BIM-extra long (BIMEL), a pro-apoptotic BH3-only protein and part of the BCL-2 family, is degraded by the proteasome following activation of the ERK1/2 signalling pathway. Although studies have demonstrated poly-ubiquitylation of BIMEL in cells, the nature of the ubiquitin chain linkage has not been defined. Using ubiquitin-binding domains (UBDs) specific for defined ubiquitin chain linkages, we show that BIMEL undergoes K48-linked poly-ubiquitylation at either of two lysine residues. Surprisingly, BIMELΔKK, which lacks both lysine residues, was not poly-ubiquitylated but still underwent ERK1/2-driven, proteasome-dependent turnover. BIM has been proposed to be an intrinsically disordered protein (IDP) and some IDPs can be degraded by uncapped 20S proteasomes in the absence of poly-ubiquitylation. We show that BIMEL is degraded by isolated 20S proteasomes but that this is prevented when BIMEL is bound to its pro-survival target protein MCL-1. Furthermore, knockdown of the proteasome cap component Rpn2 does not prevent BIMEL turnover in cells, and inhibition of the E3 ubiquitin ligase β-TrCP, which catalyses poly-Ub of BIMEL, causes Cdc25A accumulation but does not inhibit BIMEL turnover. These results provide new insights into the regulation of BIMEL by defining a novel ubiquitin-independent pathway for the proteasome-dependent destruction of this highly toxic protein.
BIM (BCL2-interacting mediator of cell death), a pro-apoptotic, BH3-only protein belonging to the BCL-2 protein family plays an important role in promoting cell death in response to several stimuli (Bouillet et al., 1999). In particular, BIM-encoding mRNA and BIM protein levels increase upon growth factor withdrawal, and BIM promotes cell death under these conditions (Bouillet et al., 1999; Whitfield et al., 2001; Ewings et al., 2007). Among the common BIM splice variants (BIM-short, BIM-long and BIM-extra long) BIM-extra long (BIMEL) is the most abundant and exhibits the most dynamic increases in expression following cytokine withdrawal (Whitfield et al., 2001; Weston et al., 2003; Ewings et al., 2007). Activation of the extracellular-signal-regulated kinases 1 and 2 (ERK1/2) pathway promotes the multisite phosphorylation of BIMEL, resulting in its proteasome-dependent degradation (Ley et al., 2003; Luciano et al., 2003; Marani et al., 2004). BIM is a tumour suppressor gene (Egle et al., 2004) and stabilization of BIMEL resulting from ERK1/2 inhibition is important in tumour cell death (Wickenden et al., 2008; Gillings et al., 2009). Consequently, understanding the mechanism of BIMEL degradation is of fundamental interest and will inform the use of new oncogene-targeted therapeutics.
The identity of the E3 ubiquitin (Ub) ligase responsible for polyubiquitylation (poly-Ub) of BIMEL has been controversial (Akiyama et al., 2003; El Chami et al., 2005; Wiggins et al., 2007; Zhang et al., 2008). However, a recent study has demonstrated that ERK1/2 and ribosomal S6 kinase (RSK) cooperate to phosphorylate BIMEL, allowing binding of the F-box protein β-TrCP, which promotes BIMEL poly-Ub (Dehan et al., 2009). Despite this, the nature of the Ub chain linkage that modifies BIMEL in cells has never been defined. Here, we have used Ub-binding domains (UBDs) with defined selectivity for lysine 48 (K48)- or K63-linked Ub chains to show that BIMEL is subject to K48-linked poly-Ub in cells following ERK1/2 activation. However, a BIMEL mutant lacking lysine residues failed to undergo poly-Ub but was still degraded in a proteasome-dependent fashion. BIMEL has been proposed to be an intrinsically disordered protein (IDP) (Hinds et al., 2007) and many IDPs can be degraded by uncapped 20S proteasomes without prior poly-Ub (Tsvetkov et al., 2008; Baugh et al., 2009). We now show that BIMEL is degraded by 20S proteasomes in the absence of poly-Ub, but is protected from this by binding to the pro-survival BCL-2 protein, MCL-1. These results provide new insights into the regulation of BIMEL by defining a novel, Ub-independent pathway for its destruction; consistent with this, we find that inhibition of cullin-based E3 ligases such as β-TrCP has no effect on BIMEL abundance or turnover. Thus, cells employ multiple mechanisms to ensure the rapid destruction of this highly toxic protein.
