Research Article

Substrate degradation by the proteasome: A single-molecule kinetic analysis

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Science  10 Apr 2015:
Vol. 348, Issue 6231, 1250834
DOI: 10.1126/science.1250834

Single-molecule assay of ubiquitylation

Many biological processes in cells are regulated by ubiquitin peptides that are attached to proteins. Measurement of single fluorescent molecules in cell extracts can be used to trace the kinetics of such reactions. Lu et al. refined assay conditions to follow ubiquitination by an E3 ubiquitin ligase (see the Perspective by Komander). They visualized the activity of the anaphase-promoting complex (APC), a ubiquitin ligase critical for control of the cell division cycle. The processive initial reaction catalyzed by the APC was replaced by slower reactions. The results show how small, commonly occurring recognition motifs can guide specific and highly controlled enzymatic events. In a companion paper, Lu et al. explored how the number and arrangement of added ubiquitin chains affected the interaction of ubiquitylated proteins with the proteasome (a protein complex that recognizes ubiquitylated proteins and degrades them). The extent of ubiquitylation determined the strength of interaction of a substrate protein with the proteasome, and the arrangement of the ubiquitin chains determined the movement of the protein into the proteasome and thus the rate of degradation.

Science, this issue 10.1126/science.1248737, 10.1126/science.1250834; see also p. 183

Structured Abstract


Protein degradation, mediated by the ubiquitin-proteasome system (UPS), plays a critical and complementary role to transcription, splicing, and translation in the control of gene expression. Effective regulation of the UPS relies on the specificity of substrate recognition, which is conferred largely by the upstream ubiquitylation process. As a specific example, the anaphase-promoting complex (APC) acting on a series of substrates promotes waves of ubiquitylation and degradation, leading to key transitions in the cell cycle. However, in this cascade the proteasome itself also plays a role in the specificity of degradation. There have been various efforts to characterize this role, leading to a commonly held view that a substrate protein must be conjugated with a chain of at least four ubiquitins to be recognized by the proteasome. Yet mass spectrometry studies show that ubiquitin chains on APC substrates (such as cyclin B) contain, on average, only two ubiquitins. But these can have complex ubiquitin configurations created by the multiplicity of ubiquitylated lysine residues. Therefore, the tetraubiquitin chain selection rule may not be generally applicable, and the mechanism by which the proteasome recognizes substrates is still shrouded in mystery.


The multiple lysines on a substrate and on ubiquitin itself generate a large number of possible ubiquitin configurations. To understand the “ubiquitin code” that must be read by the proteasome and converted into a rate of substrate degradation, we examined the kinetics of degradation with defined ubiquitin configurations by conjugating preformed ubiquitin chains on substrate molecules. Further, to reveal the molecular basis through which some ubiquitin configurations promote more efficient degradation than others, we investigated the degradation process using single-molecule (SM) methods that are capable of identifying transient intermediates and measuring their kinetic parameters and sensitivity to ubiquitin configurations.


Contrary to the tetraubiquitin chain selection rule, we find that for APC substrates with multiple ubiquitylated lysine residues, diubiquitin chains are more efficient than tetraubiquitin chains in promoting degradation, given the same number of conjugated ubiquitins. Ubiquitin chains are essential for degradation of most substrates. Nevertheless, a multiple monoubiquitylated form of securin, a regulator of chromatid separation, interacts with the proteasome as strongly as securin containing the same number of ubiquitins grouped in chains. By dissecting the degradation process using SM methods, we find that ubiquitin chain structures on substrates promote the passage of a bound substrate into the translocation channel on the proteasome.


This systematic study of synthetically constructed ubiquitylated substrates with defined configurations revealed no simple length threshold for ubiquitin chains for degradation by the proteasome. A distributed array of short ubiquitin chains, as appears naturally on APC substrates, is a superior and perhaps optimal signal for degradation; this conclusion will most likely extend to substrates of other E3 ligases. The rate of degradation is an aggregate of two sequential processes: substrate binding and kinetic postbinding events. In the past, it was widely assumed that ubiquitin chains mostly promoted binding to the proteasome. Our SM studies demonstrate that the strength of interaction with the proteasome, for substrates with multiple ubiquitylated lysines, is largely determined by the total number of ubiquitins and is less sensitive to ubiquitin chain configurations. For most substrates, binding alone is not sufficient for degradation. Rather, degradation depends strongly on a process that initiates passage into the substrate translocation channel; this transition, in contrast to binding, is determined by the particular configuration of ubiquitin chains.

Key transitions in the degradation of ubiquitylated substrates by the proteasome.

A polyubiquitylated substrate molecule explores multiple configurations on the proteasome through stochastic binding. Rearrangements of proteasomal subunits, driven by the binding and hydrolysis of adenosine triphosphate, allow a deeper engagement of the substrate with the proteasome, when there are appropriate ubiquitin chain structures on the substrate. As a result of this engagement, the substrate or its terminus is moved closer to the substrate entry port, promoting expeditious initiation of translocation and ensuing degradation.


To address how the configuration of conjugated ubiquitins determines the recognition of substrates by the proteasome, we analyzed the degradation kinetics of substrates with chemically defined ubiquitin configurations. Contrary to the view that a tetraubiquitin chain is the minimal signal for efficient degradation, we find that distributing the ubiquitins as diubiquitin chains provides a more efficient signal. To understand how the proteasome actually discriminates among ubiquitin configurations, we developed single-molecule assays that distinguished intermediate steps of degradation kinetically. The level of ubiquitin on a substrate drives proteasome-substrate interaction, whereas the chain structure of ubiquitin affects translocation into the axial channel on the proteasome. Together these two features largely determine the susceptibility of substrates for proteasomal degradation.

The propensity of a protein for degradation is largely encoded in its state of ubiquitylation (1, 2). The ubiquitylation process results in highly diverse configurations of ubiquitin chains on target proteins (3). The 26S proteasome recognizes ubiquitylated substrates and degrades them into short peptides (4). Tests of defined configurations of ubiquitins on substrates for their influence on recognition and degradation showed that a substrate protein must generally be conjugated with a chain of at least four ubiquitins to interact tightly enough with the proteasome for degradation (5). However, no compelling evidence supports the existence of a tetraubiquitin chain receptor on the proteasome. Alternative explanations for the tetraubiquitin chain selection rule mostly rely on the geometric distance between a pair of proteasomal subunits to gauge the length of a ubiquitin chain (6, 7). The requirement for a tetraubiquitin chain for degradation also differs from substrate to substrate, without a predictable relationship to a substrate’s function or structure (811).

Exit from mitosis and passage into the G1 phase of the cell cycle requires ubiquitylation mediated by the anaphase-promoting complex (APC) (12). Substrates (such as cyclin B, securin, and geminin) typically contain multiple lysine residues, to which ubiquitin moieties are conjugated, providing a very large number of possible combinations of ubiquitin chain configurations (3, 8, 13, 14). Mass spectrometry studies indicate that ubiquitin chains on cyclin B molecules generated in reconstituted reactions contain, on average, only two ubiquitins (3). Moreover, multi-monoubiquitylation (referring to ubiquitylation on multiple lysine residues without chain formation) on cyclin B leads to efficient degradation. Multiple ubiquitylation sites are commonly found on substrates of other E3 enzymes (15, 16); at least 56% of ubiquitylated human proteins contain more than one ubiquitylation site, though the relevance of these ubiquitylations to protein degradation has not been fully demonstrated (17).

