Degradation of Misfolded Proteins Prevents ER-Derived Oxidative Stress and Cell Death

Molecular Cell, Vol. 15, 767–776, September 10, 2004, Copyright 2004 by Cell Press

Degradation of Misfolded Proteins Prevents ER-Derived Oxidative Stress and Cell Death Cole M. Haynes, Eric A. Titus, and Antony A. Cooper* Division of Cell Biology and Biophysics School of Biological Sciences University of Missouri-Kansas City Kansas City, Missouri 64110

Summary A variety of debilitating diseases including diabetes, Alzheimer’s, Huntington’s, Parkinson’s, and prionbased diseases are linked to stress within the endoplasmic reticulum (ER). Using S. cerevisiae, we sought to determine the relationship between protein misfolding, ER stress, and cell death. In the absence of ERV29, a stress-induced gene required for ER associated degradation (ERAD), misfolded proteins accumulate in the ER leading to persistent ER stress and subsequent cell death. Cells alleviate ER stress through the unfolded protein response (UPR); however, if stress is sustained the UPR contributes to cell death by causing the accumulation of reactive oxygen species (ROS). ROS are generated from two sources: the UPR-regulated oxidative folding machinery in the ER and mitochondria. Our results demonstrate a direct mechanism(s) by which misfolded proteins lead to cellular damage and death. Introduction Cells have evolved a number of mechanisms to maintain the proper folding and thus functionality of important protein machinery. However, the complexities of protein folding coupled with environmental conditions that perturb folding constantly threaten the vitality of cells. Protein aggregation or alternative conformations coupled with a gain of function are common to a number of debilitating adult-onset diseases including Alzheimer’s, Parkinson’s, Huntington’s, and the prion-based diseases. To maintain proteins in a properly folded state, cells utilize a variety of chaperones that facilitate refolding, and if a protein is unable to be refolded it is degraded, often via the ubiquitin-proteasome system. If any aspect of the cellular stress responses is compromised, the potential for cellular damage increases. An organelle particularly susceptible to protein misfolding is the endoplasmic reticulum (ER), the first organelle of the secretory pathway. Once proteins have entered the ER, protein folding followed by glycosylation and disulfide bond formation commences. Proteins unable to fold correctly cause ER stress and activate the unfolded protein response (UPR). Activation of the UPR transcriptionally upregulates an array of genes required for protein folding, ER expansion, ER-Golgi trafficking, and ERAD, which all act collectively to relieve stress within the ER (Travers et al., 2000). To ensure proper processing of proteins within the ER, eukaryotic cells *Correspondence: [email protected]

employ a quality control mechanism that recognizes and degrades (ERAD) aberrantly folded proteins to prevent the aggregation and/or delivery of potentially dysfunctional or cytotoxic proteins (Plemper and Wolf, 1999). At least two ERAD systems are employed by S. cerevisiae to eliminate misfolded proteins, underscoring the importance of preventing the accumulation of misfolded proteins in the ER. HRD-dependent ERAD appears saturable as disruption of its components (HRD1, HRD3, or DER1) has little to no effect on the degradation of a model misfolded protein when highly expressed, while conversely, HIP-dependent ERAD is essential for the degradation of elevated levels of this misfolded protein (Haynes et al., 2002). The HIP pathway therefore acts as a high-capacity mechanism to accommodate increased levels of substrates. The fact that degradation of a highly expressed misfolded protein is completely dependent on the HIP pathway presented a unique opportunity to investigate the cellular consequences of the accumulation of misfolded proteins within the ER. Elevated expression of misfolded proteins in HIP-deficient cells would result in accumulation of such proteins in the ER and potentially mimic the ER stress associated with diseases such as diabetes, Alzheimer’s, Parkinson’s, Huntington’s, and the prion-based diseases. In this work, we show that the accumulation of a misfolded protein within the lumen of the ER leads to prolonged UPR activation, which in turn causes oxidative stress and finally cell death. Prolonged UPR activation leads to the accumulation of reactive oxygen species (ROS) via two sources, the UPR-regulated oxidative protein folding machinery in the ER and the mitochondria. These data highlight the essential nature of the secretory pathway, the measures cells must employ to maintain its functionality, and the consequences of prolonged ER stress. Results ER stress has been linked to a number of debilitating diseases. To determine how cells respond to such stress, we utilized HIP-deficient cells (erv29⌬) that lack the ability to eliminate misfolded proteins from the ER (Haynes et al., 2002). Low levels of the misfolded protein CPY* (CPY*-1x, a mutant misfolded form of the vacuolar protein carboxypeptidase Y) did not affect the growth of HIP-deficient cells (Figure 1A); however, increasing the number of misfolded proteins in addition to CPY* through incubation at an elevated temperature caused growth inhibition of HIP-deficient cells expressing CPY*1x (Figure 1A). Growth inhibition at 38⬚C was seen only in HIP-deficient cells expressing CPY*-1x, whereas cells lacking the HRD/DER degradation pathway (hrd1⌬) showed no growth defects. These results suggest that the ER of HIP-deficient cells expressing CPY*-1x at 30⬚C is near capacity in terms of the quantity of misfolded proteins it can tolerate as any further increase in misfolded proteins inhibited cell growth. To examine the consequences of the accumulation of a single misfolded

