Dependence of Proteasome Processing Rate on Substrate Unfolding^*

Yeast Strains, Metabolic Labeling, and Pulse-chase Analysis

Saccharomyces cerevisiae strain MHY501 (MATα his3-Δ200 leu2–3,112lys2–801 trp1–1 ura3–52) (14) was used as the genetic background for construction of wild type (YAH96) and rpn10Δ (YAH106) strains bearing a C-terminal 3xFLAG epitope at the RPN11 locus for affinity purification of yeast proteasomes (15). Test proteins were expressed from p416ADH-based vectors (16) in strain MHY501. Details of plasmid construction will be provided upon request. Pulse-chase analysis of test proteins in yeast transformants was carried out as described previously (17). Immunoprecipitations were carried out using anti-Flag (M2) affinity gel (Sigma-Aldrich). Autoradiographs were quantified using TotalLab software (Nonlinear Dynamics).

Protein Purification

Substrates containing an Rpn10 domain were expressed in Escherichia coli BL21 Rosetta strains (Novagen) as N-terminal hexahistidine-tagged proteins. Unlabeled forms were induced when host strains reached A₆₀₀ ∼1 with 0.5 mm IPTG at 30 °C overnight (16–18 h). ³⁵S-radiolabeled substrates were prepared and purified as follows. Cells were grown to A₆₀₀ ∼0.7 in LB medium containing 30 μg/ml kanamycin and 34 μg/ml chloramphenicol. After harvesting, cells were resuspended to A₆₀₀ ∼3 in M9 medium supplemented with 0.063% methionine assay medium (Difco #242310), 0.4% glucose and 30 μg/ml kanamycin and incubated at 37 °C for 1 h. 1 mm IPTG was then added. After 30 min incubation at 30 °C, [³⁵S]methionine and [³⁵S]cysteine (as EasyTag protein labeling mix (Perkin Elmer)) were added to 14 μCi/ml as well as 34 nm of unlabeled methionine and incubation continued for 1 h. The cells were then harvested and the pellet frozen at −80 °C. Labeled and unlabeled proteins were purified using Talon metal affinity resin (Clontech) according to the manufacturer's instructions. Radio-labeled proteins were eluted from Talon resin and concentrated on 10,000 molecular weight cutoff Centricon concentrators (Amicon Ultra filter devices). Substrate buffer was exchanged on BioSpin chromatography columns (Bio-Rad) into HRV Cleavage Buffer (25 mm Tris, pH 8.0, 150 mm NaCl, 2 mm DTT, 10% glycerol). After elution, the unlabeled proteins were dialyzed into HRV Cleavage Buffer. Vector-derived N-terminal sequences were removed from all substrate proteins by cleavage with 100 units HRV protease per mg of substrate protein (Turbo 3C Protease, Accelagen) at 4 °C for 24 h. HRV protease and residual uncleaved substrate proteins were removed by subsequent incubation with Talon resin.

Proteasome Purification

26 S proteasomes were purified from strains YAH96 or YAH106 in which a 3xFLAG tag has been appended to the C terminus of the Rpn11 subunit of the proteasome. Cells were grown to an A₆₀₀ of 3–4 in 1.5 liters of YPD, harvested, washed once with 50 ml of ice-cold water. The cell pellet was then resuspended in an equal volume of Proteasome Buffer (PB: 50 mm Tris pH 7.5, 25 mm NaCl, 5 mm MgCl₂, 2 mm ATP, 10% glycerol) supplemented with 2× Complete EDTA-free protease inhibitor mixture (Roche Diagnostics) and lysed by five passes through a Microfluidics microfluidizer at a pressure of 100 psI. Lysates were clarified by centrifugation at 20,000 × g at 4 °C for 30 min. 26 S proteasomes were affinity-purified by incubation with M2 anti-FLAG affinity gel (Sigma) for 1 h at 4 °C. Bound proteasome complexes were washed with 200 column volumes of PB and eluted from the resin overnight at 4 °C in an equal volume of PB supplemented with 0.2 mg/ml 3xFLAG peptide (Sigma). The eluates were concentrated on 10,000 molecular weight cuttoff Centricon concentrators. Proteasome composition was determined by denaturing gel electrophoresis followed by Coomassie staining. Complex assembly was assessed by native gel electrophoresis (18). Proteasome molarity was calculated using a molecular weight 2,400 kDa.

