The proteasome homeostasis in Saccharomyces cerevisiae is regulated by a negative feedback loop in which the transcription factor Rpn4 induces the proteasome genes and is rapidly degraded by the assembled proteasome. In addition to the proteasome genes, Rpn4 regulates numerous other genes involved in a wide range of cellular pathways. Therefore, the Rpn4–proteasome negative feedback circuit not only controls proteasome abundance, but also gauges the expression of other Rpn4 target genes. Our previous work has shown that Rpn4-induced gene expression is critical for cell viability under stressed conditions. Here we investigate whether proteasomal degradation of Rpn4 is also important for cell survival in response to stress. To this end, we generate a stabilized Rpn4 mutant (Rpn4*) that retains its transcription activity. We find that expression of Rpn4* severely reduces cell viability in response to various genotoxic and proteotoxic agents. This detrimental effect can be eliminated by a point mutation that abolishes the transcription activity of Rpn4*, suggesting that overexpression of some Rpn4 target genes weakens the cell's ability to cope with stress. Moreover, we demonstrate that inhibition of Rpn4 degradation causes synthetic growth defects when combined with proteasome impairment resulting from mutation of a proteasome gene or accumulation of misfolded endoplasmic reticulum membrane proteins. Rpn4 thus represents an important stress-responsive mediator whose degradation as well as availability are critical for cell survival under stressed conditions.
THE Saccharomyces cerevisiae RPN4 gene (also named SON1 and UFD5) was originally isolated as a suppressor of sec63-101, a temperature-sensitive mutant of SEC63, which encodes an essential component of the endoplasmic reticulum (ER) translocation channel (Nelson et al. 1993; Johnson et al. 1995; Finley et al. 1998; Fujimoro et al. 1998). Subsequent work showed that deletion of RPN4 inhibits the degradation of several model substrates of the N-end rule and UFD (Ub fusion degradation) pathways, suggesting the involvement of Rpn4 in proteasomal degradation (Johnson et al. 1995). The underlying mechanism, however, remained unsolved until recent studies revealed that Rpn4 is a transcription factor for the proteasome genes (Mannhaupt et al. 1999; Xie and Varshavsky 2001). Interestingly, Rpn4 is an extremely short-lived protein (t1/2 ≤ 2 min) and is degraded by the proteasome (Xie and Varshavsky 2001; Ju et al. 2004, 2007, 2008; Ju and Xie 2004, 2006; Wang et al. 2004). Stabilization of Rpn4 by inhibiting the proteasome results in an increase in the expression of the proteasome genes (Ju et al. 2004; London et al. 2004). Together, these observations led to a model in which the proteasome homeostasis is regulated by a negative feedback circuit. On one hand, Rpn4 upregulates the proteasome genes; on the other hand, Rpn4 is rapidly degraded by the assembled/active proteasome. The Rpn4–proteasome negative feedback circuit provides an efficient and sensitive means to control the proteasome abundance in S. cerevisiae. Whereas the homologs of Rpn4 have not yet been identified in higher eukaryotes, their proteasome genes are clearly regulated by a negative feedback mechanism (Lundgren et al. 2003; Meiners et al. 2003; Wójcik and DeMartino 2002; Xu et al. 2008).
In addition to the proteasome genes, Rpn4 appears to regulate numerous other genes involved in a wide range of cellular processes (Mannhaupt et al. 1999; Jelinsky et al. 2000; Gasch et al. 2001; Haugen et al. 2004; Guo et al. 2008; Hausmann et al. 2008; Salin et al. 2008; Teixeira et al. 2008). Interestingly, the promoter of RPN4 carries the binding sites for heat-shock transcription factor 1; multidrug resistance-related transcription factors Pdr1 and Pdr3; and Yap1, a transcription factor that plays an important role in response to oxidation and DNA damage (Owsianik et al. 2002; Haugen et al. 2004; Hahn et al. 2006). These transcription factors are activated by a variety of stressors and in turn induce RPN4 expression. Thus, Rpn4 is considered to be an important stress-responsive mediator. In fact, it has been shown that rpn4Δ cells are sensitive to different stressing agents (Jelinsky et al. 2000; Ng et al. 2000; Ju et al. 2004; London et al. 2004; Hanna et al. 2007; Wang et al. 2008).