Use of immobilized UBDs to demonstrate that BIMEL is modified by K48-linked poly-ubiquitin chains
Although K48-linked poly-Ub is recognized as a signal for degradation by the 26S proteasome, the nature of the Ub chain linkage on BIMEL has not been previously defined. Up to eight different types of Ub chain linkage might exist and this complexity is interpreted within cells by various UBDs, some of which have exquisite specificity for individual types of Ub chain linkage (Komander et al., 2009; Komander, 2009). To define the Ub chain linkage on BIMEL we used the immobilized Ub-associated (UBA) domains of GST–Dsk2 and GST–Mud1, which specifically bind K48-linked Ub chains (Ohno et al., 2005; Trempe et al., 2005; Lowe et al., 2006; Komander et al., 2009), and the NZF domain of GST–Tab2c, which is specific for K63-linked chains (Kulathu et al., 2009).
GST–UBDs on GSH-agarose beads were used in a pulldown assay to capture and partially purify poly-Ub BIM constructs from cell extracts. We first confirmed the specificity of GST–Dsk2 and GST–Mud1 for K48-linked chains using antibodies specific for K48- or K63-linked poly-ubiquitin chains (Newton et al., 2008). HEK293 cells were transfected with empty vector or BIMEL, and cell extracts were incubated with GST–Dsk2 UBA immobilized on GSH-agarose beads. These samples were readily detected by the K48-specific poly-ubiquitin antibody, which revealed a characteristic smear of poly-Ub species running up the gel, whereas the K63-specific poly-ubiquitin antibody failed to detect any material in the GST–Dsk2 pulldowns except the weak ‘ghost’ of the GST–Dsk2 protein itself, which is present in excess (Fig. 1A). The overexpression of BIMEL made no difference to the pattern of K48-linked poly-Ub species, indicating that it represents a tiny fraction of the total K48-linked poly-ubiquitin in cells. When these same samples were blotted with antibodies to BIM, we could readily detect full-length BIMEL and its poly-Ub species running as a smear up the gel in the sample from BIMEL-transfected cells. We could also detect endogenous BIMEL binding to GST–Dsk2 UBA, although the much lower levels meant that poly-Ub species of the endogenous BIMEL were difficult to detect (Fig. 1A). Identical results were obtained when we used GST–Mud1 UBA to precipitate BIMEL from control and transfected HEK293 cells (supplementary material Fig. S1A).
To define the specificity of the interaction between BIMEL and GST–Dsk2 UBA, we performed several further experiments. First, it is known that activation of the ERK1/2 pathway promotes the phosphorylation and enhances the turnover of BIMEL (Ley et al., 2003; Ewings et al., 2007). Consistent with this, when expressed in HR1 cells (HEK293 cells stably expressing the conditional kinase ΔRAF-1:ER*) (Ewings et al., 2007), the basal level of poly-Ub BIMEL, detected by binding to GST–Dsk2, was greatly enhanced when BIMEL phosphorylation was promoted by 4-hydroxytamoxifen (4-HT)-dependent activation of the ΔRAF-1:ER–MEK1/2–ERK1/2 pathway (Fig. 1B). Second, we compared BIMEL binding to GST–Dsk2 with that of BIML and BIMS (Fig. 1C), which lack the ERK1/2 and RSK phosphorylation sites that are thought to be the primary signal for poly-Ub and turnover. We again observed good binding of BIMEL, and poly-Ub species were readily apparent. However, binding was very weak with BIML and particularly BIMS and no poly-Ub species were detected (Fig. 1D), consistent with reports that BIML and BIMS stability is not regulated by ERK1/2 (Luciano et al., 2003; Ley et al., 2004; Wickenden et al., 2008). We also observed that the binding of BIMEL was drastically reduced when an inactive mutant form of GST–Dsk2 UBA was used in these pulldown experiments (Fig. 1D), and similar results were obtained with an inactive, mutant form of the GST–Mud1 UBA domain (supplementary material Fig. S1B). Finally, the GST–Tab2c NZF domain, which binds K63-linked poly-ubiquitin chains but not K48-linked chains (Kulathu et al., 2009), failed to precipitate poly-Ub BIMEL whereas GST–Dsk2 was again effective (supplementary material Fig. S1C). Thus, using the K48 linkage specificity of the Dsk2 and Mud1 UBA domains we demonstrate for the first time that BIMEL is subject to K48-linked poly-Ub in cells and this is enhanced following activation of the ERK1/2 pathway.