In Vitro Degradation Assay

Degradation reactions containing radiolabeled protein substrates were carried out at 30 °C by 50 nm 26 S proteasome in a buffer (25 mm HEPES pH 7.5, 100 mm KCl, 20 mm MgCl₂, and 10% glycerol) containing 2 mm DTT, 5 mm ATP, and an ATP regenerating system (3 mm phospho(enol)pyruvic acid, 2 mm NADH, and 0.25 units of pyruvate kinase/lactate dehydrogenase mix (Sigma)). Substrates were present at the concentrations specified in the text. Degradation was initiated by substrate addition and time-dependent degradation was assessed by periodically removing 10 μl volumes from the reaction and precipitation in 140 μl of 20% TCA for 30 min on ice. After 30 min centrifugation at 18,000 × g, the supernatant, containing the peptide products of proteolysis, was added to a scintillation vial for to quantification of radioactivity. For each experiment, the total radioactivity present in substrate was determined by adding an aliquot of the degradation assay mixture to TCA, transferring the solution to a scintillation vial and counting. Kinetic parameters were determined using the Michaelis-Menten equation by fitting the data to a hyperbolic function, using the SigmaPlot software package. For SDS-PAGE analysis of degradation, aliquots were periodically removed from the degradation assay reaction. Samples were resolved on a 12% SDS-PAGE and the distribution of radiolabeled proteins on the dried gel determined by autoradiography. Signals from autoradiographic films were quantified using TotalLab software (Nonlinear Dynamics).

ATPase Assays

Proteasome ATPase activity was measured in a coupled assay (19, 20) in 25 mm HEPES (pH 7.5), 100 mm KCl, 20 mm MgCl₂, 10% glycerol, 250 mU/ml LDH/pyruvate kinase (Sigma), 7.5 mm phospho(enol)pyruvic acid, 1 mm NADH, 2 mm DTT, and 5 mm ATP. Proteasome concentration was 30 nm and, when present, substrates or other proteins containing an Rpn10 moiety were 1.5 μm. ATPase activity was followed by loss of NADH absorbance at 340 nm. ATP consumption was calibrated using an NADH standard curve, performed with each assay. Reactions were carried out in 96-well plates in a Molecular Devices Spectra Max spectrophotometer. Rates were calculated from least squares fits of each time series. Data were collected separately for: 1) each substrate or pseudo-substrate alone or 2) while co-incubating each substrate or pseudo-substrate with proteasomes or 3) for proteasomes alone. Proteasome-specific ATP hydrolysis rates were calculated as (proteasomes + substrate rate) − (substrate-only rate).

Design and Structure of Substrates Used in These Studies

Our previous studies of the native ubiquitin-independent proteasome substrate ornithine decarboxylase (ODC) provided the basis for the design of the substrates we generated and utilized. No post-translational modification of ODC is needed for its proteasomal degradation; in particular, ubiquitin is not involved (21). Vertebrate forms of ODC have a small conserved degradation tag (22) (37 amino acids in mouse and human) at its C terminus (cODC). Degradation is strictly unidirectional and begins at the tag (5). Fusing this tag to other proteins promotes their rapid turnover by the eukaryotic proteasome (17), including those of mammals, plants, and fungi.