Ubiquitins conjugated to the substrate promote interaction with the proteasome; however, binding by itself is not sufficient to initiate degradation. Several known proteasome-interacting proteins (Usp14, Rad23, etc.) are stable (18), and autoubiquitylated Cdc34 is not degraded, despite its strong interaction with the proteasome (19). To facilitate initiation of substrate translocation, it has been proposed that substrates must have an unstructured terminal region of at least 30 amino acids (20); multiple steps occur in the proteasomal degradation process, as suggested by cryo–electron microscopy (cryo-EM) structures (6, 7, 21). Before or coincident with peptide translocation, conjugated ubiquitins are removed by deubiquitylating enzymes (DUBs) Rpn11, Usp14, and Uch37 on the proteasome (22). Whereas Usp14 and Uch37 may have an editing role to tune the rate of proteasomal degradation, they are dispensable for the degradation process. The DUB Rpn11, by contrast, is required for efficient proteasomal activity. Rpn11 is located close to the substrate entry port, where it removes ubiquitin chains en bloc from the translocating peptide (6, 23). It is unclear how the proteasome might use these features to establish selectivity in substrate recognition.

To understand the requirements for efficient protein degradation, we examined the kinetics of degradation of substrates with defined ubiquitin configurations. We found that, for APC substrates with multiple ubiquitylated lysine residues, tetraubiquitin chains were not required for efficient degradation. Rather, given the same number of total ubiquitins on a substrate molecule, diubiquitin chains were more efficient than tetraubiquitin chains in promoting degradation. To elucidate the molecular basis through which some ubiquitin configurations promote more efficient degradation than others, we investigated the intermediate steps in the degradation pathway using single-molecule (SM) methods. For substrates containing multiple lysines, the strength of interaction with the proteasome was determined largely by the total number of ubiquitins and was less sensitive to the ubiquitin configuration. However, substrate binding alone was not sufficient for rapid degradation for most substrates. Rather, degradation depended strongly on the initiation rate of passage into the substrate translocation channel, and this transition was promoted by the presence of ubiquitin chain structures on substrates.


Degradation of defined multiple ubiquitylated substrates

Most, if not all, APC substrates contain multiple ubiquitylatable lysine residues. To reduce the complexity of these mixed configurations, we sought to control the number, length, and linkage of conjugated ubiquitin chains on wild-type (WT) substrates. Purified APC was used as the ubiquitin ligase to conjugate preformed di- or tetraubiquitin chains of Lys48 (K48) linkage, which support most proteasome-mediated degradation in cells (24) (Fig. 1A). The chains were methylated on lysines before conjugation to prevent secondary elongation. We then separated the reaction products having different numbers of conjugated ubiquitins by electrophoresis. This approach limits the heterogeneity of ubiquitin configurations on the substrate to combinations of sites accepting a known number of defined ubiquitin chains. We measured the degradation rate of the ubiquitylated substrate for each electrophoretically resolved species by exposure to purified human 26S proteasomes that were free of reversibly associating ubiquitin receptors, such as Rad23 (Fig. 1A). Success of this strategy required the absence of interconversion of different ubiquitylated species generated by partial deubiquitylation by the proteasome; this was ensured by removing the DUB Usp14 on the proteasome by salt wash (25). Usp14-free proteasomes are known to efficiently degrade substrates, such as cyclin B and Sic1, without generating partially deubiquitylated products (fig. S1) (8, 25, 26). Another DUB on the proteasome, Uch37, does not appreciably deubiquitylate the substrates used here (25). Controls using a general DUB inhibitor (not active against Rpn11) further confirmed that DUB-driven interconversion of ubiquitylated species was unlikely (fig. S2).

Fig. 1 Quantitative degradation assay.

(A) The assay strategy. Preformed, methylated ubiquitin chains were conjugated to PKA-labeled securin using purified APC and E2 UbcH10. After reaction, the product was subject to degradation by purified 26S human proteasome. The decay constant for each ubiquitylated species, separated on a gel, was measured from a time (T) series. Signal from unmodified substrates (Ub0) was used as a control for loading and nonspecific dephosphorylation, which is the main reason for the decrease of Ub0 signals (Materials and methods section). Ub, ubiquitin; K, lysine residue; M-DiUb, methylated diubiquitin chain; M-TetraUb, methylated tetraubiquitin chain. (B to D) 160 nM geminin, securin, and cyclin B–NT (Xenopus) were ubiquitylated using indicated constructs of ubiquitin. Their rates of degradation by 3 nM of purified human 26S proteasome were measured and shown as a function of total ubiquitins per substrate molecule. Error bars represent the SD of three experimental replicates. The asterisk in (C) indicates that the rate for this species is 1.4. The lack of data for certain species is due either to these species not having been tested or to their signals being too weak to quantify. (E and F) Human cyclin B–NT mutants carrying lysines only at indicated positions were ubiquitylated with either methylated ubiquitin (M-Ub) or WT Ub and tested in a quantitative degradation assay. Original autoradiography for retrieving the rate information is compiled in fig. S8. In (E), the inset shows the location of the D-box (DB) on cyclin B–NT and relevant ubiquitylatable lysine residues identified by mass spectrometry.

We analyzed the rates of degradation of ubiquitylated securin, geminin, and cyclin B–NT (N-terminal fragment from cyclin B), each with a known number of ubiquitin chains of defined length. We incubated these substrates with the 26S proteasome at a nonlimiting concentration (25) (fig. S3). The measured degradation rates were similar to rates observed in cells (26, 27). The degradation rates were substrate-dependent, even for substrates with the same number of conjugated ubiquitins. For example, highly ubiquitylated securin was degraded fastest, followed by cyclin B and geminin (Fig. 1, B to D). Both methylated K48-diubiquitin chains and WT ubiquitins promoted efficient proteasomal degradation. Conjugation with K48-diubiquitin chains supported a higher rate of degradation than K48-tetraubiquitin chains when the same number of total ubiquitins were conjugated to a substrate molecule (Fig. 1, B to D). The slower degradation associated with tetraubiquitin chains was not due to methylation (fig. S4) nor to potential competition from free tetraubiquitin chains at the experimental concentration, as multidiubiquitylated securin was degraded at the same rate, even after free tetraubiquitin chains were added to the degradation reaction (fig. S4). Faster degradation was observed with diubiquitin chains, even when the two chains were closely apposed (fig. S5); this also held for K11-linked chains (fig. S6). In contrast to cyclin B, multi-monoubiquitylated securin and geminin, generated with either methylated ubiquitin or lysine-free ubiquitin (Ub0K) precluding chain formation, were degraded very slowly compared with constructs with the same ubiquitin stoichiometry but containing chains (Fig. 1, B and C, and fig. S7).

The distribution of ubiquitylated lysines on the substrate may set the degradation rate. Because our method to control ubiquitin configurations does not constrain which lysine residues receive the ubiquitin chains, we explicitly tested the contribution of the sites of ubiquitylation to the degradation rate. Multi-monoubiquitylated cyclin B–NT was degraded as efficiently as WT Ub–conjugated cyclin B–NT (8) (Fig. 1D), perhaps because of the specific configurations of lysine targets on cyclin B. We made cyclin B–NT mutants that contain only three or four lysine residues at their original locations, with other lysines mutated to arginines, and studied rates of degradation of the multi-monoubiquitylated products of those mutants. A clear pattern emerged: Given the same number of total conjugated ubiquitins, the degradation rate strongly depended on the position of the most N-terminal, ubiquitylated lysine residue; the closer to the N terminus, the higher the rate of degradation (Fig. 1E). This pattern was not explained by some mutants being inherently nondegradable, because all mutants were degraded similarly when conjugated to WT Ub at high ubiquitin multiplicity (Fig. 1F) (Ub = 7 to 8). Therefore, moving lysine residues away from the N terminus appeared to progressively convert cyclin B into a substrate whose degradation was more similar to that of securin and geminin. SM studies presented below provide mechanistic insights into these properties.