BIMELΔKK undergoes normal ERK1/2-driven proteasome-dependent turnover
Covalent attachment of Ub typically takes place at lysine residues. BIMEL contains only two lysine residues at K3 and K108 (numbered according to the rat sequence) (Fig. 1C) so we mutated either K3 (BIMELΔK) or K3 and K108 (BIMELΔKK). These mutants were transfected into HR1 cells that were then treated with 4-HT+MG132 (Fig. 2A). Wild-type BIMEL again exhibited a basal level of poly-Ub that was enhanced by 4-HT treatment. Mutation of K3 reduced the degree of basal and 4-HT-driven poly-Ub and caused the loss of certain poly-Ub species, whereas we failed to detect poly-Ub of BIMELΔKK despite overexposure of the blots (Fig. 2A). Thus, poly-Ub can take place at both lysine residues in BIMEL and mutation of both is required to generate a non-poly-Ub form.
Because BIMELΔKK was not poly-Ub in cells, we anticipated that it would accumulate at higher levels and so elicit greater cell death than wild-type BIMEL. However, we found that BIMEL and BIMELΔKK were equally effective at killing when transiently expressed in HEK293 cells (Fig. 2B). Furthermore, western blots revealed that wild-type BIMEL and BIMELΔKK were expressed at similar levels in transfected HR1 cells (Fig. 2A). These results prompted us to evaluate the turnover of the BIMELΔKK protein directly. When HR1 cells were transfected in parallel with haemagglutinin (HA)–BIMEL (Fig. 2C) or HA–BIMELΔKK (Fig. 2D) the two proteins again expressed at similar levels and exhibited very similar turnover in response to activation of the ERK1/2 pathway by 4-HT (Fig. 2C,D). As a further control, we observed that the ΔRAF-1:ER*-driven turnover of both HA–BIMEL and HA–BIMELΔKK was inhibited when ERK1/2 activation was prevented by the MEK1/2 inhibitor U0126 (supplementary material Fig. S2A,B). Thus, both wild-type BIMEL and BIMELΔKK exhibited very similar ΔRAF-1:ER*-driven, MEK1/2-dependent turnover, despite BIMELΔKK being defective for poly-Ub.
Proteasome-dependent degradation of BIMEL in the absence of poly-Ub
In considering other pathways that might contribute to the rapid turnover of BIMEL in the absence of poly-Ub we examined autophagy, a catabolic process in which the cell's own components are degraded by recruitment to autophagolysosomes. We compared immortalized mouse embryo fibroblasts (iMEFs) from wild-type and Atg5−/− mice (Kuma et al., 2004), which are defective for autophagy as judged by LC3 processing (Fig. 3A). The basal level of BIMEL was higher in Atg5−/− iMEFs compared to wild-type cells, and Atg5−/− iMEFs also exhibited a more pronounced increase in BIMEL compared to wild-type when the cells were serum starved (Fig. 3A). However, when we added cycloheximide, which both inhibits protein synthesis and activates ERK1/2, to serum-starved cells we found that the turnover of BIMEL at 3 and 6 hours was essentially identical between the two cell types (Fig. 3A). These results suggest that autophagy might contribute to determining the basal level of BIMEL but plays little or no role in acute ERK1/2-driven turnover of BIMEL.
Since the first description (Ley et al., 2003), dozens of laboratories have shown that ERK1/2-driven turnover of BIMEL is proteasome-dependent in a wide variety of cell types, as judged by the use of small molecule proteasome inhibitors including MG132, bortezomib (Velcade), and lactacystin. We used these same inhibitors to examine the acute turnover or long-term accumulation of HA–BIMELΔKK. MG132 was able to effectively inhibit the ΔRAF-1:ER*-driven turnover of HA–BIMELΔKK (Fig. 3B) and similar results were obtained with bortezomib (Fig. 3C). Furthermore, when cells were transfected with BIMEL constructs and treated chronically with lactacystin, both BIMEL and BIMELΔKK proteins accumulated to the same degree and with the same kinetics (Fig. 3D). Together these results suggested the presence of an alternative poly-Ub-independent pathway for proteasome-dependent degradation of BIMELΔKK following ERK1/2 activation.