Among the pathways involving Rpn4, the Rpn4–proteasome negative feedback loop likely plays a central role. It not only controls proteasome homeostasis, but also regulates the expression of other Rpn4 target genes through proteasomal degradation of Rpn4. Our previous work has demonstrated that disruption of Rpn4-induced proteasome expression severely reduces cell viability under stressed conditions (Wang et al. 2008). In this study, we sought to determine whether Rpn4 degradation is also important for cell survival in response to stress. We found that inhibition of Rpn4 degradation dramatically sensitizes cells to various genotoxic and proteotoxic stressors. This detrimental effect is eliminated by a point mutation that inactivates the transcription activity of Rpn4, suggesting that overexpression of Rpn4 target genes impairs the cell's ability to tolerate stress. We also demonstrated that stabilization of Rpn4 exhibits synthetic growth defects with proteasome impairment. The underlying mechanism is further discussed.
MATERIALS AND METHODS
Plasmids and yeast strains:
The details of plasmid construction are available upon request. Briefly, full-length RPN4 and a truncated mutant encoding Rpn4Δ1-10/Δ211-229 (RPN4*) were ligated with the RPN4 promoter (∼500 bp) into the low-copy vectors pRS314 and pRS315 (Sikorski and Hieter 1989), resulting in p314RPN4, p315RPN4, p314RPN4*, and p315RPN4*. For immunoblotting and immunoprecipitation assays, a DNA sequence encoding a triple ha tag (3ha) was inserted immediately upstream of the stop codons of RPN4 and RPN4* in p314RPN4 and p314RPN4* to generate p314RPN4-3ha and p314RPN4*-3ha. To construct replacement vectors, p314RPN4-3ha and p314RPN4*-3ha were cut with XhoI and SacI, and the inserts were subcloned into a XhoI/SacI-cut pRS304 vector to obtain p304RPN4-3ha and p304RPN4*-3ha. p315CUPRPN4*-C477A expresses a stabilized and transcriptionally inactive Rpn4 allele (Rpn4*-C477A) from the CUP1 promoter in pRS315 (Wang et al. 2004). The plasmids expressing Rpn4172-229-βgal and Rpn4172-229/K187R-βgal from the CUP1 promoter were previously described (Ju and Xie 2006). Plasmids expressing CPY* (pSM2215), ste6-G38D (pSM1898), and N-terminally His6-tagged VHL-L158P (pSM2016), and a control vector (pSM922) were kindly provided by Susan Michaelis (Metzger and Michaelis 2009).
To generate a strain that expresses RPN4* from the native chromosomal locus, p304RPN4*-3ha was linearized by XhoI and transformed into strain EJY140 (MATa his3-Δ200 leu2-3, 112 lys2-801 trp1-Δ63 ura3-52 rpn4Δ∷LEU2; see Johnson et al. 1995). The replacement of the rpn4Δ∷LEU2 allele by the RPN4*∷TRP1 cassette through recombination at the RPN4 promoter region and the vector sequences of pRS304 and pRS305 was illustrated by a TRP1+leu2− phenotype. An RPN4∷TRP1 wild-type strain was also constructed by using a similar approach with a XhoI-linearized p304RPN4-3ha vector. Immunoblotting analysis with an anti-ha antibody confirmed the expression of Rpn4-3ha and Rpn4*-3ha in the wild-type and RPN4* strains. Other yeast strains used in this study included JD52 (MATa his3-Δ200 leu2-3, 112 lys2-801 trp1-Δ63 ura3-52), YXY210 (MATa trp1-Δ63 ura3-52 his3-Δ200 leu2-3,112 lys2-801 PRE1-FLAG-His6∷YIplac211 rpn4Δ∷LEU2), WCG4a (MATa ura3 his3-11,15 leu2-3,112), and YHI29/1 (MATα pre1-1 ura3 his3-11,15 leu2-3,112). These yeast strains were described previously (Heinemeyer et al. 1993; Johnson et al. 1995; Ju et al. 2004).