Single-molecule studies of the kinetics of proteasomal degradation

Proteasome-substrate interactions

To understand the molecular steps that distinguish different ubiquitin configurations, we monitored the interactions of single ubiquitylated substrate molecules with the proteasome. Purified 26S proteasomes from human 293 cells were immobilized onto slides with an antibody to the core 20S proteasome (Fig. 2A); the surface was passivated with polyethylene glycol (PEG) and albumin to reduce nonspecific binding. To correlate the SM behavior with the extent of ubiquitin conjugation, each ubiquitin molecule was chemically labeled with a DyLight 550 fluorophore at the N terminus. Fluorescent labeling of ubiquitin had no measurable effect on the kinetics of ubiquitylation and degradation in bulk reactions (fig. S9). After ubiquitylation, the total fluorescence intensity of a substrate molecule was measured by total internal reflection fluorescence (TIRF) microscopy, from which the number of conjugated ubiquitins was calculated. Accuracy and linearity of this method were assessed and confirmed by photobleaching experiments, a process that randomly inactivates single fluorophores (28). Background fluctuation was less than 0.2 ubiquitin level; more than 90% of substrate-binding events can be identified with less than 30% uncertainty in measuring the number of conjugated ubiquitins (fig. S10). This uncertainty was principally due to residual uneven illumination.

Fig. 2 Proteasome-substrate interaction kinetics by the SM method.

(A) Schematics showing the experimental design, where purified 26S proteasome was immobilized on passivated coverslip using anti-20S antibody. Ubiquitin was fluorescently labeled and conjugated to substrates in solution. Sample pictures capture ubiquitin signals on the surface with either 26S proteasome (+26S) or antibody only (no 26S). (B) Average dwell time on the proteasome for different substrates (or free Ub chain) with varying numbers of conjugated ubiquitins, measured by the SM method. The distributions of individual dwell times are shown in fig. S15. Error bars represent SD of the mean. The asterisk indicates that data for chains of 6 to 9 ubiquitins are not shown due to insufficient events. (C) Cooperative and stochastic mechanisms affect binding enthalpy and entropy, respectively. By each mechanism, the expected relationship between dwell time tb and the number of ubiquitins N is shown below. ΔG, change in Gibbs free energy; ΔH, change in enthalpy; T, temperature; ΔS, change in entropy. (D) Dwell time on the proteasome (right), or its logarithm (left), for securin and cyclin B versus the number of conjugated ubiquitins. Red lines show linear fitting. The ratio of P values by fitting the red segment (in linear scale) with either a linear model (pl) or an exponential model (pe) is shown for each plot. ∆gub is the binding free energy per ubiquitin on the proteasome, calculated from the slope of the binding curve.

Ubiquitylated substrates transiently interacted with the 26S proteasome. Only background levels of binding were observed, either if the 20S proteasome was substituted for the 26S proteasome or if the substrate was omitted (fig. S11). Interaction with the proteasome requires a hydrophobic patch spanning Leu8-Ile44-Val70 on ubiquitin and is compromised by substitutions for Ile44 (29). In the SM assay, this mutation abrogated the binding of fluorescent ubiquitin to the proteasome (figs. S11 and S12). These results indicate that the interaction occurred predominantly at ubiquitin receptors on the 26S proteasome; the kinetics depended on the amount of ubiquitylation of the substrate and was insensitive to excitation laser intensities, excluding low-level fluorophore photobleaching as a complicating factor (figs. S13 and S14). To minimize systematic errors, each experiment and its controls were performed on the same slide with the same batch of proteasome and substrates.

To further validate the SM assay, we compared the affinity of ubiquitylated substrates with the proteasome using SM methods to published results measured in bulk assays. We determined the average dwell time of ubiquitylated cyclin B, securin, cyclin B with a single lysine residue (K64–cyclin B), or free ubiquitin chains on the proteasome, as a function of the total number of conjugated ubiquitins (Fig. 2B). At the same total number of ubiquitins, substrate-anchored ubiquitin chains interacted more strongly with the proteasome than did free chains, consistent with published results (5).

Higher ubiquitin stoichiometry consistently led to longer dwell times, but the quantitative relationship between dwell time and the amount of ubiquitin unexpectedly differed from substrate to substrate. Thus, although the dwell time for free ubiquitin chains or K64–cyclin B carrying a single ubiquitin chain plateaued at four to five ubiquitins, the dwell time for multi-lysine substrates, such as cyclin B and securin, increased continually as more ubiquitins were added. In a conventional competition assay, the Ki of a tetraubiquitin chain is 170 nM (5) [KiKd = koff/kon (Ki, inhibition constant; Kd, dissociation constant; koff, off rate; kon, on rate)]. The Kd value measured by the SM method is 210 ± 60 nM, given an estimated kon = 8.5 × 105 M–1 s–1 (30). A Lys6→Ala6 (K6A) mutation on ubiquitin (UbK6A) weakens the interaction of ubiquitylated substrates with the proteasome (29, 31). Consistent with this, the dwell time of UbK6A-securin on the proteasome was shorter than that of WT Ub–securin in the SM assay (fig. S16). The interaction between substrate backbone and the proteasome appears to be minimal, because the dwell time for substrates conjugated with a single ubiquitin was very short (dwell time < 300 ms) (Fig. 2B).

The dwell time obtained from SM measurements includes both a period for the initial interaction with the proteasome and, for productive interactions leading to degradation, a period for translocation and degradation. To assess the contribution of each step to the observed dwell time, we blocked enzymatically active sites of the proteasome with inhibitors that act downstream of the initial binding events (fig. S17). Neither ubiquitin-aldehyde, which inhibits deubiquitylation by Uch37, nor epoxomicin, which inhibits substrate proteolysis, changed the dwell time. Using 1,10-phenanthroline to inhibit Rpn11 activity and proteasomal degradation also caused little change in the dwell time (figs. S18 and S19). Therefore, dwell time primarily reports on the initial binding event between the ubiquitylated substrate and the proteasome.

Cooperative and stochastic features of proteasome-substrate interactions

Single-molecule studies can provide insights into mechanisms of the initial proteasome-substrate interaction, an important step for deconstructing the specificity of degradation. Only two proteasomal subunits, Rpn10 and Rpn13, contain ubiquitin-binding domains: Rpn10 has two UIM domains, and Rpn13 has one Pru domain (4). Together, they could maximally engage three ubiquitins at once. If a substrate molecule simultaneously bound to the three domains, this would constitute a type of cooperative or avidity binding process (Fig. 2C). Binding could also be enhanced by a “stochastic” mechanism, in which larger numbers of ubiquitins enhance the affinity of binding by increasing local ubiquitin concentrations, without requiring simultaneous interactions with different receptor proteins. These two mechanisms can be distinguished by dwell-time analysis, because a cooperative mechanism primarily affects the enthalpic component of the free energy of binding, whereas a stochastic mechanism should change only the entropic contribution. Kinetically, the cooperative mechanism should result in an exponential increase of dwell time with the number of conjugated ubiquitins, but a stochastic mechanism would be characterized by a linear increase, reflecting mass action (Fig. 2C) (32).