BIMEL and BIMELΔKK are degraded by 20S proteasomes
A recent structural study reported that BIM is an IDP (Hinds et al., 2007), although the functional consequences of this were not investigated. There is a growing appreciation that some disordered proteins can be degraded by uncapped 20S proteasomes independently of poly-Ub (Tsvetkov et al., 2008; Baugh et al., 2009); indeed, cleavage by the 20S proteasome has been proposed as an operational definition for IDPs (Tsvetkov et al., 2008). We used FoldIndex (Prilusky et al., 2005) to assess the distribution of folded and unfolded regions in BIMEL using p21CIP1 and PCNA (proliferating cell nuclear antigen) as comparators. This confirmed that p21CIP1 was extensively unfolded (Kriwacki et al., 1996), whereas PCNA was almost exclusively folded (Fig. 4A), consistent with the PCNA crystal structure (Gulbis et al., 1996). In comparison, BIMEL was largely unfolded, notably at the N-terminus and towards the C-terminus, though not including the C-terminal hydrophobic tail. Similar results were obtained when we used IUPred (Dosztanyi et al., 2005), to predict regions of disorder (supplementary material Fig. S3), confirming that BIMEL is an IDP (Hinds et al., 2007).
Prompted by this, we investigated whether BIMEL was degraded by 20S proteasomes. We first generated BIMEL in vitro in a coupled transcription and translation (T&T) reaction and analysed the products using the GST–Dsk2 pulldown assay. Wild-type BIMEL synthesized in vitro was poly-ubiquitylated and this was reduced in the BIMELΔK mutant and abolished in the BIMELΔKK mutant (Fig. 4B), reflecting previous observations in cells (Fig. 2A). To assess their degradation, [35S]methionine-labelled BIMEL or BIMELΔKK were synthesized in the T&T reaction and incubated with purified 20S proteasomes. In common with p21CIP1 (Touitou et al., 2001; Tsvetkov et al., 2008), both BIMEL and BIMELΔKK were rapidly degraded by 20S proteasomes whereas PCNA, a folded, globular protein was not (Fig. 4C). Thus, BIMEL is an IDP that can be degraded by 20S proteasomes in the absence of poly-Ub.
One of the earliest consequences of ERK1/2-dependent phosphorylation of BIMEL is promotion of its dissociation from pro-survival BCL-2 proteins such as MCL-1 or BCL-xL (Ewings et al., 2007); indeed, the BIMELΔBH3 mutant, with three point mutations in its BH3 domain, is defective for binding to MCL-1 or BCL-xL and exhibits accelerated turnover in cells in the absence of ERK1/2 signalling (Ewings et al., 2007). Some IDPs are protected from 20S proteasomal degradation by interactions with their partner proteins (Alvarez-Castelao and Castaño, 2005; Tsvetkov et al., 2008) and, consistent with this, we observed that when incubated with recombinant MCL-1, recombinant HA–BIMEL was protected from degradation by 20S proteasomes (Fig. 4D,E). This protection was not complete, but then it is likely that not all the MCL-1 protein produced in the T&T reaction was correctly folded to allow binding of all the BIMEL. In addition, we noted that MCL-1 was itself partially degraded by 20S proteasomes (supplementary material Fig. S4A), consistent with recent reports (Stewart et al., 2010). Thus the effects of MCL-1 in this assay are probably an underestimate. The specificity of this effect was underlined by the demonstration that BIMELΔBH3, which is defective for MCL-1 binding, was not protected by pre-incubation with recombinant MCL-1 (supplementary material Fig. S4B). Thus, binding of the BH3 domain of BIMEL to one of its biological targets, MCL-1, protects it from degradation by 20S proteasomes.
Because activation of ERK1/2 can promote the poly-Ub of BIMEL and its rapid turnover we speculated that Ub-dependent degradation by the 26S proteasome (UD/26S) and Ub-independent degradation by the 20S proteasome (UI/20S) pathways might operate in parallel, with the UD providing rapid and efficient targeting and the UI pathway serving as a failsafe, back-up pathway. Indeed, degradation of IDPs by the 20S proteasome has been described as ‘degradation by default’ (Asher et al., 2006; Tsvetkov et al., 2009b) and the fact the BIMELΔKK turned over following ERK1/2 activation served as some support for this model. On the basis of this, we reasoned that inhibition of the UD/26S pathway might not prevent BIMEL turnover. To test this we knocked down the Rpn2 subunit of the 19S proteasome cap to prevent the assembly of 26S proteasomes without affecting 20S proteasomes (Tsvetkov et al., 2009a). In cycling cells, knockdown of Rpn2 was effective and caused the accumulation of Cdc25A, a folded protein that is degraded by the UD/26S pathway (Fig. 5A); however, this had no effect on basal BIMEL expression, suggesting the possibility of 26S-independent degradation. However, we were concerned that the low basal level of ERK1/2 and RSK activity in cycling HEK293 cells was not sufficient to provide a strong signal for BIMEL turnover so that conditions were not optimal for observing any effects of Rpn2 knockdown. Instead, HR1 cells were transfected with RNAi oligonucleotides and subsequently stimulated with 4-HT to promote rapid BIMEL turnover. Knockdown of Rpn2 caused some delay in the turnover of BIMEL at early time points but ultimately did not prevent it, so that after 6 hours of stimulation the BIMEL turnover was equal in both cell populations (Fig. 5B,C). Because BIMELΔKK is turned over in a proteasome-dependent manner (Fig. 3) these results support the hypothesis that UD/26S and UI/20S pathways for BIMEL turnover can operate in parallel, and that when the UD/26S pathway is inhibited BIMEL turnover can still proceed by 20S-dependent degradation, albeit after a delay.
