Proteins are susceptible to heat-induced degradation for 30–60 min after synthesis

To determine how long after synthesis cell proteins are sensitive to thermally induced degradation, we compared the effects on proteolysis of shifting yeast from 30 to 38°C immediately after the 5-min pulse of ³⁵S-Met or 30, 60, or 90 min later (Fig. 1 B). The cells shifted at 30 min showed a smaller increase in proteolysis than those shifted immediately after synthesis, and 60 min after synthesis the effect of temperature upshift was much smaller and not statistically significant. By 90 min, the shift to 38°C failed to increase degradation. Thus, during the hour after synthesis proteins appeared to become insensitive to thermal damage, as reflected in their resistance to heat-induced degradation. Because it is unlikely that the folding of many proteins would require 30–60 min, this “fragile period” presumably represents the time necessary for postsynthetic modifications, proper multimer formation, and localization, which together contribute to thermal resistance. Presumably, as proteins achieve stable multimeric structures and their final folded conformations, they become more resistant to thermal denaturation.

ROS stimulate the degradation of recently synthesized proteins

A variety of experiments using free radical generators or hydrogen peroxide (ROS) have indicated that oxidant damage to cell proteins can enhance their degradation (Fagan et al., 1986; Davies, 1987; Davies and Goldberg, 1987). To study the effects of oxygen radicals on the breakdown of recently synthesized and long-lived yeast proteins, we used paraquat, a superoxide generator (Lee et al., 2004), as well as cadmium, which can replace the Fe²⁺ in proteins and seems to generate oxygen radicals as a consequence of the released Fe²⁺ (Thevenod and Friedmann, 1999; Vido et al., 2001). Exposure of yeast to Cd++ causes induction of enzymes that defend against ROS, molecular chaperones, ubiquitination enzymes, and Cdc48p (Vido et al., 2001). During exponential growth, wild-type (WT) yeast were exposed to 80 μg/ml paraquat or 50 μg/ml cadmium. After 90 min, these cells and untreated controls were labeled with ³⁵S-Met for 5 min, and then we compared the rates of protein degradation. At these low concentrations, neither cadmium nor paraquat reduced cell growth or protein synthesis during these experiments, but the proteins synthesized in their presence were initially degraded two- to threefold faster than controls (Fig. 2 A). As found after a shift to 38°C, the oxidant-induced acceleration of degradation was only clearly demonstrated during the first 60 min after synthesis. After this time, the proteins synthesized in the presence of cadmium or paraquat were degraded at similar rates to those in control cells (Fig. 2 A). These findings suggest that the degradation of the long-lived cell proteins is not affected by oxidative damage.

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Figure 2.

Exposure to paraquat or cadmium stimulates the degradation of recently synthesized but not long-lived proteins, and 60 min after synthesis, proteins no longer show this response to paraquat and cadmium. (A) During exponential growth at 30°C, yeast were exposed to 80 μg/ml paraquat or 50 μg/ml cadmium for 90 min, and then ³⁵S-Met was added for 5 min at 30°C to label recently synthesized proteins or for 90 min to label long-lived proteins. After washing and resuspension in chase medium, rates of protein degradation were measured as in Fig. 1 A. (B) Proteins were labeled with ³⁵S-Met for 5 min, washed, and then paraquat and cadmium were added immediately or 60 min after synthesis. The amounts of degradation after 1 h were measured. Degradation rates after 2 and 3 h showed similar differences as after 1 h (not depicted).

To verify this conclusion, growing yeast were exposed to paraquat or cadmium for 90 min. These cells and untreated controls were incubated with ³⁵S-Met for 90 min to label primarily long-lived cell components, and after washing, the rates of protein degradation were compared. As expected, overall rates of degradation of these proteins were much slower (1–2%/h) and did not increase upon exposure to paraquat or cadmium (Fig. 2 A). Thus, the degradation of most cell proteins is quite resistant to these types of oxidative damage. It is also noteworthy that the magnitudes of the increases in degradation after exposure to paraquat, cadmium, 38°C, or 42°C (Figs. 1 A and 2 A) were similar. These findings suggest that after these insults, the maximal capacity of the degradative system had been reached. Accordingly, exposure of cells to these oxidants at 38°C did not further enhance proteolysis (unpublished data).