Analysis of gene expression:
Quantitative real-time PCR (qRT–PCR) was applied to measure gene expression. RNA was prepared from cells grown to an OD600 of 0.8–1.2 as previously described (Schmitt et al. 1990). Reverse-transcription was carried out with SuperScript II reverse transcriptase (Invitrogen), and qRT–PCR was performed using the Fast SYBR Green Master Mix in the StepOne system following the manufacturer's instruction (Applied Biosystems). The primers used in qRT–PCR included YX717 (ACTGGATCCCCTAATTGACTTGGCTAATTC) and YX732 (CAAGAATTCATGGATATTATTCTGGGCATC) for PRE1; YX125 (CAGGAACTTTTTCCACCATCA) and YX126 (CTGAGGGAAAATACATCGTGG) for RPT6; and YX814 (TCCATCCAAGCCGTTTTG) and YX815 (CGGCCAAATCGATTCTCA) for ACT1, which served as an internal control to normalize the levels of gene expression.
β-Galactosidase enzymatic assay and immunoblotting analysis:
The enzymatic activity of β-galactosidase in liquid yeast cultures was determined using the chromogenic substrate o-nitrophenyl-β-d-galactopyranoside (ONPG) (Ju and Xie 2006). For induction of the CUP1 promoter driving the expression of Rpn4172-229-βgal and Rpn4172-229/K187R-βgal, yeast cells grown to an OD600 of 0.3–0.5 were treated with CuSO4 at a final concentration of 0.1 mm for 6–7 hr. For immunoblotting analysis, yeast cells were grown to an OD600 of 0.8–1.2, harvested, and resuspended in lysis buffer (150 mm NaCl, 50 mm Tris.Cl, 5 mm EDTA, 1% Triton X-100, pH 7.5) plus protease inhibitor mix (Roche). Yeast extracts were prepared using the glass-bead vortexing method. Protein concentrations were measured by Bradford assay. Approximately 50 μg of each extract was separated by SDS–PAGE, followed by immunoblotting with an anti-ha antibody (Covance) for C-terminally 3ha-tagged Rpn4 and Rpn4* and an anti-FLAG antibody (Sigma) for C-terminally FLAG-tagged Pre1 and detection with the Odyssey infrared imaging system (Li-Cor Biosciences). The blots were reprobed with an anti-yeast α-tubulin antibody (Chemicon) to verify comparable loading.
S. cerevisiae cells from 10-ml cultures (OD600 of 0.8–1.0) in SD medium with essential amino acids were harvested. The cells were resuspended in 0.3 ml of the same medium supplemented with 0.15 mCi of [35S]methionine/cysteine (EXPRESS [35S] Protein Labeling Mix, PerkinElmer) and incubated at 30° for 5 min. The cells were then pelleted and resuspended in the same medium with cycloheximide (0.2 mg/ml) and excessive cold l-methionine/l-cysteine (2 mg/ml l-methionine and 0.4 mg/ml l-cysteine) and chased at 30°. An equal volume of sample was withdrawn at each time point. Labeled cells were harvested and lysed in equal volume of 2× SDS buffer (2% SDS, 30 mm dithiothreitol, 90 mm Na–HEPES, pH 7.5) by incubation at 100° for 3 min. The supernatants were diluted 20-fold with buffer A (1% Triton X-100, 150 mm NaCl, 1 mm EDTA, 50 mm Na–HEPES, pH 7.5) before being applied to immunoprecipitation with anti-ha antibody (Sigma) combined with protein A agarose (Calbiochem). The volumes of supernatants used in immunoprecipitation were adjusted to equalize the amounts of 10% trichloroacetic acid-insoluble [35S]. The immunoprecipitates were washed three times with buffer A and resolved by SDS–PAGE, followed by autoradiography and quantitation with a PhosphorImager (Molecular Dynamics).