Inhibition of cullin-based E3 ligases has no effect on basal BIMEL expression or ERK1/2-driven BIMEL turnover
It has recently been shown that coordinated phosphorylation of BIMEL by ERK1/2 and RSK provides a binding site for SCFβ-TrCP1/2, which promotes BIMEL poly-Ub (Dehan et al., 2009). The Skp1/Cul1/F-box protein (SCF) complexes are perhaps the best-understood RING-type E3s (Cardozo and Pagano, 2004; Nakayama and Nakayama, 2006; Frescas and Pagano, 2008) and consist of a catalytic core (Cul1 and a RING protein) linked by an adaptor (SKP1) to a substrate-specific receptor subunit (the F-box protein). Recognition by F-box proteins often requires phosphorylation of the substrate, providing a link between signalling, poly-Ub and protein turnover. Examples of SCF complexes include SCFFBXW7, which promotes the destruction of cyclin E (Koepp et al., 2001); SCFSkp2, which promotes the destruction of p21CIP1 and p27KIP; and SCFβ-TrCP1/2, which promotes destruction IκB and Cdc25A (for a review, see Frescas and Pagano, 2008). If the UI/20S pathway could substitute for UD/26S, then we reasoned that disrupting BIMEL poly-Ub by targeting its E3 ligase might not affect the levels of endogenous BIMEL. To address this we inhibited cullin function using interfering mutants or RNAi-mediated knockdown of Cul1.
First, we expressed a dominant-negative interfering mutant of cullin1 (dnCul1) (Shirogane et al., 2005) in cycling HEK293 cells and examined the impact on basal protein expression. This approach was validated by showing that dnCul1 caused a substantial accumulation of p27KIP1, cyclin E and Cdc25A, all of which are recognized SCF substrates (Fig. 6A); indeed, like BIM, Cdc25A is a target of SCFβ-TrCP1/2 EL (for a review, see Frescas and Pagano, 2008). Despite this, dnCul1 did not cause an increase in the basal levels of BIMEL (Fig. 6A). Because activation of ERK1/2 promotes BIMEL turnover, we again reasoned that cycling cells were not optimal for BIMEL turnover. As an alternative we again used HR1 cells, where activation of the ΔRAF-1:ER*–MEK1/2–ERK1/2 pathway results in a rapid and robust turnover of BIMEL (Ewings et al., 2007). For these experiments we also used a dominant-negative mutant of Ubc12 (dnUbc12) (Amir et al., 2001), a protein that catalyses the NEDD8 conjugation of a conserved lysine residue that is required for the function of all cullins. The efficacy of dnUbc12 was validated by showing that it also caused accumulation of p27KIP1 when expressed in cycling HEK293 cells, just like dnCul1 (Fig. 6B). Despite this, neither dnCul1 or dnUbc12 could block ΔRAF-1:ER*-driven turnover of BIMEL (Fig. 6C). Finally, knockdown of Cul1 in HR1 cells by RNAi was also without effect on ΔRAF-1:ER*-driven turnover of BIMEL (Fig. 6D).
These results revealed that selective inhibition of cullin1 (by two strategies) or inhibition of all cullins (using dnUbc12) failed to impair ERK1/2-dependent BIMEL turnover. This data could suggest the existence of other E3 Ub ligases for BIMEL in addition to SCFβ-TrCP1/2 (Dehan et al., 2009), but they are also consistent with our demonstration of an alternative Ub-independent pathway for BIMEL degradation.