To confirm that the oxygen radicals stimulate only the degradation of newly synthesized proteins, we labeled cell proteins for 5 min during growth at 30°C, exposed a portion of the culture to paraquat or cadmium immediately or 60 min later, and then measured rates of degradation (Fig. 2 B). Addition of these ROS generators stimulated the breakdown of recently synthesized proteins. However, when the cadmium or paraquat was added 60 min later, the rates of proteolysis did not change (Fig. 2 B). The stimulation of proteolysis by this protocol was smaller than in Fig. 2 A, where the exposure to the oxidants was for 90 min, which should allow greater uptake of the oxidants. Because cycloheximide was present after the proteins were labeled, this resistance of proteolysis to the stimulation by ROS (Fig. 2) or heat shock (Fig. 1) after 1 h cannot be caused by the induction of proteins that protect against the oxidant or thermal damage. Instead, it probably reflects some maturational change in the newly synthesized proteins during this hour. Most likely, these insults can enhance degradation of newly synthesized proteins for only 1 h because, with time, cell proteins succeed in forming more stable multimeric complexes that are more resistant to ROS or thermal damage.

These findings raise two possibilities: either long-lived proteins are not chemically damaged by ROS or chemical damage occurs but does not trigger their unfolding and degradation, as it does to the recently synthesized proteins. To examine these alternatives, we assayed whether the long-lived proteins undergo oxidative damage using OxyBlot, which detects the presence of carbonyl groups in proteins (Sturtz et al., 2001). Exposure to paraquat for 90 min dramatically increased the presence of carbonyls in cell proteins and did so similarly, or even to a great extent, after treatment of the yeast with cycloheximide for 2 h to prevent the synthesis of new proteins and allow the degradation of short-lived polypeptides. Thus, the remaining long-lived cell proteins are certainly damaged by the oxygen-free radicals, although this modification fails to trigger rapid proteolysis (Fig. 3 A). The apparently greater appearance of protein carbonyls after cycloheximide treatment in Fig. 3 A is probably because some antioxidant enzymes are short-lived proteins (e.g., Sod1p).

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Figure 3.

Although long-lived proteins are not degraded more rapidly after exposure to paraquat, they are oxidatively damaged, and a lack of SOD1 also enhances the degradation of recently synthesized proteins. (A) Yeast were grown exponentially, and a portion of cells were treated with cycloheximide for 2 h to allow the degradation of short-lived proteins (the times studied in Fig. 2 A). Both the control and treated cells were then exposed to paraquat for 90 or 150 min. The presence of carbonylated proteins in equal amounts of cell proteins was assayed after derivitization with DNP-hydrazine (DNPH) and then Western blotting with an anti–DNP-hydrazone antibody. A control lane without the treatments with DNPH and paraquat is included to show the specificity of the antibody. Equal loading of lanes was shown with an eIF5A antibody. (B) WT, Δsod1 mutant, and the Δsod1 mutant expressing SOD1 from a plasmid were labeled with ³⁵S-Met for 5 min at 30°C. Rates of protein degradation were measured at 30°C as in Fig. 1 A.

Superoxide dismutase protects newly synthesized proteins against degradation

Prior studies on the effects of ROS on protein degradation used exogenous agents that cause free radical generation. However, oxygen radicals are also continually generated by cell metabolism. Cells have multiple enzyme systems to inactivate such radicals including the cytosolic Cu-Zn–containing superoxide dismutase 1 (Sod1p) and the mitochondrial Mn-containing superoxide dismutase 2 (Sod2p; Sturtz et al., 2001). Mutant strains lacking these enzymes, Δsod1 and Δsod2, accumulate oxidant-damaged proteins (Sturtz et al., 2001). To evaluate whether these enzymes help protect cell proteins against ROS-induced degradation and whether superoxide radicals can stimulate proteolysis, we compared rates of degradation in WT yeast and strains lacking either of these dismutases. After a 5-min pulse of ³⁵S-Met, newly synthesized proteins were degraded at least twice as rapidly in the Δsod1 strain as in the WT (Fig. 3 B), but by 1 h after synthesis, the rates of proteolysis resembled those in the WT. Thus, Sod1p appears to be important in protecting newly synthesized proteins from degradation induced by endogenously generated superoxide radicals.