Cell growth and survival assays:
Cell growth was assessed by serial dilution assays. Yeast cells were grown to exponential phase (OD600 of 0.8–1.2) and normalized by optical density. Fivefold serial dilutions of cell cultures were spotted on selective plates and incubated at 30°. Alternatively, cell growth was directly measured in liquid cultures. Specifically, overnight cultures were diluted to an OD600 of ∼0.2 and continued to grow to stationary phase. Cell density (OD600) was monitored at different time points and applied to plotting cell-growth curves. Cell survival was measured by colony formation assay. Yeast cultures were grown to exponential phase before being subjected to stresses. Cells were treated with various doses of MMS and t-BuOOH for 1 hr, extensively washed with medium, and plated on selective synthetic complete medium. For UV stress, cells were plated and incubated for 4 hr before being exposed to UV radiation. The plates were kept in darkness afterward. To examine canavanine sensitivity, cells were directly seeded on minimal medium (SD) plates containing different concentrations of canavanine. Colonies were counted 72 hr after plating. The survival rates of treated cells were normalized against that of untreated cells, which was set at 100%.
Generation of a stabilized and transcriptionally active Rpn4 mutant:
An ideal experimental setting to examine the phenotypes caused by inhibition of Rpn4 degradation is to express a stabilized and transcriptionally active version of Rpn4. This has been a difficult task because Rpn4 is extremely short-lived in vivo and can be degraded by two distinct mechanisms, ubiquitin (Ub)-dependent and -independent (Ju and Xie 2004). Our recent studies showed that the N-terminal 10 amino acids of Rpn4 are required for the Ub-independent degradation mode, whereas residues 211–229 constitute the Ub-dependent degradation signal (Ju and Xie 2004, 2006; Wang et al. 2004; Ju et al. 2007, 2008). Using pulse-chase analysis, we demonstrated that Rpn4Δ1-10/Δ211-229, an Rpn4 mutant simultaneously deleted of residues 1–10 and 211–229 (hereafter referred to as Rpn4*; Figure 1A), was substantially stabilized (Figure 1, B and C). We then decided to examine whether Rpn4* retains transcriptional activity. Low-copy vectors expressing RPN4 or RPN4* from the native RPN4 promoter or a control vector were transformed into an rpn4Δ strain, and the expression levels of two proteasome genes, PRE1 and RPT6, were measured by qRT–PCR. As expected, the proteasome genes were induced by Rpn4 (Figure 1D). We found that the RPN4* transformants had higher mRNA levels of PRE1 and RPT6 than their RPN4 counterparts (Figure 1D). In line with the qRT–PCR results, the Pre1 protein level was also relatively higher in the RPN4* transformants than in the RPN4 counterparts (Figure 1E; compare lanes 2 and 3). These results indicated that Rpn4* is competent in activating its target genes. The elevated expression of PRE1 and RPT6 in the RPN4* transformants likely reflected a higher steady-state level of Rpn4* due to its stabilization.
To further examine the effect of Rpn4* on the expression of other proteasome genes, we compared the proteasome activity of the three rpn4Δ transformants. A plasmid expressing Rpn4172-229-βgal, a short-lived βgal fusion protein with the Ub-dependent degradation signal of Rpn4 (Ju and Xie 2006), was introduced into the rpn4Δ transformants. The degradation of Rpn4172-229-βgal was assessed by ONPG assay, which measures the remaining βgal enzymatic activity (Ju and Xie 2006). Rpn4172-229/R187K-βgal, a stabilized derivative of Rpn4172-229-βgal with a K187R substitution (Ju and Xie 2006), was used as a control to ensure that the disparity of βgal activity is not due to variation of transcription and/or translation of the βgal substrates. Whereas its degradation was inefficient in the absence of Rpn4, Rpn4172-229-βgal was rapidly degraded in the cells expressing RPN4 or RPN4* (Figure 1F). These results again demonstrated that Rpn4* is capable of inducing its target genes. In fact, the proteasome activity was slightly higher in the RPN4* transformants than in their RPN4 counterparts, consistent with the increase in proteasome expression in the RPN4* transformants (Figure 1, D and E).