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Figure 1. HIP-Deficient Cells Are Sensitive to the Accumulation of Misfolded Proteins in the ER (A) Wild-type (KHY163), hrd1⌬ (KHY171), and erv29⌬ (KHY270) cells expressing CPY*-1x and erv29⌬ (KHY271) cells expressing wtCPY1x were serially diluted, spotted onto plates, and grown at 30⬚C or 38⬚C. (B) Western blot analysis of CPY* quantities in wild-type cells. 1x, CPY* expressed from the native promoter; 8x, CPY* expressed from the native promoter on a high copy (2 ␮) plasmid; 25x, CPY* expressed from the CUP1 promoter in the presence of 150 ␮M CuSO4. (C) A growth curve of wild-type, hrd1⌬, and erv29⌬ cells expressing CPY*-25x (pAC595) following the addition of CuSO4. (D) Wild-type and erv29⌬ cells, expressing either CPY*-1x or CPY*-25x, were grown at 30⬚C or 38⬚C for 3 hr and UPR activation levels measured as described in Experimental Procedures. As a control for full UPR activation, wild-type cells were grown in 3 mM DTT for 3 hr. (E) CPY*-25x expression was induced in wildtype and erv29⌬ cells harboring pBG15 (CPY*1x-HA, Caldwell et al., 2001). Cells were then harvested and fixed for indirect immunofluorescence at the times indicated. Cells were stained with anti-HA antibodies in combination with AlexaFluor 488 to detect CPY*-1xHA, anti-Eug1p antibodies in combination with AlexaFluor 568 to detect the ER luminal marker, Eug1p, and DAPI to detect nuclei. Cell morphology was observed through differential interference contrast (DIC). At 4 hr, the fluorescence signal was overexposed to detect the diminishing signal.

protein in the ER, CPY* was expressed from the Cu2⫹ inducible CUP1 promoter, which elevated expression approximately 25-fold (CPY*-25x, Figure 1B). The growth rate of wild-type or hrd1⌬ cells expressing CPY*25x was slightly impaired, while cells lacking the HIP pathway failed to grow (Figure 1C). HIP-deficient cells are therefore sensitive to protein misfolding in the ER and fail to grow when high levels of misfolded proteins accumulate. The UPR upregulates an array of genes that act to relieve stress within the ER (Travers et al., 2000). The extent of UPR activation is tailored to the degree of stress and therefore can be used as a relative measure of ER stress. HIP-deficient cells expressing either CPY*1x at 38⬚C or CPY*-25x at 30⬚C showed a striking degree of UPR activation, nearly 9-fold greater than that in wildtype cells (Figure 1D) and was similar to that of wildtype cells grown in 3 mM DTT, a reducing agent known to fully activate the UPR (Friedlander et al., 2000). Therefore, severe ER stress, as indicated by high UPR activation, correlates with reduced growth rates. To understand the relationship between ER stress and growth inhibition, we examined the subcellullar localization of CPY*. CPY*-25x expression was induced in HIPdeficient cells constitutively expressing epitope-tagged CPY* (CPY*-HA-1x), and cells were fixed at different times. HA-tagged CPY* was observed by indirect immunofluorescence, and the signal compared to that of an

ER-resident protein, Eug1p (Tachibana and Stevens, 1992) (Figure 1E). As expected, CPY*-HA localized to the ER prior to induction of CPY*-25x; however, after 2–3 hr of induction, CPY*-HA was observed only at the cell periphery in the cortical ER and was no longer localized as a perinuclear ring. By 3–4 hr, the cortical ER signal had also diminished. The loss of perinuclear CPY*-HA staining at 2–3 hr coincided with the loss of perinuclear signal from the ER marker protein. Cells were also stained with DAPI to view DNA, and intriguingly after 4 hr, over 50% of the HIP-deficient cells expressing CPY*-25x displayed a completely delocalized DAPI signal, suggesting that the nuclear DNA had dispersed. This loss of organelle and nuclear structure was only observed in cells with severe ER stress as the ER and nucleus remained intact in wild-type cells expressing CPY*-25x. ER Stress Causes Cell Death Dependent on the UPR Expression of CPY*-25x in HIP-deficient cells causes UPR activation, the loss of ER and nuclear integrity, and growth inhibition. The possibility that sustained ER stress is lethal was investigated using a survivorship assay. Figure 2A shows that erv29⌬ cells began to die after 2–3 hr of CPY*-25x induction, with few cells surviving after 24 hr, whereas wild-type cells expressing CPY*25x were relatively unaffected. Figure 2B shows that the level of UPR activation in the erv29⌬ cells was fully

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investigated the role in cell death of two signaling components of the UPR: Ire1p, the ER stress signal transducer, and Hac1p, the downstream transcription factor (Chapman et al., 1998). Both IRE1 and HAC1 were disrupted in erv29⌬ cells expressing CPY*-25x, and the effect on growth rate determined. Figure 2C shows growth curves of wild-type, erv29⌬, erv29⌬ ire1⌬, and erv29⌬ hac1⌬ cells expressing CPY*-25x. Elimination of the UPR permitted erv29⌬ cells to grow, demonstrating that UPR signaling via Ire1p and Hac1p contributes to the loss of viability following the accumulation of a misfolded protein in the ER. These cells grew somewhat slower than wild-type cells, but this was not unexpected as the cells remain under ER stress due to the inability to degrade CPY* and yet cannot upregulate the complement of chaperones and degradative machinery under UPR control. To confirm erv29⌬ ire1⌬ and erv29⌬ hac1⌬ cells were unable to activate the UPR, UPR activation assays were performed following exposure to 3 mM DTT. These cells exhibited no UPR activation (see Supplemental Figure S1 at http://www.molecule.org/cgi/content/ full/15/5/767/DC1/).