Proteasomal degradation, whether dependent on (23) or independent of ubiquitin (24), requires two distinct substrate properties: proteasome association and an unstructured element at which degradation begins. Both of these functions are provided by the cODC degron. Mutating a critical cysteine residue (C441) in the mouse cODC sequence prevents proteasome association (25). However, proteins stabilized by a Cys-441 mutation could be restored as proteasome substrates by appending them to the Rpn10 subunit of the proteasome (24). Such fusions bind at the proteasome site which normally accommodates Rpn10. Once tethered in this way these chimeric proteins were degraded only if they also provided an unstructured element of some minimal length. Here we used similarly designed fusions to deliver domains of varying mechanical stability to the proteasome within a uniform context, and thus determined the effect of this parameter on turnover. Fig. 1 shows diagrammatically the composition of the substrates used in these studies.

Titin I27 Degradation in Yeast Cells

The domain we first utilized for analysis of unfolding was the I27 module of the giant titin protein, which is composed of more than two hundred individually folded domains connected by unstructured linkers. I27 provides several advantages. Its mechanical unfolding has been extensively investigated by biophysical methods and is among the most stable proteins that have been studied by these means (26–28). It thus presents a formidable challenge to a biochemical unfoldase. Importantly, single residue mutations have been characterized that do not significantly alter the structure of I27 but do reduce the force required for mechanical unfolding. We compared wild-type I27 and a destabilized mutant containing a single residue mutation, V13P. Compared with the wild-type form, I27^V13P requires less pulling force for unfolding (29).

We first asked whether proteasomes could degrade I27 in living cells and whether the V13P mutation altered turnover. Because the ubiquitin independent cODC tag can direct degradation in yeast (17), we appended that degron to the C terminus of I27 and I27^V13P (Fig. 1) and expressed these proteins in Saccharomyces cerevisiae. Cells were incubated with radio-labeled amino acids and pulse-chase analysis used to measure turnover (Fig. 2, A and B). During a 1-h chase period more than 90% of the labeled I27^V13P-cODC was degraded during the first 30 min of the chase, and its turnover appeared to be first order, with a half-life of about 6 min. At the 60 min time point, residual protein was below our detection limit. In marked contrast, the wild-type I27 form showed little degradation during the 60 min chase period; at most, 20–30% of the labeled I27-cODC present at the beginning of the chase disappeared. We conclude that, as for other proteins, the cODC degron can direct degradation of I27, and that a destabilizing mutation significantly accelerates its turnover.

Although cODC is an effective degron in vivo, it has not in our hands proven reliably effective for use with purified yeast proteasomes. For biochemical analysis we therefore chose an alternative means to target substrates to yeast proteasomes, which depends on fusion to the Rpn10 protein, as previously described (24) and further developed here. Before exploiting that approach in vitro we tested it in yeast cells, again asking whether these fusions provided an effective delivery vehicle for I27 and whether the wild type and mutant form of the folded domain were degraded differently. R-I27 was expressed in yeast, with or without the V13P mutation. As a control for specificity, these were expressed either with or without an unstructured C-terminal extension (see Fig. 1 for diagram of substrate structures). This consisted of cODC^C441S, which has been shown to support proteasomal degradation of tightly folded Rpn10-tethered proteins in yeast (24). Pulse-chase analysis was used, as above, to assess turnover (Fig. 2, C and D). Without a C-terminal extension we observed no degradation of either wild-type I27 or its destabilized V13P mutant form. When the extension was present, both forms of I27 were degraded. However, the rate and extent of degradation of the two was very different, and closely matched the results observed using the cODC degradation tag. Only 20–30% of labeled substrates containing wild-type I27 were degraded during a 2 h chase. In contrast the I27^V13P substrate that was otherwise identical, but for a single residue mutation, was degraded with a half-life of about 15 min, and none was detected after 2 h. These data show that distinct means of targeting substrates to proteasomes (cODC, a native eukaryotic degron, and fusion to Rpn10, an endogenous mechanism for delivery of ubiquitin-conjugated proteins (30, 31)) give concordant results in yeast cells. We therefore determined unfolding kinetics of Rpn10 tethered substrates by yeast proteasomes in vitro.