Proteasomal degradation of Rpn4 is crucial for cell survival under stressed conditions:
To study the biological significance of Rpn4 degradation, we constructed a yeast strain that expresses RPN4* from the native chromosomal locus. Specifically, we replaced the rpn4Δ∷LEU2 allele in strain EJY140 with an RPN4*∷TRP1 cassette via site-specific recombination (Figure 2A). Using a similar approach, we generated a wild-type control strain by replacing the rpn4Δ∷LEU2 allele with an RPN4∷TRP1 cassette. The RPN4* strain grew relatively slower than its wild-type counterpart under normal conditions (Figure 2B). The growth rate difference between these two strains was most notable at an early stage. RPN4* acted as a dominant mutant because expression of wild-type RPN4 from an episomal vector could not suppress the slow-growth phenotype of the RPN4* strain (data not shown).
We then assessed the effect of the inhibition of Rpn4 degradation on cell viability under stressed conditions. Wild-type and RPN4* strains were treated with various doses of DNA-damaging agents, including UV radiation and the alkylating agent MMS. The survival rates were measured by colony formation assays (Figure 3, A and B). We found that RPN4* cells were much more sensitive to UV and MMS than wild-type cells, especially at higher doses. The survival rate of RPN4* cells after treatment with 180 J/m2 UV or 0.2% MMS was ∼0.1%, whereas >1% of wild-type cells survived under the same conditions. We also compared the sensitivity of RPN4 and RPN4* cells to the oxidizing agent t-BuOOH. Only ∼0.03% of RPN4* cells grew to form colonies after treatment with 30 mm t-BuOOH, whereas >0.3% of wild-type cells survived with the same treatment (Figure 3C).
We further examined whether proteasomal degradation of Rpn4 is critical for cell survival in response to proteotoxic stress. Specifically, we challenged RPN4 and RPN4* cells with canavanine, an arginine analog that can be efficiently incorporated into nascent proteins in the place of arginine, thereby producing structurally aberrant proteins that may not function properly. As shown in Figure 3D, the survival rate of RPN4* cells was significantly lower than that of wild-type cells in the presence of canavanine. Thus, proteasomal Rpn4 degradation is crucial for cell survival in response to genotoxic and proteotoxic stressors.
The detrimental effect of Rpn4* depends on its transcriptional activity:
To gain insight into the mechanism underlying the stress hypersensitivity caused by the inhibition of Rpn4 degradation, we wanted to determine whether the detrimental effect of Rpn4* is related to its transcriptional activity. To this end, we introduced a Cys-to-Ala substitution at codon 477 of the C2H2 DNA-binding motif of Rpn4*, thereby generating a stable but transcriptionally inactive Rpn4 mutant (Rpn4*-C477A). (Our previous study has shown that the C477A mutation abolishes the transcription activity of Rpn4; see Wang et al. 2004.) Low-copy plasmids expressing RPN4, RPN4*, or RPN4*-C477A were transformed into a wild-type (RPN4) strain. The transformants were challenged with UV radiation, and the survival rates were measured by a colony formation assay. While Rpn4* sensitized cells to UV exposure, Rpn4*-C477A displayed no effect (Figure 4A). Similarly, Rpn4*-C477A did not reduce cell viability in response to MMS, t-BuOOH, and canavanine (data not shown). These results indicate that the negative effect of Rpn4* on cell survival under stressed conditions is related to its transcription activity.