Figure 2. CPY*-25x Expression in HIP-Deficient Cells Leads to Cell Death that Is Dependent on UPR Activation (A) A survivorship curve in which wild-type and erv29⌬ cells were grown in liquid for the indicated amount of time (x axis) following induction of CPY*-25x expression. Cell survivorship was determined by growing wild-type (KHY163) and erv29⌬ (KHY270) cells overnight in synthetic media to log phase (30⬚C) followed by the addition of 150 ␮M CuSO4 to induce expression of CPY*-25x (pAC595). To maintain wild-type cells in log phase, several dilutions were made throughout the time course. At the described times, 1 OD600nm was harvested, diluted 1:1000, and 300 ␮l of these cells were plated on synthetic media lacking CuSO4 and incubated at 30⬚C. Colonyforming units were then determined and a survivorship curve generated. (B) UPR activation assays of wild-type and erv29⌬ cells following induction of CPY*-25x expression. As a control for UPR activation, wild-type cells were grown in 3 mM DTT for 3 hr. (C) A growth curve comparing wild-type, erv29⌬, erv29⌬ ire1⌬ (KHY438), and erv29⌬ hac1⌬ (KHY439) cells following induction of CPY*-25x expression.

elevated after 1–2 hr of CPY*-25x induction, indicating that UPR activation preceded cell death. The activation of the UPR prior to cell death suggested that in the presence of high quantities of misfolded proteins the UPR may contribute to cell death. We therefore

ER Stress Causes Programmed Cell Death-like Features Our results indicate that HIP-deficient cells expressing CPY*-25x exhibit UPR activation, loss of DNA staining, and subsequent death. Loss of nuclear DNA in conjunction with cell death is consistent with programmed cell death (PCD) (Wyllie et al., 1980). Two hallmarks of PCD are DNA fragmentation and the externalization of phosphatidylserine (PS) on the plasma membrane (Martin et al., 1995). We examined wild-type and HIP-deficient cells for DNA fragmentation with TUNEL assays. After 4 hr of CPY*-25x expression, erv29⌬ cells showed extensive DNA fragmentation, while wild-type cells did not (Figure 3A). Detection of TUNEL-positive cells correlated with the timing of the loss of both nuclear/ER structure (Figure 1E) and cell viability (Figure 2A). Wild-type and erv29⌬ cells expressing CPY*-25x were also examined for PS exposure on the plasma membrane. Two hours after CPY*-25x induction, erv29⌬ but not wild-type cells bound fluorescently labeled annexin V, indicating that PS was now exposed extracellularly (Figure 3B). While DNA fragmentation and PS exposure are hallmarks of PCD, we have yet to identify any known regulatory proteins such as caspases that affect this process in S. cerevisiae, including the yeast metacaspase YCA1 (Madeo et al., 2002) whose absence had no effect in our studies (see Supplemental Figure S2 at http://www. molecule.org/cgi/content/full/15/5/767/DC1/). Therefore we cannot eliminate necrosis. We also examined the potential role of the UPR in the appearance of PCD-like phenotypes by TUNEL assays performed over a time course following CPY*-25x expression. Figure 3C shows the percentage of TUNEL staining over 6 hr in strains expressing CPY*-25x. erv29⌬ cells display TUNEL staining at 3–4 hr, while erv29⌬ cells lacking a functional UPR (erv29 ire1⌬ or erv29⌬ hac1⌬) show no greater percentage of DNA fragmentation than wild-type cells, demonstrating that the UPR is required for DNA fragmentation.

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Figure 3. ER Stress Causes Cells to Display Some Hallmarks of Programmed Cell Death TUNEL and annexin V assays (see Experimental Procedures) comparing wild-type (KHY163), erv29⌬ (KHY270), erv29⌬ ire1⌬ (KHY438), and erv29⌬ hac1⌬ (KHY439) cells harboring pAC595 to express CPY*-25x. (A) TUNEL assays to observe DNA fragmentation in wild-type and erv29⌬ cells following induction of CPY*-25x for the indicated times. These cells did not appear to be necrotic as less than 5% of similarly treated cells did not stain with propidium iodide, indicating that the integrity of the plasma membrane was preserved (Madeo et al., 1997a). (B) Annexin V assays (Madeo et al., 1997b) to examine PS externalization on the plasma membrane of wild-type and erv29⌬ cells harvested following induction of CPY*-25x for the indicated times. (C) DNA fragmentation in wild-type, erv29⌬, erv29⌬ ire1⌬, and erv29⌬ hac1⌬ cells following induction of CPY*-25x. Cells were harvested at the indicated time points and TUNEL assays performed. At least 300 cells were counted to determine the percentage of TUNEL-positive cells at each time point.

ER Stress Leads to Accumulation of Reactive Oxygen Species Our data indicating that the UPR contributes to cell death during prolonged ER stress suggested the existence of a downstream event dependent on Ire1p. Two recent reports have linked ER metabolism with the generation of reactive oxygen species (ROS) through oxidative protein folding (Harding et al., 2003; Tu and Weissman, 2004). Intriguingly, ER components involved in this process are regulated by the UPR (Travers et al., 2000).

To test for the presence of ROS, we utilized ROS-sensitive fluorescent dyes, including dihydroxyrhodamine 123. Staining of both wild-type and erv29⌬-deficient cells following 1.5 hr of CPY*-25x expression found that erv29⌬ but not wild-type cells accumulated ROS (Figure 4A). Based on the sequence of observed events, we propose the following timeline: CPY* accumulation→ER stress→UPR activation→ROS generation→organelle breakdown and DNA fragmentation→cell death. In this model sustained ER stress causes prolonged UPR activation resulting in the production of ROS. To test this model, the presence of ROS in erv29⌬ ire1⌬ and erv29⌬ hac1⌬ cells was examined following expression of CPY*-25x. Figure 4B shows that cells unable to activate the UPR did not accumulate ROS despite the presence of ER stress. To determine the importance of ROS in cell death, erv29⌬ cells expressing CPY*-25x were treated in a manner that prevented ROS accumulation while not diminishing ER stress. To relieve oxidative stress, cells utilize a mechanism to increase the concentration of reduced glutathione (GSH), which acts as a reductant of endogenous peroxides (Hayes and McLellan, 1999). If ROS accumulation is the cause of cell death under these conditions, then ROS elimination via increased GSH levels should prevent death. Figure 4C shows growth curves of erv29⌬ cells expressing CPY*-25x in the presence or absence of 1 mM GSH. As previously demonstrated, growth of erv29⌬ cells expressing CPY*25x is inhibited. However, following 1 hr of CPY*-25x induction, the addition of 1 mM GSH permitted the continued growth of these cells. Figure 4D shows that ROS cannot be detected in erv29⌬ cells expressing CPY*25x following exposure to 1 mM GSH for 2 hr. Addition of 1 mM GSH did not diminish UPR activation in these cells (see Supplemental Figure S3 at http://www. molecule.org/cgi/content/full/15/5/767/DC1/). Oxidative Folding in the ER Contributes to ROS Production and Accumulation Due to prolonged UPR activation, ROS accumulate during ER stress and lead to cell death. The ER may directly contribute to ROS production through the oxidation and reduction of disulfide bonds (Tu and Weissman, 2004). During disulfide bond formation electrons are passed, through a series of thiol-disulfide exchange reactions, from the thiols of the substrate protein to protein disulfide isomerase (Pdi1p) then to Ero1p and finally to molecular oxygen (Tu and Weissman, 2002). The reduction of molecular oxygen would likely result in ROS as a byproduct (Figure 5A). The upregulation of both Ero1p and Pdi1p by the UPR (Travers et al., 2000) could provide the basis for the observed accumulation of ROS upon prolonged UPR activation. This model predicts that overexpression of ERO1 would cause an increase in ROS accumulation in the absence of UPR activation, and/or removal of Ero1p would inhibit ROS generation. To determine if ERO1 overexpression, uncoupled from UPR regulated expression, could contribute to ROS accumulation, ERO1 was placed on a high-copy plasmid. Figure 5B shows flow cytometry analysis of erv29⌬ ire1⌬ cells expressing CPY*-25x with and without overex-