For both cyclin B–NT and securin, the dwell time initially increased exponentially as a function of the number of conjugated ubiquitins, up to a total of three (Fig. 2D), consistent with a cooperative mechanism (Fig. 2C). By contrast, further ubiquitylation (from four to nine ubiquitins) led to a linear increase in dwell time, consistent with a stochastic mechanism (Fig. 2, C and D). From the slope of the binding curve on a semi-log plot, we calculated the mean free energy of binding per bound ubiquitin to the proteasome: 0.92 kcal/mol for securin and 0.97 kcal/mol for cyclin B, suggesting weak interactions. The cooperative behavior at low-ubiquitin stoichiometry and the linear behavior at higher levels provide keys to understanding how ubiquitin configurations are discriminated.

The role of the ubiquitin chain structure in proteasome-substrate interactions

Ubiquitin chains interact more strongly with the proteasome than do ubiquitin monomers (5). Most proteasomal substrates require conjugation of ubiquitin chains for degradation. As we showed, multi-monoubiquitylated securin (or geminin) was degraded much more slowly than securin with ubiquitin chains (Fig. 1, C and B). The simplest explanation would be that this difference is due to weaker interaction of the multi-monoubiquitylated forms with the proteasome. To test this explanation, we measured the interaction between Ub0K-conjugated securin and the proteasome and compared it to securin linked to WT ubiquitin. Surprisingly, Ub0K-conjugated securin and WT Ub–conjugated securin showed almost identical dwell times on the proteasome when we compared substrates with the same total number of ubiquitins (Fig. 3A). This result was not caused by surface immobilization of the proteasome because it also occurred in binding assays performed in bulk, measuring interactions with the ubiquitin receptor Rpn10 on the proteasome (fig. S20). Likewise, cyclin B, which can be degraded without forming ubiquitin chains, had a similar binding affinity whether conjugated with WT Ub or Ub0K (Fig. 3B).

Fig. 3 Interaction with the proteasome is mainly determined by the total number of ubiquitins on a substrate, insensitive to Ub chain structures.

(A and B) Securin or cyclin B–NT was ubiquitylated by APC with either WT Ub or Ub0K and was tested for interaction with the proteasome, as in Fig. 2B. (C and D) Interaction of WT Ub– or Ub0K-conjugated cyclin B with the proteasome in the presence of ATP or ATP-γ-S. rep1 and rep2 are two experimental replicates. The data for the ATP-proteasome experiment are plotted for comparison; these data are identical to those shown in (B). Cyclin B–ATP data in (C) are identical to those in (B). In (A) to (D), error bars denote SD of the mean. (E) Degradation rate and proteasomal dwell-time relationship for WT Ub– and Ub0K-conjugated securin. Degradation rates were measured in the quantitative degradation assay (Fig. 1C), and dwell time on the proteasome was measured using the SM method. The inset shows the Ub0K result on a smaller y axis. (F) Ratio of “commitment efficiency” for securin–WT Ub over securin-Ub0K, as a function of total conjugated ubiquitins. Commitment efficiency is defined as the degradation rate divided by the dwell time.

Although degradation of most substrates appears to be much more efficient if chains are formed, our results show that the presence of chains does not alter the affinity of substrate binding to the proteasome. Instead, affinity is primarily determined by the total number of ubiquitins on a substrate molecule, irrespective of their configurations except in the special case of single-chain substrates. Nevertheless, the increase in degradation rate when chains are present implies that ubiquitin configuration has an important role in the fate of the substrate after its binding to the proteasome. Therefore, we used SM methods to examine how the presence of ubiquitin chains affects various steps in the degradation process, with the goal of understanding how ubiquitin chains stimulate substrate degradation.

During the process of degradation, the proteasomal subunits in the 19S regulatory particle alternate among different adenosine triphosphatase (ATPase)–driven conformational states (3335). To test whether the recognition of ubiquitin chains may require a particular proteasome conformation, we locked the proteasome in the adenosine triphosphate (ATP)–bound state with the nonhydrolyzable analog adenosine 5′-O-(3-thiotriphosphate) (ATP-γ-S) (fig. S18) and studied the proteasome’s interaction with ubiquitylated substrates. There was a consistent increase of ~25% in dwell time for binding of WT Ub–conjugated cyclin B to the ATP-γ-S proteasome, as compared with proteasomes in ATP buffer. This increase in affinity was not observed when cyclin B was conjugated with Ub0K (Fig. 3, C and D). As a more sensitive metric for this effect, 29% of the binding events between ATP-γ-S proteasome and WT Ub–conjugated cyclin B lasted longer than 10 s, whereas only 10% of binding interactions had such persistence under the three control conditions (fig. S21). Another substrate, securin, gave similar results (fig. S22). Proteasomes in buffer containing adenosine diphosphate (ADP) showed weaker interactions with ubiquitylated cyclin B than those in ATP-containing buffer, and the binding of WT Ub–conjugated cyclin B to ADP proteasomes was not different from that of Ub0K-conjugated cyclin B (fig. S23). We thus conclude that the discrimination of different configurations of ubiquitin on a substrate is enhanced in the ATP-bound state of the proteasome and therefore may result from rearrangement of the dynamic conformations of the proteasomal subunits. This discrimination was observed as a small binding enhancement, which by itself is unlikely to explain the generally large degradation rate enhancement for Ub chain–containing substrates. Perhaps the rearrangement of proteasomal subunits may expose or activate a hidden ubiquitin chain receptor that contributes to substrate discrimination. Two alternative mechanisms for the enhancement of protein degradation in the presence of ubiquitin chains—allosteric opening of the gate of 20S complex and stimulation of proteasomal ATPase activity (3638)—were found in control studies to have little effect when tested with ubiquitylated Ube2S as the substrate (figs. S24 and S25). Therefore, these mechanisms are unlikely to explain the much larger effect of ubiquitin chains in promoting degradation (Fig. 1, B and C).

The role of ubiquitin chains in initiating substrate translocation

If binding strength was the only determinant for degradation, the degradation rate should be proportional to the dwell time on the proteasome, as predicted by a probabilistic model of chemical reactions. For Ub0K-conjugated securin, this appeared to be the case over most of the range of ubiquitylation (Fig. 3E), where the rate versus dwell time was linear. By contrast, the degradation rate of WT Ub–conjugated securin increased superlinearly with dwell time (Fig. 3E), suggesting that ubiquitin chains, in addition to providing interaction with the proteasome, may also increase the efficiency of commitment of a substrate to degradation once bound to the proteasome [i.e., a turnover number (kcat) effect] (Fig. 3F). Notably, at high modification levels, WT Ub was up to 40 times more efficient in promoting securin degradation than Ub0K, per unit dwell time.

In addition to binding kinetics, SM traces can provide critical mechanistic information about events that follow binding, such as substrate translocation and degradation by the proteasome, in the form of the kinetics of ubiquitin removal from the substrate. We observed two different types of signals after binding of substrate to the proteasome: (i) structureless, in which there was a complete loss of ubiquitin within 200 ms (the sampling rate of the camera), and (ii) stepped, in which the ubiquitin signal decreased in several discrete steps with very short intervals (Fig. 4A). These effects were not attributable to photobleaching or the limited speed of camera detection (figs. S26 and S27).