We suspected that Rpn4* over-induces its target genes and that some of the protein products are toxic if not rapidly degraded by the proteasome. To test this hypothesis, we transformed low-copy plasmids expressing RPN4, RPN4*, or RPN4*-C477A or a control vector into a proteasome mutant, pre1-1, and its wild-type counterpart. The pre1-1 mutant bears a mutation in the proteasome subunit Pre1, and its proteasome activity is compromised (Heinemeyer et al. 1993). Whereas Rpn4* had only a mild effect on wild-type cells, it severely impaired the growth of pre1-1 cells (Figure 4B). Notably, Rpn4*-C477A did not affect the growth of pre1-1. These results reveal that overproduction of Rpn4 target proteins and proteasome impairment exhibit a synthetic effect on cell growth. This synthetic effect likely contributes to the vulnerability of RPN4* cells to stress. Under stressed conditions, the proteasome may be choked by damaged proteins and fail to remove the toxic Rpn4 target proteins in a timely manner, thereby reducing cell viability.
Inhibition of Rpn4 degradation sensitizes cells to overexpression of misfolded ER membrane proteins:
The hypersensitivity of RPN4* cells to canavanine suggests that inhibition of Rpn4 degradation impairs the cell's ability to tolerate misfolded proteins. We went on to examine whether RPN4* cells are sensitive to misfolded proteins located in a specific cellular compartment. Specifically, we compared the sensitivity of RPN4* cells to overexpression of three different classes of misfolded proteins. CPY*, a misfolded form of carboxypeptidase Y, is soluble and retained in the ER lumen (Hiller et al. 1996). ste6-G38D, which bears a mutation in the first transmembrane span of the ABC transporter Ste6, represents a misfolded ER membrane protein (Metzger and Michaelis 2009). VHL-L158P, a mutated allele of the mammalian von Hippel Lindau tumor suppressor protein (VHL) serves as a cytosolic misfolded protein (McClellan et al. 2005). Plasmids expressing these mutated alleles from the galactose-inducible GAL1 promoter or an empty vector were introduced into RPN4, RPN4*, and rpn4Δ cells, respectively. The transformants were seeded on glucose and galactose plates, and the relative survival rates (galactose vs. glucose) were measured by colony formation assay (Figure 5A). In the absence of the overexpression of misfolded proteins, RPN4, RPN4*, and rpn4Δ cells displayed a similar colony formation efficiency. Overexpression of CPY* and ste6-G38D modestly reduced the viability of RPN4 cells. Interestingly, RPN4* cells were much more sensitive to overexpression of ste6-G38D than their RPN4 counterparts. By contrast, they tolerated CPY* accumulation slightly better than RPN4 cells. We found that rpn4Δ cells were also very sensitive to overexpression of ste6-G38D. However, unlike RPN4* cells, rpn4Δ cells were relatively more sensitive to CPY* than RPN4 cells. Differing from ste6-G38D and CPY*, overexpression of VHL-L158P had no appreciable effect on the survival of RPN4, RPN4*, and rpn4Δ cells. Immunoblotting analysis confirmed the expression of VHL-L158P upon induction by galactose (supporting information, Figure S1). These results suggest that inhibition of Rpn4 degradation sensitizes cells specifically to the accumulation of misfolded ER membrane proteins.
Since Rpn4* slightly but reproducibly mitigated the toxicity of CPY* overexpression (Figure 5A), we wanted to examine whether Rpn4* could lessen the detrimental effect of the accumulation of other misfolded ER lumenal proteins. To this end, we compared the sensitivity of RPN4, RPN4*, and rpn4Δ cells to tunicamycin, a well-known inhibitor that blocks protein folding in the ER lumen. As shown in Figure 5B, RPN4* cells were indeed relatively more resistant to tunicamycin than RPN4 cells. Consistent with the data from CPY* analysis, rpn4Δ cells were more sensitive to tunicamycin than RPN4 cells. Thus, whereas inhibition of Rpn4 degradation severely reduces cell viability in the presence of the accumulation of misfolded ER membrane proteins, it modestly enhances the cell's ability to cope with misfolded lumenal proteins.