Unlike the Δsod1 strain, the Δsod2 mutant showed very similar rates of degradation to those in isogenic WT cells (Fig. S1 A, available at http://www.jcb.org/cgi/content/full/jcb.200803022/DC1). Thus, the mitochondrial Sod2p is not important in the protection of newly synthesized proteins, presumably because these proteins are restricted to the cytosolic compartment. Upon addition of paraquat or cadmium to the Δsod1 strain, there was a further increase in proteolysis, and its capacity to degrade damaged proteins exceeded those in the paraquat- or cadmium-treated WT (Fig. S1 B).

Thermal and ROS-induced degradation involves the ubiquitin–proteasome system

Although early studies indicated a requirement for ATP for the degradation of oxidant-damaged proteins (Goldberg and Boches, 1982), subsequent studies, primarily in mature red cells, indicated that oxidant-damaged proteins could be degraded by an energy- and ubiquitin-independent process (Fagan et al., 1986; Davies and Goldberg, 1987; Matthews et al., 1989). This process was reported to involve degradation of the damaged proteins by 20S proteasomes (Grune et al., 1996; Shringarpure et al., 2003). To learn if the rapid elimination of these oxidant- and heat-damaged proteins requires the ubiquitin–proteasome pathway (like the degradation of misfolded proteins containing amino acid analogues), we tested whether degradation of these postsynthetically damaged proteins requires the ubiquitin-conjugating enzymes (E2s) UBC4 and UBC5 (Seufert and Jentsch, 1990) and also RPN10, a cytosolic ubiquitin binding protein that also functions as a subunit of the 26S proteasome's 19S regulatory subcomplex (Glickman et al., 1998).

Involvement of these components seemed likely because both Δrpn10 and Δubc4Δubc5 strains have been reported to show growth defects at 38°C and, in the presence of amino acid analogues (Seufert and Jentsch, 1990; Cagney et al., 2001; Elsasser et al., 2004), Δubc4Δubc5 mutant is also sensitive to cadmium (Jungmann et al., 1993). However, at the concentrations of paraquat and cadmium used here or after exposure to 38°C, these mutants grew at similar rates to the WT for several hours. Nevertheless, these mutants were defective in their capacity to degrade the oxidant- and heat-damaged proteins. After a 5-min pulse of ³⁵S-Met, degradation of short-lived and long-lived components at 30°C were not different in the WT and Δrpn10 strains, although degradation in the first hour was consistently slightly lower in the Δubc4Δubc5 double mutant, as was noted previously (Seufert and Jentsch, 1990). However, upon shift to 38°C (Fig. 4) or upon treatment with paraquat or cadmium (Fig. 5 A), both mutants failed to increase degradation of recently synthesized proteins (Figs. 4 and 5 A), which is in sharp contrast to WT cells.

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Figure 4.

Shift to 38°C stimulates the degradation of recently synthesized proteins in WT but not in Δrpn10 and Δubc4Δubc5 strains. Cells were labeled with ³⁵S-Met for 5 min at 30°C and shifted to 38°C or maintained at 30°C. Rates of degradation were measured as in Fig. 1 A.

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Figure 5.

Paraquat and cadmium fail to stimulate the degradation of recently synthesized proteins in Δrpn10 and Δubc4Δubc5 strains and these strains contain higher amounts of oxidant-damaged proteins than WT during growth and after exposure to paraquat. (A) WT and mutants were grown exponentially at 30°C and treated with paraquat or cadmium for 90 min. Then cell proteins were labeled with ³⁵S-Met for 5 min, and rates of degradation measured as in Fig. 2 A. (B) WT and mutants were grown exponentially, and a portion was exposed to paraquat for 90 min. The presence of carbonylated proteins in equal amounts of cell extracts was assayed by OxyBlot as in Fig. 3 A. To visualize oxidant damage in the control cells, approximately three times more protein was loaded than in cells treated with paraquat.