Fig. 4 Ubiquitin chains on substrates promote translocation initiation.

(A) Examples of structureless and stepped traces from a SM experiment on WT Ub–conjugated securin. Stepped traces are due to progressive deubiquitylation by Rpn11, as illustrated in (B). (C) The fraction of stepped traces, as a function of the number of ubiquitins, for WT Ub (N = 663) or Ub0K-conjugated (N = 133) securin. Error bars represent SD of the mean.

Although the likely explanation for structureless events is dissociation of substrate from the proteasome before ubiquitin removal, the stepped decrease of the ubiquitin signal could represent a series of deubiquitylation events catalyzed by Rpn11, the only effective DUB in our samples (Fig. 4B). In support of this interpretation, inhibitors of Rpn11 activity, such as 1,10-phenanthroline and ATP-γ-S, almost completely eliminated the stepped decreases (fig. S28). When cyclin B–NT was conjugated with multiple diubiquitin chains, we observed with each step the removal of two ubiquitins rather than one (fig. S29). Thus, the proposal that Rpn11 cleaves ubiquitin chains en bloc (23) is strongly supported at the SM level. Accordingly, we did not observe the stepped loss of ubiquitin signals for ubiquitylated K64–cyclin B, a single-lysine substrate that carries only one ubiquitin chain (fig. S27). Presumably, if all ubiquitins on a multi-lysine substrate happened to form a single chain, only structureless events would be observed. However, such cases should be rare for APC substrates, because the E2 UbcH10 forms long ubiquitin chains with low efficiency. E2 UbcH10 acts broadly to increase the number of sites of mono- or diubiquitylation on multiple lysines (3).

Kinetic studies also reveal the sequence of events in degradation. Before the very processive phase of deubiquitylation, there was always a delay. The distribution of delay time was not exponential, as would be expected for a single-step reaction. Instead, the distribution corresponded closely to a Γ distribution with a modal delay of 2 s (fig. S30), suggesting intermediate rate-limiting steps between initial binding and the onset of deubiquitylation.

Rpn11 sits at the substrate entry port, where it has a critical role in removing ubiquitin chains on translocating peptides (6, 3942). Translocation may signal irreversible engagement of the substrate into the degradation process. This tight coupling between deubiquitylation and translocation was supported by the absence of deubiquitylated but nondegraded substrates in our bulk assays (fig. S1). We therefore used the processive deubiquitylation mediated by Rpn11 as a kinetic indicator for substrate translocation. To test whether the substrate was cleaved by the proteasome contemporaneously with deubiquitylation, we labeled both ends of cyclin B–NT with DyLight 550. We observed a stepped decrease of fluorescence on cyclin B that was sensitive to proteasome inhibition by MG132 and was coincident with the deubiquitylation, suggesting cotranslocational activity of Rpn11 (figs. S31 and S3). We compared the fraction of stepped events among all binding events for WT Ub–conjugated securin and Ub0K-conjugated securin to assess their relative probability of undergoing deubiquitylation and translocation steps. At increased amounts of conjugated ubiquitin, this probability increased dramatically for securin conjugated with WT ubiquitin capable of forming chains. By contrast, it remained low if Ub0K, which precludes chain formation, was used (Fig. 4C). This difference was large enough to account for the 12-fold degradation rate disparity between WT Ub–conjugated and Ub0K-conjugated securin. Multi-monoubiquitylated securin was not intrinsically refractory to deubiquitylation, which still occurred processively in the few cases of stepped events on multi-monoubiquitylated securin (fig. S32). Thus, ubiquitin chains on substrates appear to specifically promote initiation of translocation and degradation.


The multiple lysines on a substrate and on ubiquitin itself generate a large number of possible ubiquitin configurations. These configurations represent a “ubiquitin code” of unknown degeneracy that must be read by the proteasome and converted into a rate of substrate degradation. We have approached this problem with two methods: construction of substrates with defined ubiquitin configurations and SM techniques. We found that tetraubiquitin chains are not essential for rapid proteasomal degradation of APC substrates, which would explain why a tetraubiquitin receptor on a proteasome has not been found. In fact, ubiquitin chains on cyclin B, and possibly other APC substrates, are typically short (3), and multiple ubiquitylatable lysine residues are a common feature of these substrates. A distributed array of short ubiquitin chains appears to be a superior and perhaps an optimal signal for proteasomal degradation; this conclusion could probably extend to substrates of other E3 ligases. Although a single ubiquitin chain may be sufficient for degrading certain substrates, such as Sic1 mutants and IκB (16, 43), increasing the number of ubiquitylated lysine residues of the canonical single-chain substrate β-galactosidase greatly accelerates its degradation (44). Similarly, WT cyclin B was degraded faster than mutants with fewer lysines at the same total amount of ubiquitylation (Fig. 1F) (Ub = 7 to 8). Besides K48 chains, the APC also establishes K11 and K63 linkages on substrates (3). We found that K48 chains promoted more efficient degradation than K11, K27, and K63 chains (fig. S6).

By studying cyclin B mutants, we found that proximity of the first ubiquitylated lysine to the N terminus was associated with faster degradation (Fig. 1E), suggesting that the degradation rate is sensitive to the position of ubiquitylated lysine residues. There was a correspondence between long dwell times and elevated rates of degradation (fig. S33A), suggesting that ubiquitin chain position could control the rate of degradation, at least partially, through controlling affinity with the proteasome. Single-lysine mutants of cyclin B with a ubiquitin chain at different positions had indistinguishable binding kinetics to the proteasome, suggesting that monochain and multichain substrates may interact with the proteasome by different mechanisms (fig. S33B). For WT substrates, our current method of constructing defined ubiquitin configurations does not specify the chain positions. The results are understood as a populational average of all actual combinations of positions, of which the vast majority can promote efficient degradation.

Comparison of the Kd values for tetraubiquitin chains measured by our SM methods with those from bulk assays suggests that surface immobilization of the proteasome is unlikely to distort kinetic rate constants. In addition, ubiquitylated securin took, on average, 10 s to complete translocation and possibly the degradation process on the surface-bound proteasome (fig. S34). This result is consistent with time for degrading similar-size proteins, such as dihydrofolate reductase and Sic1, measured in bulk assays (45), suggesting that the surface-bound proteasome is unimpaired for unbiased kinetic studies.

The rate of degradation is determined by both binding (i.e., Kd effect) and postbinding (i.e., kcat effect) events on the proteasome. For APC substrates, multiple diubiquitin chains were more efficient degradation signals than tetraubiquitin chains, given the same total number of conjugated ubiquitins (Fig. 1, B to D). The explanation for this distinction may be their different binding strength with the proteasome. Using SM methods, we observed weaker binding if ubiquitins were assembled into a single chain on K64–cyclin B when compared to that for WT cyclin B, which had short and distributive chain configurations (Fig. 2B). High-resolution cryo-EM structures of the proteasome are consistent with the potential effectiveness of multiple short ubiquitin chains. Because proteasomal ubiquitin receptors Rpn10 and Rpn13 are distant from each other (6, 7), distributive configurations of ubiquitins on a substrate molecule might promote the use of more ubiquitin receptors on the proteasome; a single ubiquitin chain might also be less effective due to steric constraint. Furthermore, Rpn10 and Rpn13 may not be the only ubiquitin receptors on proteasome because budding yeast can tolerate mutations of both (46). Additional receptors or shuttle factors for the proteasome are also thought to contribute to the binding of ubiquitylated proteins (47, 48) (see below).