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Figure 5. Oxidative Protein Folding in the ER Contributes to ROS Accumulation and Cell Death (A) A depiction of disulfide bond oxidation and reduction in the lumen of the ER indicating the flow of electrons and oxidizing equivalents (Cuozzo and Kaiser, 1999). (B) Flow cytometry analysis of DH123-stained erv29⌬ ire1⌬ (KHY438) cells expressing CPY*-25x (pAC595) in the presence (black line) or absence (gray line) of ERO1 overexpression (pAC672). Flow cytometry was performed as described (Madeo et al., 1999). (C) Growth curves comparing wild-type (KHY298) and erv29⌬ (KHY318) cells expressing CPY*-25x or cysteine-less CPY*-25x (pAC774).

Figure 4. ER Stress and Prolonged UPR Activation Leads to ROS Accumulation (A) Fluorescence microscopy comparing DH123 stained wild-type (KHY163) and erv29⌬ (KHY270) cells following expression of CPY*25x for the indicated amount of time. Cells were stained with DH123 for 2 hr then visualized using filter sets specific for rhodamine and DIC to observe cell morphology. (B) Fluorescence microscopy comparing DH123 stained erv29⌬, erv29⌬ ire1⌬ (KHY438), and erv29⌬ hac1⌬ (KHY439) following expression of CPY*-25x for 2 hr. (C) A growth curve comparing erv29⌬ cells expressing CPY*-25x incubated with 0 or 1 mM GSH. GSH was added 1 hr after induction of CPY*-25x. (D) Fluorescence microscopy of DH123 stained erv29⌬ cells following addition of 1 mM GSH. The cells were induced to express CPY*25x, an aliquot of cells was harvested at 1 hr, and 1 mM GSH was added to the remaining cells which were harvested 2 hr later and stained with DH123.

pressed ERO1 stained for ROS. Overexpression of ERO1 caused a significant increase in ROS, suggesting that upregulation of the oxidative protein folding machinery by the UPR contributes to ROS accumulation. Our ability to test if the removal of Ero1p would inhibit ROS generation has proven impossible due to a strong synthetic interaction between the temperature-sensitive ero1-1 allele (Frand and Kaiser, 1999) and CPY*. Even at the permissive temperature the erv29⌬ ero1-1 cells expressing CPY*-1x were severely retarded for growth (see Supplemental Figure S4 at http://www.molecule. org/cgi/content/full/15/5/767/DC1/), and transformation of the CPY*-25x plasmid was never successful. As ERO1 is an essential gene and necessary for disulfide bond formation in a number of proteins, we predict this phenotype is due to protein folding and secretion defects in the ER. We suspect that the reduced level of Ero1p activity available for forming disulfide bonds in ero1-1 cells, even at the permissive temperature, is limiting, and the repeated interactions of CPY* with Pdi1p (see

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below) effectively depletes the cell of Ero1p activity available for forming essential disulfide bonds in other proteins. Our model predicts that thiol oxidation of cysteines and subsequent disulfide bond formation in CPY* by the UPR-regulated ER oxidative folding machinery leads to ROS generation and cell death. To directly test the importance of thiol oxidation and disulfide bond formation, we mutated all 11 cysteine residues in CPY* to alanines and determined the effect on growth. Figure 5C shows growth curves comparing wild-type and erv29⌬ strains following induction of CPY*-25x or cysteine-less CPY*-25x (cys-less CPY*-25x). The lack of cysteines and thus disulfide bonds in CPY* allowed growth of the erv29⌬ cells. The growth difference between the cysless-CPY*-25x and CPY*-25x is lost when the cells are grown in the presence of GSH (data not shown). Cysless CPY*-25x is expressed to the same level as CPY*25x and is degraded at the same rate as CPY*-25x (see Supplemental Figure S5 at http://www.molecule.org/ cgi/content/full/15/5/767/DC1/) indicating that the difference in growth rate was due solely to the lack of cysteines. While ER oxidative protein folding may directly generate ROS, oxidative folding may also contribute to ROS accumulation through depletion of GSH (see below; Cuozzo and Kaiser, 1999). Elimination of ROS through the oxidation of GSH would be expected to increase the ratio of oxidized glutathione (GSSG) to GSH. This ratio, along with total glutathione, was examined in wild-type cells and erv29⌬ cells following induction of CPY*-25x (Figure 6A). Prior to induction of CPY*-25x, 10-fold more total glutathione existed in erv29⌬ cells relative to wildtype cells (see Supplemental Figure S6 at http://www. molecule.org/cgi/content/full/15/5/767/DC1/). The low level of CPY* (1x) constitutively expressed in preinduced erv29⌬ cells is not growth inhibitory (Figure 1A) but is likely causing a degree of oxidative stress that the cells tolerate by significantly upregulating GSH synthesis. Upon induction of CPY*-25x in erv29⌬ cells, GSH is quickly oxidized as reflected by an increase in the GSSG/GSH ratio. The basis of GSH oxidation is unlikely due to neutralizing ROS produced from de novo disulfide bond formation as overexpression of correctly folded