Thus, the selective degradation of oxidant- and thermally damaged proteins appears to involve the same components of the ubiquitin–proteasome pathway required for breakdown of proteins misfolded because of the incorporation of amino acid analogues (Seufert and Jentsch, 1990). Additional experiments using the autophagy-deficient mutant Δatg8 showed that autophagy is not involved in the degradation of heat damaged proteins (Fig. S2, available at http://www.jcb.org/cgi/content/full/jcb.200803022/DC1). This mutant showed a small reduction in breakdown of long-lived proteins which is catalyzed by vacuolar proteases (Lee et al., 1996). Further strong evidence that RPN10, UBC4, and UBC5 are important in the elimination of oxidant-damaged proteins came from analyzing extracts of these mutant cells by OxyBlot (Fig. 5 B). As predicted, the Δrpn10 and Δubc4Δubc5 cells during normal growth showed a higher content of protein carbonyl groups than WT cells, which increased markedly after exposure to paraquat for 1.5 or 2.5 h. These findings confirm that the ubiquitin–proteasome system is important in degradation of proteins damaged by endogenously generated as well as chemically induced ROS (Fig. 5 B).

The CDC48 ATPase and its partners ubiquitin fusion degradation (UFD) 1 and NPL4, but not UBC7 or UFD4, are essential for degradation of damaged proteins

CDC48, a member of the AAA family of hexameric ATPases (the homologue of mammalian p97), and its associated partners UFD1 and NPL4, have been previously implicated in the degradation of misfolded secretory and resident proteins by the ERAD pathway (Ye et al., 2001), as well as certain cytosolic ubiquitin fusion proteins by the UFD pathway (Ghislain et al., 1996). Interestingly, the expression of CDC48 is enhanced by cadmium (Vido et al., 2001). To test if it also functions in this breakdown of oxidant- and thermally damaged cytosolic proteins, we studied their degradation in WT and cdc48-3, ufd1-1, and npl4-1 strains upon shift to 38°C and exposure to paraquat or cadmium. At 30°C, rates of degradation of pulse-labeled proteins were slower in the cdc48-3 (Fig. 6 A), ufd1-1 (Fig. 7), and npl4-1 mutants (Fig. 8) and, upon shift to 38°C or exposure to cadmium and paraquat, a very large defect in the degradation of recently synthesized proteins was evident (Fig. 6 A, Fig. 7, and Fig. 8). However, the degradation of long-lived proteins in these mutants occurred at approximately similar rates as in the WT. Further evidence for an important role of CDC48 and its cofactors in elimination of oxidant-damaged proteins was obtained by OxyBlot analysis on the cdc48-3 mutant. During normal growth, this mutant contained a greater content of oxidant-damaged proteins than did the WT and, after exposure to paraquat for 90 min, the content of protein carbonyls was also much greater than in WT yeast (Fig. 6 B). The mutants of CDC48 or its associated partners UFD1 and NPL4 all showed similar defects in proteolysis, which further supports the conclusions that together they play an important role in the clearance of heat- and ROS-damaged proteins in a process also involving RPN10, UBC4, and UBC5.

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Figure 6.

Shifting cells to 38°C and exposure to paraquat or cadmium fail to stimulate the degradation of recently synthesized proteins in the cdc48-3 mutant, and the cdc48-3 strain contains higher amounts of oxidant-damaged proteins than WT during growth and after exposure to paraquat. (A) WT and cdc48-3 strains were labeled with ³⁵S-Met for 5 min at 30°C, and rates of degradation at 30 or 38°C were measured as in Fig. 1 A. To measure the degradation of oxidant-damaged proteins, WT and cdc48-3 strains were exposed to paraquat or cadmium for 90 min, followed by labeling with ³⁵S-Met for 5 min, and rates of degradation were measured as in Fig. 2 A. (B) WT and the cdc48-3 mutant were grown exponentially, and a portion was exposed to paraquat for 90 min. The presence of carbonylated proteins in equal amounts of cell proteins was assayed by OxyBlot as in Fig. 3 A.

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Figure 7.

Shifting cells to 38°C and exposure to paraquat or cadmium fail to stimulate the degradation of recently synthesized proteins in the ufd1-1 mutant. WT and ufd1-1 strains were labeled with ³⁵S-Met for 5 min at 30°C, and rates of degradation were measured at 30 or 38°C as in Fig. 1 A. To measure the degradation of oxidant-damaged proteins, WT and ufd1-1 strains were exposed to paraquat or cadmium for 90 min, followed by labeling with ³⁵S-Met for 5 min, and rates of degradation were measured as in Fig. 2 A.

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Figure 8.