Poly-Ub of a target protein determines the fate of that protein; K48-linked chains provide a signal for proteasomal degradation whereas K63-linked chains are important in the assembly of pro-inflammatory signalling complexes and protein trafficking (Ikeda and Dikic, 2008; Komander, 2009). The nature of the polyubiquitin chain linkage is typically defined by using individual Ub point mutants (K48R, K63R, etc.) to compete with endogenous Ub. However, such approaches have limitations (see Newton et al., 2008), require substantial overexpression of the mutant Ub (which can be difficult to achieve), and in our hands gave variable results. As an alternative we have made use of the ability of certain UBDs to discriminate between different Ub chain linkages (Komander et al., 2009). Such specificity underpins the use of ubiquitylation as a regulatory signal in a variety of cellular processes; however, in this instance we have used it simply as a diagnostic tool. The Dsk2 and Mud1 UBA domains have been defined as K48-specific by comparing their ability to bind chemically synthesized K48-linked, K63-linked and linear poly-Ub chains; similar studies have defined the K63 specificity of the TAB2C NZF domain (Kulathu et al., 2009). In addition, crystal structures have revealed the molecular basis by which the UBA domains from Dsk2 and Mud1 utilize the unique conformational features of K48-linked chains for specific recognition (Trempe et al., 2005; Lowe et al., 2006). Although the data are still at an early stage, emerging studies suggest that K11 linkages might be a second proteasomal degradation signal (Xu et al., 2009; Wu et al., 2010). K11-linked chains are compact and structurally distinct from K48-linked chains (Bremm et al., 2010) and neither Mud1 UBA or TAB2C NZF domains are able to interact with K11-linked poly-ubiquitin chains in pulldown experiments (Kulathu et al., 2009) (D.K., unpublished). Thus, the results of testing against all currently available Ub chain types strongly suggest that the Dsk2 and Mud1 UBA domains are specific for K48 chains, thereby validating our approach of using these UBA domains as diagnostic tools. Accordingly, our results demonstrate for the first time that BIMEL is subject to K48-linked poly-Ub and define a simple assay for monitoring this linkage specificity in cells. We believe that such an assay might be more generally applicable to the study of K48-linked poly-Ub of proteins and are currently testing this.
Two pathways for ERK1/2-driven proteasomal degradation of BIMEL
The ERK1/2 signalling pathway is the major pathway controlling the proteolytic turnover of BIMEL (Ley et al., 2003) (for a review, see Gillings et al., 2009). Coordinated phosphorylation by ERK1/2 and RSK1/2 targets BIM for poly-Ub by SCFβ-TrCP EL (Dehan et al., 2009). Despite this, we found that a BIMELΔKK mutant failed to undergo poly-Ub but was still subject to ERK1/2-driven, proteasome-dependent turnover. Prompted by the suggestion that BIMEL is an IDP (Hinds et al., 2007), we found that BIMEL (wild-type or ΔKK) could be degraded in a Ub-independent manner by 20S proteasomes. Further evidence for Ub-independent turnover of BIMEL came from the demonstration that Rpn2 knockdown delayed, but did not prevent, BIMEL turnover. Finally, inhibition of cullin-based E3 Ub ligases disrupted the turnover of p27KIP1, cyclin E and Cdc25A (the latter a validated target of SCFβ-TrCP1/2) but had no effect on BIMEL turnover. Taken together, these results provide strong, independent lines of evidence for an additional Ub-independent pathway for BIMEL turnover by the proteasome.
We suggest that ERK1/2-driven degradation of BIMEL proceeds by two pathways: classical Ub-dependent degradation by the 26S proteasome (UD/26S) or Ub-independent degradation by the 20S proteasome (UI/20S). In the former, phosphorylation of BIMEL by ERK1/2 and RSK will allow the E3 ligase SCFβTrCP to promote the poly-Ub and 26S-dependent destruction of BIMEL (Dehan et al., 2009). However, because ΔRAF-1:ER* can still promote the turnover of BIMELΔKK, it is apparent that ERK1/2 activation can also promote BIMEL turnover by the proteasome, independently of poly-Ub. UI/20S degradation would explain why we could not prevent BIMEL turnover by ΔKK mutation, Rpn2 knockdown or inhibition of cullin-based E3 ligases.
UI/20S is thought to be an important and evolutionarily conserved proteolytic pathway. Uncapped 20S proteasomes are abundant in mammalian cells and degrade up to 20% of cellular proteins (Baugh et al., 2009), including some proteins involved in cell cycle control and apoptosis. For example, the tumour suppressor p53 is degraded by a classical UD/26S mechanism but its N-terminus is disordered, allowing degradation by 20S proteasomes (Asher et al., 2005a; