. In this context, yeast may be an excellent tool for the development of new high throughput screening strategies for the identification of therapeutic molecules in the field of lipid-induced ER stress diseases.

Materials and Methods

Yeast strains and culture conditions

S. cerevisiae strains used in this study are listed in Table 1. Cells bearing the hem1Δ mutation were grown aerobically at 28°C in YPD medium [1% yeast extract (w/v), 1% peptone (w/v), 2% glucose (w/v)] supplemented with 80 μg.mL−1δ-aminolevulinate (ALA, referred to as lipid-competent conditions). Haem-induced ergosterol and UFA depletions (referred to as lipid depletion) were obtained by cultivating the cells in unsupplemented YPD. Alternatively, YPD was supplemented with 80 μg.mL−1 ergosterol and/or 1% Tween-80 (v/v), used as the source of oleic acid (C18:1). Where indicated, Tween-80 was replaced by myristoleic acid (C14:1) at a final concentration of 2 mM (Sigma). For competition experiments, Tween-80 amounts were set at 0.1‰ (v/v) and Tween-40, as a source of palmitate (C16:0), was added at 0.2‰ (v/v) or 0.6‰ (v/v), as indicated. ole1Δ cells were grown in YPD with or without supplementation with 1% Tween-80 (v/v) as indicated. The KMY81 strain bearing the kar2-1 mutation (37) was grown in YPD + adenine (50 μg.mL−1) at 23°C to stationary phase before transfer to the non-permissive temperature of 37°C for 8 h. When indicated, 4-phenylbutyric acid (4-PBA; Alfa Aesar), dithiothreitol (DTT; Sigma) and tunicamycin (Sigma) were added from aqueous stock solutions (equilibrated to pH 7.4 with Na0H for the 4-PBA stock) to the appropriate final concentrations.

Table 1.  Yeast strains used in this study.
hemaMating type (MAT)αtrp1 his3 ura3 leu2 hem1::LEU2Progeny of FY1679α x FYHO4 (17)
hem1Δα(MAT)αtrp1 his3 ura3 leu2 hem1::LEU2Progeny of FY1679α x FYHO4 (17)
ole(MAT)αhis3 leu2 ura3 YGL055w::kanMXProgeny of diploid Y24422 (EUROSCARF)
hac(MAT)α his3 leu2 lys2 ura3 YFL031w::kanMX4Y15650 (EUROSCARF)
hem1Δ, hac(MAT)ahis3 ura3 leu2 trp1 hem1::LEU2 YFL031w::kanMX4This study, progeny of hem1Δ a x Y15650
ire(MAT)α his3 leu2 lys2 ura3 YHR079c::kanMX4Y11907 (EUROSCARF)
hem1Δ, ire(MAT)αhis3 ura3 leu2 trp1 lys2 hem1::LEU2 YHR079c::kanMX4This study, progeny of hem1Δ a x Y11907
KMY81(MAT)ahis3-11 his4?115 ura3-1 leu2-3,112 leu1 ade2-1 can1-100 kar2-1Kimata et al. (37)

β-galactosidase assays

To assay UPR induction, cells were transformed with plasmid pJC104 bearing a UPRE-CYC-LacZ gene fusion, provided by Dr Peter Walter (University of California, USA) (18). β-galactosidase assays were performed at 30°C as previously described (56) by measuring the increase in absorbance at 420 nm with o-nitrophenyl-β-D-galactoside as the substrate, after permeabilization of the cells with chloroform and sodium dodecyl sulfate.

Preparation of microsomal fractions

Microsomal fractions were prepared as described by Zinser et al. (27) with slight modifications. Briefly, approximately 2 × 1010 cells grown under the conditions indicated were harvested, washed twice with water and disrupted with glass beads (diameter 0.3–0.4 mm; Sigma) in 5 mL 250 mM Tris, 5 mM ethylenediaminetetraacetic acid (EDTA), 500 μM phenylmethylsulfonyl fluoride, containing 20 μL of protease inhibitor cocktail (Sigma). After the addition of 5 mL glycerol 20% (v/v), the homogenate was centrifuged for 10 min at 700 ×g to remove cellular debris and the supernatant centrifuged for a further 20 min at 20 000 ×g. The 20 000 ×g supernatant was centrifuged for 1 h at 100 000 ×g (TFT 50.38 rotor; Beckman). The resulting pellet was suspended in 10 mM Tris–HCl, pH 7.4, in a 10-mL potter with a tightly fitting pestle. The suspension was then centrifuged at 20 000 ×g for 20 min to remove most of the remaining contaminating plasma membrane and mitochondria. The resulting supernatant was centrifuged at 100 000 ×g for 1 h, yielding a colorless, opaque pellet that consisted mostly of microsomal membranes. This pellet was resuspended in 500 μL Glycerol Tris-HCl (GTH) buffer [glycerol 20% (v/v), 10 mM Tris–HCl, pH 7.4]. The total protein amount ranges typically between 2 and 3 mg per sample, as determined by the Bicinchoninic acid kit (Sigma). Purity of the microsomal fractions was evaluated by measuring the relative enrichment of the ER-resident protein Sec 61p versus Pma1p (a canonical marker of plasma membranes) in crude homogenate and in the microsomal fractions. The relative amounts of each protein were estimated by western blotting using polyclonal antibodies against Sec 61p (Dr Randy Schekman; University of California, USA) and Pma1p (Dr Carolyn Slayman; Yale University, USA) in 10 and 100 μg protein samples, with the ImageJ 1.40 g software (NIH).

Lipid analysis and mass spectrometry

Lipid extracts were prepared from ≅2 × 109 (sterols), or 108 (phospholipids) yeast cells grown as indicated, or approximately 1 mg (sterols) and 200 μg (phospholipids) of proteins from microsomal fractions. Cells were harvested, washed with distilled water, suspended in 1 mL of cold water and shock-frozen. They were broken by vigorous shaking with 500 μL of glass beads (diameter 0.3–0.4 mm; Sigma) using a mini-beadbeaterTM (Biospec Products) for 1 min at 5000 rev/min. Cellular lipids were extracted using chloroform/methanol (2:1, v:v) as described by Folch et al. (57). The final organic phase was evaporated and lipids were dissolved either in 100 μL hexane (sterols) or 200 μL chloroform/methanol/H2O (16:16:5, v:v:v) (phospholipids).

For specific sterol identification and quantification, total lipid extracts were saponified as previously reported (17). The different sterol species were then separated by gas chromatography using a 25 m ×0.32 mm AT-1 capillary column (Alltech), and identified by means of their retention times relative to cholesterol as a standard. The results are expressed as μg of sterol per 109 cells.

To analyze phospholipid species by MS, total lipid extracts were reconstituted at a concentration of approximately 50 μg.mL−1 of phospholipids in chloroform/methanol/H2O (16:16:5, v:v:v), with 1% (v/v) formic acid for analysis in the positive ion mode. For routine single-stage MS, samples were analyzed with a triple quadrupole instrument model API 165 (Perkin Elmer Sciex) equipped with an ion-spray source. Analysis of spectra was performed using the Biomultiview 1.2 software (Perkin Elmer Sciex) and raw identification of phospholipid species was carried out based on their expected m/z by means of a home-made software program. The various species were unambiguously identified by tandem MS with a Deca XP Max equipped with an ion trap source (Thermo Electron) by precursor ion scan analysis, as previously described (58). The molecular profile of PC species was specifically obtained by scanning for the positive ion precursors of m/z 184, characteristic of choline phosphate.

Fluorescence microscopy

For Pma1p immunolocalization studies, a HA-tagged version of PMA1 was placed under control of a heat-shock promoter in the integrative plasmid YIplac204 (TRP1; Yiplac204-2HSEpr-HA-PMA1; 54). As desired, expression of the tagged PMA1 gene was induced by incubation of the cells for 15 min at 39°C (54). HA-tagged Pma1p was visualized by immunofluorescence as previously described (22). The primary antibodies used were HA.11 monoclonal 16B12 from raw ascites fluid (Babco), diluted 1:150. Goat anti-mouse Texas Red IgG (Jackson Immunoresearch) served as fluorescent secondary antibodies and were diluted 1:100. Samples were examined by confocal laser scanning microscopy using a Bio-Rad MRC 1024 equipped with a 15-mW argon-krypton gas laser. The confocal unit was attached to an inverted microscope (Olympus IX70). Images were obtained with Olympus plan apo x60 water, 1.3 numerical aperture objective lens. Fluorescence signal output was measured using the control software (Lasersharp 3.2; Bio-Rad). The Texas Red fluorochrome was excited with the 568 nm yellow line and emission of the dye was collected via a photomultiplier through a 605-nm pass band filter (35 nm width).

Invertase secretion assays

The invertase secretion assays were performed according to the method described by Munn et al. (59). Yeast cells were grown to an OD600 of 0.2–0.5 in the indicated medium containing 2% glucose. After washing, 10 OD600 units of cells were induced for invertase expression by resuspension in selective medium containing 0.05% glucose and 2% sucrose. Cell samples were taken at 0, 15, 30, 45 and 60 min after transfer to low glucose medium. Invertase activity was determined as described by Munn et al. (59), using an enzyme reporter assay. Intracellular invertase activity was calculated by subtracting extracellular activity, measured using unbroken cells, from total invertase activity determined after permeabilization of the cells by freezing in liquid nitrogen in the presence of 10% Triton X-100.

Electron microscopy

hem1Δ cells were fixed with 2% paraformaldehyde (m/v) and 0.5% glutaraldehyde (v/v) in 0.1 M phosphate buffer (pH 6.8) for 1 h at 25°C. The samples were postfixed in 1% (v/v) osmium tetroxide for 15 min at 4°C, dehydrated with alcohol and embedded in LR White resin. Polymerization was achieved at 4°C under UV light. Ultrathin sections were stained with uranyl acetate and lead citrate and observed using a JEOL 100 S microscope under 80 kV.

Laurdan and DPH labeling of microsomal membranes

The concentration of microsomal preparations was adjusted to 140 μg.mL−1 in GTH buffer. Laurdan (Sigma) was prepared as a 2 mM stock solution in dimethyl sulphoxide (DMSO). Three millilitres of microsomal preparations in GTH buffer were transferred to a quartz cuvette before labelling with 2 μL of the Laurdan stock solution in the dark at 27°C for 10 min to allow for an appropriate level of staining of the microsomes. A Fluorolog-3 spectrometer (Jobin Yvon, Horiba group, USA) was used for the fluorescence measurements. During measurements, samples were stirred and equilibrated in a temperature-controlled chamber using a thermoelectric Peltier junction. The excitation wavelength used during this study was 350 nm and emission spectra were acquired between 420 and 550 nm at 7 and 27°C. Blank spectra were obtained with unlabeled microsomes and subtracted from the spectra of labelled microsomes.

Membrane fluidity was assessed by the steady-state fluorescence polarization of 1,6-DPH (60). DPH (Sigma) was prepared as a 1 mM stock solution in tetrahydrofuran (Sigma). Before labeling microsomal membranes, DPH was dispersed by injection of 4 μL from the stock solution into 2 mL of GTH buffer. After 15 min stirring, 1 mL of microsomes in GTH buffer (400 μg.mL−1) was added to the suspension before incubation for 30 min at 27°C. Steady-state anisotropy of DPH was measured in a Fluorolog-3 spectrometer (Jobin-Yvon, Horiba Group, USA), using T-format fluorescence polarizers. The excitation and emission wavelengths were 360 nm (5 nm bandwidth) and 431 nm (5 nm bandwidth), respectively. Steady-state fluorescence anisotropy (r) was calculated as:


Where I stands for the fluorescence intensity and the first and second subscripts refer to the setting of the excitation and emission polarisers, respectively. G = Ihv/Ihh is a correction factor for the monochromator's transmission efficiency for vertically and horizontally polarized light.


We are very grateful to Daniel Guyonnet for excellent technical assistance with electrospray mass spectrometry and Dr Anne Cantereau (Confocal Microscopy facility of UMR CNRS 6187, Poitiers) for her precious advice regarding confocal microscopy imaging. Florence Thibault, Jean-Michel Pérault and Emile Béré (Service Interdisciplinaire de Microscopie et d'Imagerie Scientifique, Université de Poitiers) are acknowledged for their worthy help with sample preparation and image processing regarding electron microscopy experiments. We thank Dr Peter Walter (University of California, USA), Dr Kenji Kohno (Nara Institute of Science and Technology, Ikoma, Japan), Dr Carolyn Slayman (Yale University, USA) and DrRandy Schekman (University of California, USA) for strains, plasmids and antibodies. Miss Lucy Mills is acknowledged for her help in writing this article in English and Dr Alan Brett Mason (Yale University, USA) for his precious comments and for proofreading the manuscript. This work was supported by the French MENRT (with a grant to LP), the Région Poitou-Charente and the CNRS.