The SM binding measurements suggest a model wherein a substrate molecule samples multiple modes of binding during its interaction with the proteasome (Fig. 5). Evidence for such a mechanism comes from measurement of the dwell time as a function of ubiquitylation levels on the substrate. Beyond the ubiquitin-binding capacity of the proteasome, most likely limited to three or four ubiquitins by available ubiquitin receptors, a further increase of binding affinity relies on an increase in the local ubiquitin concentration on the substrate: the stochastic interaction. This stochastic mechanism stabilizes the bound state by increasing its entropy, or the number of microscopic states, because entropy is proportional to the logarithm of the number of these states. In this system, an increase in relevant entropy may occur if the substrate molecule can explore multiple conformations on the proteasome through intramolecular diffusion while remaining associated with the proteasome (Fig. 5). Such dynamic sampling should also increase the likelihood that the peptide terminus would be captured by the substrate entry port on the ATPases, thereby facilitating initiation of translocation. A cooperative process is implied by the exponential increase of dwell time as a function of the number of conjugated ubiquitins, whereas a stochastic process is implied by a linear increase (Fig. 2, C and D). A similar, biphasic binding relationship (1/Ki ~ Ub number) was suggested in an early publication using competition assays, though the interpretation was different (5) (fig. S35). An exponential increase of dwell time involving greater than three simultaneous interactions would further increase the discrimination of ubiquitin levels over a linear increase. Why, then, is the process no longer exponential after four ubiquitins? Cooperative mechanisms tend to promote tight binding, which has potential risks for the cell. If a highly ubiquitylated substrate could not be degraded by the proteasome, it would stably block the proteasome. Such an inhibition by stably binding complexes has been proposed to underlie the accumulation of ubiquitylated intermediates in various neurodegenerative diseases (49). A linear increase in affinity at high-ubiquitin stoichiometry, though less discriminating, is also less prone to form unproductive, inhibitory substrate-proteasome complexes.

Fig. 5 An integrated model for the degradation of ubiquitylated substrates by the 26S proteasome.

Polyubiquitylated substrates could simultaneously interact with multiple ubiquitin receptors on the proteasome. While remaining bound, a substrate molecule explores multiple configurations on the proteasome through intramolecular diffusion (1). Rearrangements of proteasomal subunits, fueled by the ATPases in response to binding and hydrolysis of ATP, expose or activate a deeper ubiquitin chain receptor and facilitate the transfer of ubiquitylated substrates from initial binding to a deeper engagement (2 and 3). In this engagement, the substrate or its terminus is closer to the substrate entry port, which expedites translocation initiation (3) and ensuing degradation (4).

Although binding to the proteasome is a prerequisite for degradation, it does not in itself determine the rate of degradation. For example, WT Ub–conjugated securin and Ub0K-conjugated securin bind equally tightly, but the former is degraded much faster than the latter. The specificity of degradation must also reflect postbinding events. The SM analysis of Rpn11-dependent deubiquitylation indicates that the chain structure of ubiquitin promotes the initiation of translocation, a requirement for degradation. This effect of Ub chains also applies to cyclin B, a special substrate that can be degraded even without ubiquitin chains. We observed a shorter delay between binding and the initial deubiquitylation event for WT Ub–conjugated cyclin B than for Ub0K conjugates, consistent with the translocation-promoting activity of ubiquitin chains (fig. S36).

Most substrate-proteasome encounters do not lead to degradation, especially for substrates with a low number of conjugated ubiquitins (Fig. 4C). Even for highly ubiquitylated substrates, binding events sometimes lasted for tens of seconds without leading to degradation. Thus, there may be a latent state of the proteasome, in which heterogeneous ubiquitin chain conformations might affect deubiquitylation (50) or, perhaps more likely, might affect the orientation of a bound substrate, placing the translocation-initiating element far away from the substrate entry port. In this context, the presence of a short flexible domain at a substrate’s terminus should substantially accelerate its rate of degradation (20, 51). Therefore, engagement of the translocation-initiating element by force-generating pore loops of the proteasomal ATPases, which are reached via the substrate entry port, may generally be a rate-limiting step in degradation. Translocation initiation has been proposed to underlie “commitment,” a hypothetical point at which substrates are irreversibly destined to degradation (39). We argue that translocation initiation, sensitive to the configuration of ubiquitin groups on the substrate, is either the commitment step or is closely coupled to it.

To understand how ubiquitin chains promote translocation initiation, we propose a model based on our experimental observations (Fig. 5). Conformational changes of the proteasome, driven by the ATPases quickly transiting through different nucleotide-bound states (33, 40), may activate a ubiquitin chain receptor(s) that participates in substrate recognition (52). Candidates for such a receptor include ATPase Rpt5, which can be cross-linked to bound ubiquitin chains (53). The same result can also be explained by rearrangement of ubiquitin receptors (Rpn10 and Rpn13) into a higher-affinity state for ubiquitin chains. It would make sense if the additional ubiquitin chain receptor were closer to the substrate entry port than Rpn10 or Rpn13 to facilitate translocation initiation by engaging the substrate into a deeper conformation after the initial interaction involving mainly Rpn10 and Rpn13 (Fig. 5). Such an intermediate step is indicated by the delay before deubiquitylation or translocation (fig. S30). Consistently, Rpt5 is very close to the substrate entry port, and conformational changes induced by ATP-γ-S dramatically reduce the distance between Rpn10 and Rpt4/5, indicating a possible direct transfer of substrates from initial binding to deeper engagement (33, 54) (fig. S37).

We have shown that there is no simple length threshold for ubiquitin chains for degradation by the proteasome. Rather, there are at least two requirements: a minimal number of ubiquitins to result in tight binding and a certain number or length of chains to promote translocation into the axial channel. The ultimate rate of degradation is probably set by ubiquitin stoichiometry, chain configuration, and properties of the substrate that affect not only the capacity to be ubiquitylated and the configuration of chains but also the orientation of the chains and translocation-initiating elements once bound to the proteasome.

Materials and methods

Protein purification and labeling

Recombinant, full-length human securin, Ube2S, GST-Emi1 (297-447), Xenopus geminin, Xenopus N-terminal cyclin B (amino acids 1 to 104), human N-terminal cyclin B (amino acids 1 to 88), and mutants were purified from Escherichia coli cells using an IMPACT kit (NEB, E6901S). A PKA (protein kinase A) site (RRASV) was also placed at the N terminus of the substrates used in degradation assays. Human ubiquitin, UbK48, Ub0K, UBK6A, and UbI44A mutants, each with a cysteine residue and a His tag at the N terminus, were purified from E. coli cells using cation-exchange chromatography (GE, 17-1152-01) labeled with DyLight 550 maleimide (Pierce, 62290). After removing unreacted dyes, labeled ubiquitin was subjected to anion-exchange chromatography (GE, 17-1153-01) to separate labeled and unlabeled species. Finally, the N-terminal His tag was cleaved off using thrombin.

Fluorescently labeled ubiquitin chains were synthesized using E2-25K in reactions containing 30 μM DyLight 550–labeled UbK48 and 10 μM DyLight 550–labeled Ub0K; carried out in UBAB buffer [25 mM Tris-HCl (pH 7.5), 50 mM NaCl, 10 mM MgCl2], 3 mM ATP, 1 μM E1, and 10 μM E2-25K; and incubated at 37°C for 16 hours. The product was purified and separated using anion-exchange chromatography. For binding assays with the proteasome, different fractions of ubiquitin chains were mixed as the sample.

Anti-20S antibody (MCP21) was biotinylated using biotin-NHS (Pierce, 20217) and was purified using desalting columns.

Radioactive 33P-ATP was used to label substrates with a PKA site at the N terminus for in vitro ubiquitylation and degradation assays.

E1, E2s (UbcH10, E2-25K), WT ubiquitin, biotin ubiquitin, methylated ubiquitin, K48-diubiquitin chains, and K48-tetraubiquitin chains were from Boston Biochem. Purified streptavidin was from Invitrogen.

For biotinylating substrates, a biotin-containing peptide (NEB, Bio-P1) was ligated to the substrate C terminus, which had been activated by intein cleavage during purification, resulting in ~90% ligation efficiency.

Ubiquitin chain methylation

Ubiquitin chain samples were buffer-exchanged into a mixture of 50 mM Hepes-Na (pH 7.5), 150 mM NaCl, and 2% glycerol. Dimethylamine borane and formaldehyde were added to the sample to final concentrations of 20 and 40 mM, respectively. After 2-hour incubation on ice, another 20 mM dimethylamine borane and 40 mM formaldehyde were again added, bringing final concentrations to 40 and 80 mM, respectively, and incubation was continued for 16 hours on ice. Methylated proteins were buffer-exchanged to tris-buffered saline [20 mM Tris-HCL (pH 7.5), 150 mM NaCl].

APC purification from HeLa cell G1 extract

HeLa cell G1 extract preparation and APC purification were performed as described previously (13). Briefly, 2L HeLa S3 cell spinner culture was synchronized at prometaphase using thymidine/nocodazole double block and was then released for 3 hours into G1. Cells were harvested and subjected to nitrogen cavitation in 75% volume swelling buffer [20 mM Tris-HCl (pH 7.5), 5 mM KCl, 1.5 mM MgCl2, 1 mM dithiothreitol (DTT), protease-inhibitor tablet (Roche, 05892953001)]. APC was purified from cell extracts using anti-Cdc27 (AF3.1, custom-made) agarose and eluted using a competitive peptide.

Affinity purification of the 26S human proteasome

Human proteasomes were affinity-purified on a large scale from a stable human embryonic kidney 293 cell line harboring HTBH-tagged hRPN11 (a gift from L. Huang). The cells were Dounce-homogenized in lysis buffer [50 mM NaH2PO4 (pH 7.5), 100 mM NaCl, 10% glycerol, 5 mM MgCl2, 0.5% NP-40, 5 mM ATP, and 1 mM DTT] containing protease inhibitors. Lysates were cleared and then incubated with NeutrAvidin agarose resin (Thermo Scientific) overnight at 4°C. The beads were then washed with excess lysis buffer, followed by the wash buffer [50 mM Tris-HCl (pH 7.5), 1 mM MgCl2, and 1 mM ATP]. Usp14 was removed from proteasomes using the wash buffer plus 100 mM NaCl for 30 min. 26S proteasomes were eluted from the beads by cleavage, using TEV protease (Invitrogen).

In vitro ubiquitylation reaction

The APC ubiquitylation reactions were carried out in UBAB buffer [25 mM Tris-HCl (pH 7.5), 50 mM NaCl, 10 mM MgCl2] containing 5 nM APC, 100 nM E1, 2 μM UbcH10, 2 mg/ml bovine serum albumin (BSA), an energy regenerating system, and either ubiquitin or ubiquitin chains: 10 μM ubiquitin, methylated ubiquitin, or DyLight 550–Ub0K; 5 μM methylated K48-diubiquitin; and 1 μM methylated K48-tetraubiquitin. Reactions were incubated at 30°C. Results were quantified using a phosphorimager. For PKA-labeled substrates, calyculin A (EMD, 19-139), a broad-spectrum phosphatase inhibitor, was added at 10 μg/ml.

Ube2S autoubiquitylation reactions were carried out in UBAB buffer containing 6 μM Ube2S, 0.5 μM E1, and 5 mM ATP. 100 μM ubiquitin, methylated ubiquitin, or 40 μM methylated K48-diubiquitin chain were also added. Reactions were incubated at 37°C for 4 hours.

Quantitative degradation assay

PKA-labeled substrates (800 nM) were ubiquitylated by the APC using various forms of ubiquitin or ubiquitin chains for 45 min at 30°C. Reaction products were then mixed with proteasome-containing solution (UBAB buffer, 3 nM 26S proteasome, 2 mg/ml BSA, 1 μM GST-Emi1, 3 mM ATP, 10 μg/ml calyculin A) at a ratio of 1:4. The degradation reaction was sampled at 0, 3, 6, and 15 min at 37°C and was quantified using a phosphorimager.

To calculate the degradation rate, the intensity of each ubiquitylated species was quantified using ImageJ. These values were normalized to the intensity of the unmodified substrate (Ub0) to correct for loading error and nonspecific dephosphorylation. Finally, traces of the time course of degradation were fitted with an exponential decay function to obtain the rate constant.

Peptidase assay

Products of Ube2S autoubiquitylation reaction (6 μM) were mixed with proteasome-containing solution (UBAB buffer, 6 nM 26S proteasome, 2 mg/ml BSA, 3 mM ATP, 30 μM Z-GGL-AMC, 0.4 mM DTT) at a 1:1 ratio. Hydrolysis of GGL-AMC was monitored at 37°C using a fluorescence spectrometer.

Binding assay for ubiquitylated substrate

Wild-type securin was ubiquitylated in a standard APC reaction with either WT Ub or Ub0K in a total volume of 20 μl for 1.5 hours at 30°C. The product was incubated with GST-magnetic beads conjugated with GST-Rpn10 for 40 min at 4°C (8). After 2× wash with buffer [25 mM Tris-HCl (pH 7.5), 100 mM NaCl, 0.5 mM EDTA, 0.05% Tween-20], the supernatant and bead-bound fraction were assayed for securin using anti-securin antibody by Western blot.

Slide passivation

We followed a basic slide passivation protocol using 5-kD PEG plus 2.5% 5-kD biotin-PEG (LaysanBio, MPEG-SVA-5000; Biotin-PEG-SVA-5000) in a “clouding-point” solution on amino-silanized slides for 4 hours, as described previously (55, 56). In addition, we passivated them again using 1-kD PEG-NHS (Nacocs, PG1-SC-1k) for 1 hour, followed by 10% (w/v) BSA for 30 min before use. After passivation, slides were assembled into reaction chambers (55). Streptavidin (0.2 mg/ml) was incubated with passivated slides for 5 min for immobilizing biotinylated substrates. The quality of slide passivation was tested by incubating with HeLa cell extracts supplemented with DyLight 550–Ub and measuring the level of nonspecific binding.

Single-molecule proteasome assay

Purified substrates (200 nM) were ubiquitylated by the APC with 5 μM fluorescent ubiquitin (WT Ub, Ub0K, UbK6A, or UbI44A) for 40 min (60 min for K64–cyclin B) at 30°C. The reaction product was diluted 100× into imaging buffer (UBAB, 3 mM ATP, 40 mM imidazole, 5 mg/ml BSA). Imidazole in the imaging buffer serves to reduce the nonspecific binding of fluorescently labeled proteins.

26S proteasome (15 nM) and biotinylated MCP21 antibody (15 nM) were mixed and incubated at room temperature for 15 min. The proteasome-antibody mix was loaded onto passivated slides coated with streptavidin. After a brief incubation, unbound protein was washed off and replaced with imaging buffer containing diluted ubiquitylation product. Image acquisition was started immediately with <15 s delay.

For experiments involving ATP-γ-S, ATP in imaging buffer was replaced with 2.5 mM ADP plus 0.5 mM ATP-γ-S. For experiments involving ADP, proteasome was incubated in 3 mM ADP buffer. ADP buffer and ubiquitylation product were also incubated with 30 mM glucose and 2 mg/ml hexokinase for 60 min at 30°C to remove residual ATP in the solution. For experiments involving 1,10-phenanthroline, 3 mM phenanthroline was added to the imaging buffer and proteasomal mixture; the sample was incubated for 20 min at room temperature before performing the experiment.

For the SM degradation assay, human cyclin B–NT was engineered with two cysteine residues close to its N and C termini and was labeled with DyLight 550–maleimide. Doubly labeled cyclin B was enriched via anion-exchange chromatography (HiTrap Q HP, pH 8.0). After ubiquitylation with unlabeled WT ubiquitin, 2 nM cyclin B was studied in the SM setup, as described above. Data analysis was identical to that described for the study of deubiquitylation.

We used a Nikon Ti TIRF microscope equipped with three laser lines of 491 nM (27 mW, full output from objective), 561 nM (30 mW), and 640 nm (59 mW), as well as an Andor DU-897 EMCCD camera. Time series were acquired at 200 ms per frame for 2 min, unless indicated.

Single-fluorophore intensity calibration

The fluorescence intensity of a single DyLight 550 molecule on ubiquitin was calibrated through photobleaching preformed ubiquitin chains (27). Ubiquitin chains were synthesized in a standard E2-25K reaction (see above) containing 30 μM DyLight 550–labeled ubiquitin and 5 μM biotin-ubiquitin in UBAB buffer supplemented with 3 mM ATP, 1 μM E1, and 5 μM E2-25K. The reaction was incubated at 37°C for 16 hours.

The E2-25K reaction product was diluted 10,000× in imaging buffer (above) and loaded on a passivated slide via streptavidin. Photobleaching was observed with the use of a TIRF microscope, illuminated with a laser-intensity level 2.5× higher than that in proteasome experiments (to achieve faster photobleaching).

The uncertainty of measuring ubiquitin stoichiometry is ~30%, as suggested by the standard deviation of the intensity distribution of single fluorescent ubiquitin. This uncertainty is primarily due to uncorrected uneven illumination.

Single-molecule data reproducibility

To test the reproducibility of dwell time data as in fig. S13, the same experiment with slightly varying concentrations (±50%) of ubiquitylated substrates was repeated three times on the same coverslip but at different positions.

Long-term (<1 year) reproducibility tests of the results of binding measurements carry 10 to 20% variation, probably due to batch-to-batch variability of proteasome samples or laser intensity drifts. Therefore, to minimize systematic variation in the measurements, each experiment with its controls was performed on the same slide using the same batch of proteasome and substrates.

Data analysis

The workflow of data analysis is illustrated in fig. S38.

Image processing

Raw images were first corrected for stage drifting after subtracting a uniform-intensity background. Stage drifting was corrected by subtracting the stage motion, which was derived using successive image registration to calculate the shift (step 1 in fig. S38). A custom-built algorithm was applied to identify binding spots of ubiquitylated molecules to the proteasome on the basis of their absolute intensity and local signal-to-noise ratio (step 2 in fig. S38). The spot-identification algorithm is based on finding the local maxima of ubiquitin fluorescence intensity in a field of view by applying a filter requiring the local signal-to-noise ratio to be larger than 6 and no other spots within four pixels around it. In this way, >90% of substrate-binding events can be identified compared with a blind manual identification (fig. S10). The false-positive rate is less than 5%, as shown by the nonsubstrate control, considering that one ubiquitin generates an average of ~20 photons on the camera per frame. After each substrate-binding spot has been recognized in a time lapse, spots are registered along the time axis according to their coordinate to designate substrate-binding events (step 3 in fig. S38). Specifically, if the coordinates of two substrate spots in two consecutive frames are less than or equal to one pixel apart, they are considered to be the same binding event. The absolute intensity of each spot was obtained by fitting the SM diffraction pattern with a two-dimensional Gaussian function, which generates both the signal intensity I(s) and the local background level I(bg) (step 4 in fig. S38). The signal intensity I(s) is then corrected for inhomogeneous illumination (step 5 in fig. S38). Inhomogeneous illumination was corrected by a self-adaptive algorithm that separates the entire field of view into 15-by-15 identical squares and uses the average fluorescent spot intensity in each square as a surrogate for the relative illumination intensity in the center of that square. Because the illumination intensity varies slowly across the field of view, we interpolated the illumination intensity at a given position from the center values in each square. The corrected signal intensity was then converted to absolute ubiquitin numbers by normalizing with the intensity value of a single ubiquitin obtained in the calibration step. The resulting traces were directly used for subsequent analysis or represented in figures, with no further processing.

Dwell-time analysis

A custom-built algorithm was used to measure the duration of substrate-binding events (dwell time), based on the background noise intensity plus a cutoff of 0.7 Ub level. The binding measurement is insensitive to the choice of the cutoff (0.5 to 0.9 Ub) or laser intensity (fig. S14). The maximal fluorescence intensity (converted to the number of ubiquitins using a calibrated single-fluorophore intensity value) during each binding event was measured and plotted versus the dwell time.

Stepped-event identification

A custom-built step-detection algorithm was used to identify stepped events (fig. S39). The formal definition of a stepped event is as follows: If two consecutive drops of fluorescent signal (>0.7 Ub) occur within 5 s, this is designated as a stepped event. Also, it is required that each intermediate state must last longer than one frame. Traces (~80%) can be identified correctly as compared with manual identification. Obvious classification errors were later corrected manually.

For counting the fraction of stepped events, only those events whose ubiquitin number was ≥3 were included in the analysis. We also excluded binding events that lasted longer than 90 s (5 to 10% of total binding events).

The same step-detection algorithm was also used in the analysis of photobleaching traces, to extract single-fluorophore intensity values.

Curve fitting in Fig. 2D

Segments of the curve (indicated in the figure) showing dwell time versus number of ubiquitin relationships were fitted with a linear function or an exponential function. A χ2 test Embedded Image (yi, data point i; Embedded Image, data point on the fitted line; ei, standard error) was performed to obtain the P value for each fitting model.

Photobleaching experiment on ubiquitylated substrates

Biotinylated securin was ubiquitylated by APC using DyLight 550–Ub and was immobilized on passivated coverslips via streptavidin. Time-lapse images were acquired under conditions identical to the SM proteasomal assay, but in the absence of proteasome. The total signal intensity in a central region of the field was analyzed to extract photobleaching information (fig. S26).

Supplementary Materials

Supplementary Text

Figs. S1 to S39


References and Notes

  1. See supplementary text (“Estimating the on-rate of tetraubiquitin chains with immobilized 26S proteasome in SM experiments”).
  2. See supplementary text (“Background information on the stochastic binding mechanism”).
  3. Acknowledgments: We thank L. Bai and W. Ma for commenting on the manuscript. We thank the Nikon Imaging Center at Harvard Medical School for providing instruments. Y.L. is supported by a Damon Runyon Research Fellowship and is a Lallage Feazel Wall Fellow. This work was supported in part NIH grants GM43601 (to D.F.) and GM66492 (to R.W.K.)
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