Titin I27 Degradation by Purified Proteasomes

Because in vitro experiments can be performed with purified components at known concentrations, they make it possible to calculate kinetic parameters, such as absolute turnover rates, rather than relative rates, as in the in vivo experiments just described. We therefore affinity purified yeast proteasomes and generated recombinant substrates in bacteria with the same structure (Rpn10-“folded domain”-extension) as those used with intact yeast cells. Purified proteasomes were analyzed by SDS-PAGE and by native gel electrophoresis (supplemental Fig. S1). These data confirmed that proteasomes had the expected polypeptide composition and that >95% were doubly capped, with two 19 S regulatory complexes per 20 S proteolytic core complex. SDS-PAGE confirmed that the substrates were homogeneous and had the expected electrophoretic mobility (supplemental Fig. S1).

We initially analyzed a substrate pair containing the I27 domain or its V13P mutant; each produced with or without a C-terminal extension (see Fig. 1). We monitored degradation in reactions with 50 nm 26 S proteasomes and 250 nm radio-labeled Rpn10-tethered proteins, measuring the production of acid soluble radiolabeled peptide products (Fig. 3A). (In initial experiments we compared isogenic Rpn10 and rpn10Δ proteasomes; the results were similar (Ref. 24 and supplemental Fig. S2). We subsequently used only wild-type proteasomes). Time-dependent progress curves were qualitatively similar to the data observed with the cognate substrates in vivo. No degradation occurred in the absence of a C-terminal extension. Both I27 substrates with extensions were degraded, but the rate and extent of destruction of the destabilized mutant was about 8–10-fold greater. These results show that relative degradation rates in our in vitro system match those observed in intact cells.

DHFR

We next asked whether these observations and conclusions could be extended from I27 to other proteins. Mammalian dihydrofolate reductase (DHFR) is well-suited for testing the generality of our findings. DHFR, like I27, has a well characterized structure, the stability of which can be experimentally altered. The antimetabolite methotrexate binds tightly to DHFR and stabilizes its structure, a conclusion supported by numerous biochemical (7, 32–34) as well as biophysical (35) studies. We investigated the degradation of Rpn10-tethered DHFR by purified proteasomes (Fig. 3B). Again we compared a series of substrates: an extension was present or absent, and methotrexate was present in or omitted from the reaction. The results, although similar to those seen with I27, were quantitatively different. An extension was not an absolute requirement for degradation, but its presence accelerated degradation. Regardless of the presence or absence of an extension, the stabilizing ligand methotrexate slowed degradation.

For both DHFR and I27 tethered substrates, we performed two tests of degradation specificity. First we asked whether the Rpn10 moiety of substrates determines proteasome association by adding a fifty-fold excess of Rpn10-GFP as competitor. (Rpn10-GFP can functionally replace Rpn10, but is not an active substrate because it lacks an unstructured extension (24)). The rate of degradation of substrates was reduced more than 10-fold by Rpn10-GFP (supplemental Fig. S3), demonstrating that Rpn10 binding is required for substrate activity in this system. We also tested whether the proteasome rather than an adventitiously co-purified protease was responsible for substrate degradation by preincubating reactions with the highly specific proteasome inhibitor epoxomicin (36). Epoxomicin (200 μm) reduced degradation by more than 95% compared with control reactions without the inhibitor.

Substrate Degradation Kinetics

We performed kinetic analysis using proteasomes and the substrates described above. For all substrates tested, degradation rates were determined using 50 nm proteasomes and reaction conditions that produced linear initial velocities (representative primary data is shown in supplemental Fig. S4). The resultant substrate concentration-versus-velocity data were well-approximated by a hyperbolic function, a result descriptive of Michaelis-Menten kinetics (Fig. 4). Despite their great structural complexity, proteasome activity is approximated quite well by classical Michaelis-Menten kinetics, a property commonly observed with simpler enzymes. Inspection of the kinetic parameters derived from curve fitting (Table 1) reveals that the four substrate conditions yielded broadly similar K_D values, indicating that the variable C-terminal domain in these substrates does not significantly perturb the Rpn10-proteasome interaction. In contrast, the V_max values of the various substrates were more diverse. The V_max values exhibit a 4.8-fold faster turnover rate of the destabilized I27^V13P substrate compared with its wild-type counterpart and a 2.7-fold slower turnover rate of the DHFR substrate when associated with a stabilizing ligand.