Recent studies have revealed a stress-response network in S. cerevisiae where Rpn4 may serve as a major mediator (Jelinsky et al. 2000; Ng et al. 2000; Owsianik et al. 2002; Haugen et al. 2004; Ju et al. 2004; London et al. 2004; Hahn et al. 2006; Hanna et al. 2007; Salin et al. 2008; Teixeira et al. 2008; Wang et al. 2008; Metzger and Michaelis 2009). RPN4 is activated by a variety of stressors; and Rpn4 in turn upregulates a large number of stress-responsive genes, including the proteasome genes and others involved in protein ubiquitylation, DNA repair, and other cellular processes.
The induction of RPN4 under stressed conditions suggests that an increase in the Rpn4 protein level is important for cell survival in response to stress. One would expect that stabilization of Rpn4 might increase cell viability under stressed conditions. Surprisingly, we found that inhibition of Rpn4 degradation actually sensitizes cells to various genotoxic and proteotoxic agents. It is likely that, whereas a transient increase in Rpn4 expression benefits the cell by inducing relevant stress-responsive genes, some of the Rpn4 target proteins may be toxic to the cell if overproduced and accumulated. In support of this argument, we showed that the toxicity of Rpn4* can be eliminated by a point mutation that inactivates its transcription activity. We further demonstrated that expression of RPN4* severely impairs the growth of proteasome mutant cells but has no significant effect on wild-type cells. In addition, we found that cells expressing Rpn4* are hypersensitive to accumulation of ste6-G38D, a misfolded ER membrane protein. Interestingly, a recent study reported that the proteasome activity is impaired in cells overexpressing ste6-G38D (Metzger and Michaelis 2009). These results indicate that over-induction of Rpn4 target genes is toxic if the protein products cannot be removed rapidly by the proteasome. We infer from these results that the toxic Rpn4 target proteins are normally short-lived but may be stabilized under stressed conditions because the proteasome could be occupied by damaged proteins.
It is currently unclear why RPN4* cells tolerate the accumulation of misfolded lumenal proteins, such as CPY*, relatively better than wild-type cells (Figure 5). Unlike ste6-G38D, overexpression of CPY* does not impair the proteasome activity (Metzger and Michaelis 2009). Therefore, the Rpn4 target proteins are presumably not accumulated in the cells overexpressing CPY*. On the other hand, the relatively higher proteasome expression level induced by Rpn4* may facilitate the degradation of misfolded lumenal proteins via the ER-associated degradation pathway.
It is worthy of note that rpn4Δ cells are also hypersensitive to the stressors that impair the growth and viability of RPN4* cells, such as UV, MMS, t-BuOOH, and ste6-G38D (Figures 3 and 5; also see Wang et al. 2008). Thus, both availability and degradation of Rpn4 are critical for cell survival under stressed conditions. This underscores the biological significance of the Rpn4–proteasome negative feedback circuit. On one hand, Rpn4 induces proteasome and other stress-responsive genes to cope with stress; on the other hand, proteasomal degradation prevents Rpn4 target genes from being over-induced by keeping the Rpn4 protein level in check. The importance of Rpn4 degradation in cellular stress is also reflected by the fact that Rpn4 can be degraded by two distinct pathways that are Ub-dependent and -independent (Ju and Xie 2004). This dual degradation mode guarantees the efficient removal of Rpn4 even if one of the degradation pathways is impaired by stressors. This study provides an interesting model molecule for understanding the molecular basis of the cellular stress response.
We thank Susan Michaelis for plasmids and Li Li for access to the qRT–PCR instrument. This study was supported by a National Science Foundation grant (MCB-0816974) to Y.X.
Supporting information is available online at http://www.genetics.org/cgi/content/full/genetics.109.112227/DC1.
↵1 These authors contributed equally to this work.
↵2 Present address: Department of Pediatrics, University of Texas Southwestern Medical Center, Dallas, TX 75390.
Communicating editor: A. P. Mitchell
- Received September 24, 2009.
- Accepted November 22, 2009.
- Copyright © 2010 by the Genetics Society of America