Figure 6. Oxidation of Glutathione Is Associated with ER Stress and Prolonged Interactions between CPY* and Pdi1p (A) Quantification of GSSG/GSH following induction of CPY*-25x in erv29⌬ (KHY270) cells compared to wild-type cells (SEY6211a). Quantification was performed as described (Cuozzo and Kaiser, 1999). (B) Autoradiograph and quantification of Pdi1p interactions with either wtCPY-25x or CPY*-25x via mixed disulfide bond. erv29⌬ cells containing pAC595 or erv29⌬ (KHY271) containing pAC667 were incubated in 150 ␮M CuSO4 for 30 min followed by radiolabeling for 10 min, chase for 2 min, cycloheximide (50 ␮g/ml) addition for 2 min, and cells were harvested at the described time points. The top panel shows the amount of CPY* or wtCPY bound to Pdi1p via a mixed disulfide. Isolation and quantification of mixed disulfides was performed as described (Frand and Kaiser, 1999) (see Experimental Procedures). In short, cells were harvested, suspended in 10% TCA to block thiol exchange, lysed in a buffer containing SDS under nonreducing conditions, divided into four aliquots, and Pdi1p was immunoprecipitated under nonreducing conditions from one of

the aliquots. The samples were then exposed to 100 mM DTT to break disulfide bonds, diluted, and incubated with anti-CPY antibodies to immunoprecipitate the wtCPY/CPY* that interacted with Pdi1p via mixed disulfide bond. The samples were run on a reducing 10% SDS page gel. The remaining three aliquots which represent 1/7 the cell lysate used in the top panel were immunoprecipitated under reducing conditions for the indicated antigens, and they represent the total relative amount of each protein. Vma1p was used as a loading/lysis control. The graph represents the percentage of wtCPY or CPY* in a cell that is bound to Pdi1p via mixed disulfide bond at the indicated time following the chase. (C) Fluorescence microscopy comparing DH123 staining in erv29⌬ cells expressing CPY*-25x and erv29⌬ cells expressing wtCPY-25x for 2 hr. (D) Growth curves comparing CPY*-25x expression to that of wtCPY-25x in wild-type (SEY6211a) and erv29⌬ (KHY271) cells. The PRC1 ORF encoding wtCPY was inserted downstream of the CUP1 promoter so the expression level would be identical to that of CPY*25x (data not shown).

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wtCPY (wtCPY-25x), which contains five disulfide bonds, was not growth inhibitory in erv29⌬ cells (Figure 6D). Instead the rapid oxidation of GSH may result from GSH acting directly to reduce improper disulfide bonds within misfolded CPY* (Figure 5A). This results in the consumption of GSH and would return the thiols of CPY* to the reduced form allowing them to again interact with Ero1pPdi1p to be reoxidized. This futile cycle of disulfide formation and breakage would continue with each cycle generating ROS and consuming GSH (Figure 5A). Our model of GSH depletion and ROS accumulation through the actions of Ero1p-Pdi1p predicts multiple interactions between CPY* and Pdi1p. It has been shown previously that wtCPY interacts with Pdi1p via mixed disulfides and is required for wtCPY folding and export (Frand and Kaiser, 1999). To examine differences between the mixed-disulfide interactions of wtCPY or CPY* and Pdi1p, erv29⌬ cells expressing either wtCPY-25x or CPY*-25x were radiolabeled, and the amount of CPY*/ wtCPY interacting with Pdi1p through mixed disulfide bond(s) was determined over time. Figure 6B shows that (1) the amount of CPY* interacting with Pdi1p is much greater than that of wtCPY and (2) that CPY* interacts much longer with Pdi1p than wtCPY. At the time point immediately following the chase, 5-fold more CPY* interacts with Pdi1p via a mixed disulfide bond, while 30 min later there is 14-fold more CPY* interacting with Pdi1p than wtCPY. These observations match our predictions of multiple, repeated interactions between CPY* and Pdi1p. The shorter, more limited, interactions between wtCPY and Pdi1p in erv29⌬ cells expressing wtCPY25x did not result in ROS accumulation (Figure 6C) nor inhibit growth (Figure 6D).

Mitochondria Contribute to ROS Accumulation Respiring mitochondria are known producers of ROS and could potentially contribute to ROS that accumulate during ER stress. The above experiments were performed in media containing high concentrations of glucose where mitochondrial respiration is reduced (Fiechter et al., 1981). However, ROS accumulation can occur under conditions of reduced mitochondrial activity, and mitochondrial respiration is a major source of ROS in yeast cells (Richter et al., 1995). To test if mitochondria are an additional source of ROS during ER stress, erv29⌬ cells were converted to ␳0 cells, which lack mitochondrial DNA and are therefore devoid of respiring mitochondria (Davermann et al., 2002). Respiration-deficient (␳0) wild-type and erv29⌬ cells expressing CPY*-25x were examined for the production of ROS and growth. ␳0 erv29⌬ cells did not accumulate detectable levels of ROS (Figure 7A) and were able to grow as well as ␳0 wild-type cells (Figure 7B), suggesting that respiring mitochondria contribute to ROS accumulation and cell death following ER stress. Figure 7C shows that both ␳0 wild-type and erv29⌬ cells expressing CPY*-25x did not show DNA fragmentation and confirmed that mitochondria and respiration contribute to PCD-like phenotypes. It should be noted that respiration deficiency does not affect the level of UPR activation in these cells (see Supplemental Figure S7 at http://www. molecule.org/cgi/content/full/15/5/767/DC1/).