Proteasome ATPase Activity

We also determined the ATP hydrolysis activity of proteasomes and asked whether substrate processing affected that activity (Table 2). The rate of ATP turnover was ∼110/proteasome/min. and varied modestly among several independently prepared lots. The effect on ATP hydrolysis of incubating proteasomes with Rpn10-tethered substrates or with pseudo-substrate controls (those lacking a C-terminal extension) was examined. Reactions were carried out under the same conditions as used for degradation, but the added proteins were present at 1.5 μm, 5–10-fold above the substrate K_m values. The measured rates therefore reflect ATPase activity in the presence of near-saturating substrate. The apparent stimulation of ATPase activity by substrates and pseudo-substrates was at best modest. The Rpn10 tethered proteins produced ∼10–30% stimulation above basal rates (Table 2; see supplemental Fig. S5 for primary data). The modest stimulation of ATPase activity observed is close to the boundary of experimental error. Furthermore, this is unlikely to be physiologically significant for substrate degradation because of its small magnitude and the lack of systematic correlation of the extent of stimulation with the properties of Rpn10-tethered proteins important for their degradation: domain stability and the requirement for an unstructured extension.

What Limits the Turnover Rate of More Slowly Degraded Substrates?

The kinetic rate parameter can be expressed as the catalytic constant, k_cat, or as its reciprocal, the time required at steady state for a proteasome to complete the reaction cycle (Table 1). The steady-state turnover time of the wild-type I27 substrate is 40 min. What is status of the substrate-proteasome pair during this prolonged period? It is likely that there are multiple iterations of unfolding attempts associated with cycles of ATP hydrolysis, each with a low probability of success. One important question is whether substrate and proteasome remain continuously associated during this process. To test this, we initiated degradation of radio-labeled R-I27-ext, then after 30 min added a 10-fold excess of the same substrate, but in unlabeled form (Fig. 5, arrow). Before addition of the unlabeled competitor, radiolabeled substrate was present at a concentration of 250 nm, in excess of the K_m value for this substrate, which is 145 nm. Hence proteasomes were more than half saturated. Addition of competitor immediately and markedly inhibited the subsequent generation of labeled proteolysis products. In contrast, under the control condition without competitor, production of labeled peptides continued unabated (Fig. 5). This result implies that the substrate must undergo a rapid on/off process. Excess competitor quickly re-equilibrates proteasomes with substrate that is 11-fold less radioactive and the generation of labeled proteolysis products slows commensurately. The rapid inhibitory effect of adding excess unlabeled substrate is inconsistent with stable substrate association and clearance of substrate from proteasomes solely by proteolysis: the mean time for that clearance mechanism exceeds 30 min. Therefore, the Rpn10-tethered substrate undergoes a rapid on/off process that may contribute to the measured duration of steady-state degradation.

Are Stability-dependent Rate Differences an Artifact of Interrupted Degradation?

Lastly, we considered whether non-processive degradation could have artifactually distorted the turnover times reported here. We (6, 37) and others (8, 38) have shown that a tightly folded protein domain can halt degradation, producing partial degradation products with termini containing short extensions of the folded domain. Such partial degradation products could act as competitive inhibitors. SDS-PAGE and autoradiography was used detect the production from R-I27-ext of terminally trimmed products. Reactions were performed as in Fig. 3A; incubation without addition of proteasomes was done in parallel to control for non-proteasomal degradation. We observed time- and proteasome-dependent disappearance of the substrate band (supplemental Fig. S6). Densitometry showed that less than 20% of the radiolabeled substrate initially present accumulated as intermediates; the rest was degraded to peptide fragments not resolved by SDS-PAGE and autoradiography. Therefore, both R-I27-ext and R-I27^V13P-ext are predominantly degraded processively and non-processive outcomes did not markedly alter the reported kinetic parameters.