NPL4 is required for increased degradation at 38°C and after exposure to paraquat or cadmium. WT and npl4-1 strains were labeled with ³⁵S-Met for 5 min at 30°C, and rates of degradation at 30 or 38°C and upon exposure to paraquat or cadmium were measured as in Fig. 6 A.

The E2 UBC7 plays an important role together with CDC48, UFD1, and NPL4 in the ubiquitination of misfolded ER resident proteins in the ERAD pathway (Hiller et al., 1996). To verify that the degradation of proteins damaged by heat and oxidative conditions involves proteins in the cytosol and is distinct from ERAD (as was suggested by its requirement for UBC4 and UBC5), we measured rates of degradation in the Δubc7 mutant upon shift to 38°C or exposure to paraquat or cadmium. A similar rapid increase in proteolysis was observed under these conditions in Δubc7 mutant as in WT cells (Fig. 9 A). Similar experiments tested whether the degradation of damaged cytosolic proteins is catalyzed by the UFD pathway. In Δufd4 mutants that lack the E3 ligase essential for the degradation of UFD substrates (Johnson et al., 1995), exposure to 38°C or to paraquat or cadmium caused a similar increase in breakdown of newly synthesized proteins to that in the WT (Fig. 9 B). Thus, this degradative process is distinct from the ERAD and UFD pathways, although all three require the same ATPase complex.

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Figure 9.

UBC7 and UFD4 are not required for increased degradation at 38°C and after exposure to paraquat or cadmium. (A) WT and Δubc7 strains were labeled with ³⁵S-Met for 5 min at 30°C, and rates of degradation at 38°C and upon exposure to paraquat or cadmium were measured as in Fig. 6 A. (B) WT and Δufd4 strains were labeled with ³⁵S-Met for 5 min at 30°C, and rates of degradation at 38°C and upon exposure to paraquat or cadmium were measured as in Fig. 6 A.

Protein damage and degradation induced by superoxide radicals

To examine the effects of ROS, we used paraquat and cadmium, because prior studies in yeast or mammalian cells had shown that these free radical generators cause oxidative damage to polypeptide chains. In addition, cadmium ions are known to cause induction of heat-shock proteins, the ubiquitin-activating enzyme E1, and the proteasome subunit PRE1, as well as CDC48 and SOD1, which we show are essential for the degradation of the damaged newly synthesized proteins (Vido et al., 2001). Although paraquat or cadmium stimulated the degradation of newly synthesized proteins, by 1 h after synthesis, their degradation could no longer be stimulated by these ROS generators.

Endogenous production of superoxide radicals by metabolism also can damage recently synthesized proteins and, as shown in this paper, cause their rapid degradation. The mutant strains lacking SOD1 showed a very similar large increase in their degradation as was induced with paraquat or cadmium (Fig. 3 B). In other words, SOD1, by removing damaging superoxide radicals, seems to enhance the chances that newly synthesized proteins can assume their final conformations. Thus, SOD1, by eliminating this highly reactive species, functions together with the ubiquitin–proteasome pathway to prevent the accumulation of oxidant-damaged polypeptides and to help cells withstand the consequences of increased levels of ROS, as occurs in mammals with neurodegenerative disease and inflammation, as well as during normal aging.

Most likely, these modifications by themselves are not recognized by the degradative system but, instead, these modifications lead to unfolding or irreversible denaturation of the newly synthesized polypeptides. Accordingly, a large fraction of the thermally and oxidant-damaged proteins that were not degraded in the Δubc4Δubc5, Δrpn10, and cdc48-3 mutants accumulated as aggregates in rapidly sedimenting fractions (Fig. 10, A and C; and Fig. S3). These observations also suggest that the oxidant-damaged proteins are recognized by their ability to cause unfolding of newly synthesized proteins. Resistance of the long-lived proteins to ROS-induced degradation is not because they fail to react with the ROS. On the contrary, under conditions where no newly synthesized proteins were present (after exposure to cycloheximide for 2 h), exposure to paraquat caused as much or even more oxidant damage to cell proteins (appearance of carbonyl groups) as in control cells (Fig. 3 A). However, after such oxidant damage, the chemically modified long-lived proteins seem not to denature because in the degradation-deficient mutants, these proteins did not accumulate in the particulate fraction, which is in contrast to the oxidant-damaged newly synthesized proteins which tend to aggregate when not degraded.