Figure 7. Respiring Mitochondria Contribute to ROS Accumulation and Cell Death during ER Stress All cells were grown in synthetic media containing 2% glycerol and 0.5% glucose. (A) Fluorescence microscopy of ␳0-derived wild-type (KHY163) and ␳0-derived erv29⌬ (KHY270) cells stained with dihydroethidium for 10 min following 1.5 hr of CPY*-25x (pAC595) expression (Madeo et al., 1999). (B) Growth curves of ␳0-derived wild-type and ␳0-derived erv29⌬ cells following induction of CPY*-25x expression. (C) TUNEL assays comparing ␳0-derived wild-type and ␳0-derived erv29⌬ cells following CPY*-25x expression for 5 hr.

Discussion This work examines the consequences of prolonged ER stress and demonstrates that the accumulation of an aberrantly folded protein within the ER lumen can lead directly to cell death. Cells deficient in the HIP pathway, a component of ERAD upregulated in response to high quantities of misfolded proteins (Haynes et al., 2002), are unable to survive the cellular consequences of misfolded CPY* accumulation. Oxidative Protein Folding Derived ROS and Glutathione Depletion Our results support a model in which the ultimate phenotype of sustained, severe ER stress is cell death due to prolonged UPR activation and the subsequent accumulation of ROS. ROS accumulate because of events from at least two sources: the oxidative folding machinery in the ER and mitochondria. In the ER, two mechanisms likely contribute to ROS accumulation with Ero1p and Pdi1p playing a central role in both. During the formation of disulfide bonds, Ero1p and Pdi1p act in concert to transfer electrons from the thiol groups of substrate proteins to molecular oxygen with the generation of ROS as a byproduct (Tu and Weissman, 2004). An additional

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process leading to ROS accumulation derives from the proposed role of glutathione to directly reduce unstable and improper disulfide bonds (Cuozzo and Kaiser, 1999), such as those likely to occur in misfolded CPY* but not in wtCPY. Reduction of disulfides in CPY* by GSH has two consequences: the depletion of cellular GSH diminishes the ability of the cell to eliminate ROS, and it restores thiol groups to again interact with Ero1p-Pdi1p in a manner that generates ROS. This cycle of disulfide bond formation and reduction (Figure 5A) would be repeated indefinitely as CPY* degradation is impaired in erv29⌬ cells. The importance of cysteine residues and disulfide bond oxidation in this process is illustrated by the significantly improved growth of erv29⌬ cells when the cysteines of CPY* are replaced with alanines (Figure 5C). Misfolded cysteine-less CPY*-25x continues to cause ER stress and is therefore somewhat growth inhibitory relative to correctly folded wtCPY-25x. The combination of ROS generation and GSH depletion act synergistically to increase the net total of ROS. Correctly folded wtCPY likely forms stable disulfide bonds relatively immune to reduction by GSH and therefore fails to enter this cycle. Increased Ero1p expression results in both enhanced ROS levels (Figure 5B) and a decrease in GSH (Cuozzo and Kaiser, 1999). Therefore the 5- to 8-fold upregulation of Ero1p and Pdi1p by the activated UPR (Travers et al., 2000) would greatly exacerbate the oxidative stress in the ER by further increasing the rate of ROS production and GSH expenditure to reduce improper disulfides. The significance of prolonged UPR activation is demonstrated by our findings that UPR-deficient cells under sustained ER stress failed to accumulate ROS and were able to grow, while cells with a functional UPR accumulated ROS resulting in cell death. While our data implicate Ero1p and Pdi1p as the UPR target genes responsible for ROS accumulation, we cannot exclude the possibility that other UPR targets also contribute. Mitochondrial Involvement in ER Stress-Induced ROS Accumulation Mitochondria contributed significantly to lethal levels of ROS during periods of sustained ER stress as inactivation of respiration prevented the accumulation of ROS and cell death yet did not abolish ER stress. Depletion of GSH through disulfide bond reduction in the ER would allow ROS generated from mitochondria to accumulate unchecked and contribute to cell death. ROS originating from mitochondria may be derived through typical ROS production rates associated with respiration. Alternatively, ER stress or increased ROS may in turn signal or cause mitochondrial dysfunction, resulting in increased rates of ROS production. This would be consistent with previous observations in which GSH depletion resulted in subsequent mitochondrial dysfunction and consequential ROS accumulation (Merad-Boudia et al., 1998). Cell Death Is Independent of Proteasome Inhibition or Trafficking Defects Our model of ER stress-induced cell death is distinct from previous reports showing that overexpression of ERAD substrates, including mutant CFTR, lead to cellular defects or death due to cytosolic aggregation and

subsequent proteasome inhibition (Bence et al., 2001). In erv29⌬ cells expressing CPY*-25x, the capacity of the ubiquitin-proteasome system appears to be undiminished (see Supplemental Figure S8A at http://www. molecule.org/cgi/content/full/15/5/767/DC1/). Despite substantial ER stress, the secretory pathway also appeared to be functional, with the exception of ER-Golgi transport of Erv29p-dependent cargo, as the ER-Golgivacuole transport of alkaline phosphatase in erv29⌬ strains expressing CPY*-25x was very similar to transport in wild-type cells (Supplemental Figure S8B). These studies demonstrate that cell death caused by accumulation of misfolded CPY* in the ER is not due to a general impairment of the secretory pathway nor a deficiency in the ubiquitin-proteasome system. ER Stress, ROS Accumulation, and Human Disease It is intriguing that the UPR program employed by cells to alleviate ER stress, thus promoting cell survival, can also contribute to cell death if the program is activated for extended periods. Why is cell death or impairment not the typical outcome when the UPR is activated in cells expressing misfolded proteins in the ER? What distinguishes the many cell types in which the UPR is activated yet continue to grow from those cells that undergo PCD in the presence of ER stress? Contributing factors likely include the quantity of misfolded protein(s) being expressed, how many unstable disulfide bonds are formed within such protein(s), and the efficiency by which misfolded protein(s) are removed. This in turn affects the duration and level of UPR activation. From this perspective it is apparent that eukaryotic cells expressing immense quantities of correctly folded secretory proteins containing disulfide bonds such as insulinproducing pancreatic ␤ cells would also be at risk for ROS accumulation from prolonged ER stress as suggested previously (Harding et al., 2003). Such cells attempt to match molecular chaperone levels (including Pdi1p and Ero1p) to the high levels of nascent client proteins so as to provide sufficient folding capacity. These cells are presumably exposed to levels of ERderived oxidative stress proportional to the number of nascent disulfide bonds being formed. Such correctly folded proteins would not enter a thiol oxidation/reduction cycle, yet the sheer number of de novo disulfides formed may put such cells at risk. If these cells expressing large quantities of a single protein also possess a mutant allele for that secreted protein then the problem would be immensely exacerbated. The cell would now produce enormous quantities of an ER-retained misfolded protein that would almost certainly overwhelm the ERAD capacity and continually (re)-engage the oxidative folding machinery causing GSH depletion and ROS accumulation (similar to the effect of CPY*-25x). Several examples include insulin-secreting pancreatic ␤-cells (Oyadomari et al., 2002; Ron, 2002), arginine vasopressin (AVP)-secreting neurons (Ito and Jameson, 1997), and proteolipid protein (PLP)-secreting oligodendrocytes (Gow et al., 1998). In all these cases large quantities of proteins with disulfide bonds are secreted, and if misfolded, ER retention has been shown. Most intriguingly, ER retention in each particular cell type leads to cell death.

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A particularly well-studied example exists in the Akita diabetic mouse in which a mutant form of insulin (C96Y) causes diabetes mellitus likely because of pancreatic ␤-cell death (Oyadomari et al., 2002). The Ins2C96Y mutation prevents proper folding; however, mice have two insulin genes, which should be able to compensate if Ins2C96Y were simply a loss-of-function mutation instead of a dominant-negative effect (Leroux et al., 2001). Applying our model for ER stress-induced cell death, we propose that the dominant-negative phenotype of Ins2C96Y is due to accumulation in the ER causing UPR activation and an increase in the oxidative protein folding machinery. Constant/futile interactions between Ins2C96Y and PDI in the ER might deplete the cell of GSH and/or generate ROS leading to oxidative stress and contribute to cell death. While this may not account for all contributors to cell death in these scenarios, as genes that do not have obvious homologs in S. cerevisiae have been implicated, the generation of ROS due to an overactive UPR and oxidative folding machinery is likely. Persistent ER stress, GSH depletion, ROS accumulation, and cell death are also associated with a number of age-related neurological diseases including Alzheimer’s, Huntington’s, Parkinson’s, and the prion-based diseases (Merad-Boudia et al., 1998; Paschen and Frandsen, 2001). Our data suggest a mechanism by which persistent ER stress and prolonged UPR activation can account for some of these phenotypes. Experimental Procedures Chemicals were purchased from either Sigma or Fischer. Plasmids, Media, and Strain Construction Media were prepared as described previously (Hill and Stevens, 1994). SEY6211a, KHY30, KHY163, KHY271, KHY270, KHY281, KHY298, and KHY318 were described previously (Caldwell et al., 2001; Haynes et al., 2002). To introduce the ire1⌬::KAN, hac1⌬::KAN, and yca1⌬::KAN alleles into strains, oligonucleotides were used to amplify the disrupted allele DNA from strains of the S. cerevisiae Genome Deletion Project (Research Genetics, Invitrogen) that was then introduced into cells with selection on YEPD media containing 200 ␮g/ml G418 (Calbiochem). Disruptions were confirmed by PCR analysis on genomic DNA using oligonucleotides flanking each disrupted locus. In this manner, ire1⌬::KAN or hac1⌬::KAN alleles were introduced into SEY6211a to create KHY235 and KHY233, respectively. KHY438 and KHY439 were created by integrating prc1-1 (the gene that expresses CPY* [Finger et al., 1993]) into KHY422 and KHY430, which were created by introducing erv29⌬::HIS3 into KHY235 and KHY233. ero1-1 was introduced into KHY271 via an ERO1 knockout followed by a plasmid shuffle to create strain KHY490. ero1⌬::HIS3 was introduced into KHY270 harboring pAC693. The chromosomal disruption strain was selected on media lacking histidine and confirmed by PCR. pAC693 was replaced by pAC695 via plasmid shuffling. ␳⬚ cells were generated by growing cells overnight in media containing 5 mg/ml ethidium bromide, and respiration deficiency was confirmed by lack of growth on media containing 3% glycerol and 2% ethanol. To create pAC595, a CEN plasmid in which the prc1-1 allele is preceded by the CUP1 promoter, the prc1-1 allele was amplified by PCR from KHY163-genomic DNA such that a ClaI site was introduced immediately preceding the start codon of prc1-1, and the product inserted into pTOPO (Invitrogen) to create pAC587. The XhoI-SacI fragment containing prc1-1 was excised from pAC587 and inserted into pRS316 at the corresponding restriction sites to create pAC594. The CUP1 promoter (an 865 bp fragment immediately preceding the CUP1 gene) was amplified by PCR from KHY163 introducing flanking HindIII (5⬘) and ClaI (3⬘) restriction sites and inserted into pTOPO to create pAC586. pAC595 was created by

excising the SacI-ClaI CUP1 promoter containing fragment from pAC586 and inserting it into pAC587 cut with SacI and ClaI. pAC667 is identical to pAC595 with prc1-1 replaced with PRC1. PRC1 was amplified from SEY6211a-genomic DNA and ligated into pTOPO to create pAC666. To create pAC667, the CUP1 promoter from pAC586 (as a SacI-ClaI fragment) was ligated with PRC1 (a ClaI-XhoI fragment from pAC666) and SacI-XhoI-digested pRS316. To construct the cysteine-less allele of CPY* (Cys-less CPY*-25x), all eleven cysteine residues of CPY* were mutated to alanine within pAC595 by a combination of site-specific mutagenesis (Quik Change, Stratagene, La Jolla, CA) and sequence replacement with a synthesized mutagenized gene fragment (Retrogen Inc, San Diego, CA). pAC693 is a CEN (pRS316 digested with SacI-XhoI) plasmid containing ERO1, which was PCR amplified, with oligonucleotides which introduced flanking 5⬘ XhoI and 3⬘ SacI restriction sites from KHY163 and ligated into pTOPO to create pAC669. pAC695 is a CEN plasmid containing ero1-1. ero1-1 was PCR amplified from CKY559 (a gift from Dr. Chris Kaiser) and ligated into pTOPO to create pAC670. The SacI-XhoI ero1-1 fragment from pAC670 was ligated into SacI-XhoI-digested pRS314. The ero1⌬::HIS3 disruption cassette was created by ligating the BamHI-BamHI HIS3 fragment from pJJ217 into BglII-digested pAC672. pAC672 was created by excising the XhoI-SacI ERO1 fragment from pAC669 and ligated into the SalI-SacI sites of pTV3, a 2 ␮ plasmid. Growth Rates and UPR Activation Assays Growth curves were generated by growing the described strains overnight in synthetic media at 30⬚C to log phase and then diluting them to 0.1–0.3 OD600nm. CuSO4 was added to 150 ␮M, and subsequent OD600nm readings were taken at the indicated time points. UPR activation was measured as described (Friedlander et al., 2000). Glutathione Assays, Mixed Disulfide Trapping, Protein Preparation, Antibodies, Western Blotting, and Immunofluorescence Total glutathione and oxidized glutathione levels were measured as described (Cuozzo and Kaiser, 1999). Radiolabel pulse chase experiments were performed as described (Haynes et al., 2002). Mixed disulfide trapping to determine the quantity of CPY* or wtCPY bound to Pdi1p was performed as described (Frand and Kaiser, 1999). In short, erv29⌬ cells expressing either CPY*-25x or wtCPY-25x were radiolabeled for 10 min. This was followed by the addition of cycloheximide (50 ␮g/ml) to prevent further CPY synthesis and chased with nonradioactive methionine and cysteine. Six OD600nm were harvested at the described times, suspended in 10% TCA, washed, cell walls removed, and lysed in a buffer containing SDS under nonreducing conditions. The lysate was split into 4 aliquots. One-tenth was immunoprecipitated with anti-Pdi1p antibodies, 1/10 was immunoprecipitated with anti-Vma1p antibodies, and 1/10 was immunoprecipitated with anti-CPY antibodies, and all were loaded onto a reducing 10% SDS-PAGE gel. These three samples represented total amounts of the respected proteins. The remaining 7/10 was immunoprecipitated with anti-Pdi1p antibodies under nonreducing conditions. The samples were then reduced with 100 mM DTT and sequentially immunoprecipitated with anti-CPY antibodies. Quantitation was performed using a phosphoimager and is expressed as the amount of wtCPY/CPY* bound to Pdi1p via mixed disulfide divided by the total amount of wtCPY/CPY*. This number was multiplied by 100 to give the percentage of wtCPY or CPY* in the cell that is interacting with Pdi1p via mixed disulfide bonds at the indicated time. Western blots were performed as described (Haynes et al., 2002). Monoclonal HA antibodies were purchased from Molecular Probes. Polyclonal Eug1p and Pdi1p antibodies were a gift from Dr. Tom Stevens. The secondary antibodies used for Western blots were purchased from Bio-Rad and those used for immunofluorescence from Molecular Probes. TUNEL, Annexin V, and ROS Detection TUNEL assays were performed by growing cells overnight in media lacking CuSO4 to log phase; 150 ␮M CuSO4 was added, and at the described times 5 OD600nm cells were harvested and exposed to 4% formaldehyde (Sigma) for 1 hr. The cell walls were removed using

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oxalyticase (Enzogenetics) and zymolyase (Seikagaku Corporation), washed in 1.2 M sorbitol, and subsequently adhered to polylysine coated microscope slides. The TUNEL assay was performed using the ApoAlert DNA Fragmentation Assay kit (Clontech/BD Biosciences). The annexin V assay and ROS detection were performed as described (Madeo et al., 1997a). Supplemental Data Supplemental Data, including additional data and detailed Experimental Procedures used in this work, are available at http://www. molecule.org/cgi/content/full/15/5/767/DC1/. Acknowledgments We would like to thank Drs. David Ron and Chris Kaiser for helpful discussions, Kathryn Hill for critical reading of the manuscript, and Drs. Kazutoshi Mori and Tom Stevens for reagents. C.M.H. was supported by the UMKC Distinguished Dissertation fellowship. This work was supported by NIH grant GM55848 (to A.A.C). Received: December 24, 2003 Revised: June 16, 2004 Accepted: June 21, 2004 Published: September 9, 2004 References Bence, N.F., Sampat, R.M., and Kopito, R.R. (2001). Impairment of the ubiquitin-proteasome system by protein aggregation. Science 292, 1552–1555. Caldwell, S.R., Hill, K.J., and Cooper, A.A. (2001). Degradation of endoplasmic reticulum (ER) quality control substrates requires transport between the ER and Golgi. J. Biol. Chem. 276, 23296– 23303. Chapman, R., Sidrauski, C., and Walter, P. (1998). Intracellular signaling from the endoplasmic reticulum to the nucleus. Annu. Rev. Cell Dev. Biol. 14, 459–485.

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