Measurements of total protein degradation

Yeast cells were grown exponentially to OD₆₀₀ 0.8–1 at 30°C in SD medium supplemented with all amino acids (Sigma-Aldrich). 10 ml of cells were collected by centrifugation and resuspended in 2 ml of the same media, but without methionone, for 50 min. Then the cells were incubated with 50 μCi/ml ³⁵S-Met for 5 min to label recently synthesized proteins or 90 min to label long-lived proteins. After two washes with ice-cold chase media at 4°C, cells were resuspended in 2 ml chase media with a high concentration of Met (6 mg/ml) and cycloheximide (0.5 mg/ml) to prevent reutilization of ³⁵S-Met. At different time intervals, equal aliquots (450 μl) of cells were taken and mixed with 50 μl of 100% TCA to a final concentration of 10%. After incubation at 4°C overnight, the samples were centrifuged at 14,000 rpm for 10 min at 4°C, and the radioactivity in both the TCA-soluble and -insoluble fractions was measured by suspending the pellets in 4 ml of scintillation fluid (PerkinElmer). The rate of protein degradation is expressed as the percentage of initially incorporated radioactivity converted into acid-soluble fragments each time. The TCA-soluble cpm at time zero was substrated from each subsequent measurement. To measure the effects of temperature shift, cells were shifted to 38, 42, or 20°C during the chase period with chase media preadjusted to the respective temperatures. To measure the effect of 80 μg/ml paraquat or 50 μg/ml cadmium, cells were exposed to these agents for 90 min before the labeling with ³⁵S-Met, except in Fig. 2 B where these agents were added after the pulse of ³⁵S-Met. TCA-soluble and -precipitable cpm for typical experiments (Figs. 1 and ​and2)2) are shown in Tables S1 and S2 (available at http://www.jcb.org/cgi/content/full/jcb.200803022/DC1).

Detection of carbonyl groups in proteins

To detect the presence of carbonyl groups in proteins, yeast cells were grown exponentially at 30°C in YPD media. 20 ml of cells were then exposed to 80 μg/ml paraquat for 90 min. Cells were collected by centrifugation and lysed with glass beads in the presence of ice-cold sorbitol buffer (700 mM sorbitol, 50 mM Tris-HCl, pH 7.5, and 1 mM PMSF) at 4°C. Cell debris was removed by centrifugation at 2,000 g for 5 min at 4°C. Equal amounts of cell proteins were derivatized to yield 2,4-DNP-hydrazones by incubation with 2,4-DNPH for 15 min at room temperature. By using the OxyBlot detection kit (Millipore), proteins were separated by SDS-polyacrylamide gel electrophoresis, and the derivatized carbonyl groups were detected with an anti–DNP-hydrazone–specific antibody (Millipore).

Measurement of radiolabeled proteins in the ultracentrifugal pellets

Yeast cells were grown exponentially at 30°C, and 20 ml of cells were labeled with ³⁵S-Met for 5 min, washed, and resuspended in chase media containing cycloheximide and high concentrations of Met (as described in Measurements of total protein degradation). Cells were shifted to 38°C or exposed to paraquat, and aliquots were collected at the indicated time points. Lysates were prepared by breaking the cells with glass beads in the presence of ice-cold sorbitol buffer at 4°C. Cell debris was removed by centrifugation at 2,000 g for 5 min. Lysates containing equal amounts of proteins were subjected to differential centrifugation. The amount of radiolabeled proteins present in the pellets was obtained by centrifuging for 20 min at 9,300 g, and after ultracentrifugation at 275,000 g for 60 min, they were measured by suspending the pellets in scintillation fluid. The accumulation of the ³⁵S-Met protein in these fractions was expressed relative to the total amount of the radiolabeled protein at time zero. The presence of carbonyl groups in the 275,000 g pellet was detected by OxyBlot analysis.

We thank D. Finley, T. Rapoport, and V. Culotta for providing yeast strains and plasmids, D. Finley, O. Kandror, and J. Brault for helpful suggestions, and M. Dethavong for valuable assistance.

This research was supported by grants from the National Institutes of Health (GM5123-12), the Fund for Innovation from Elan, Inc., and the Ellison Medical Foundation.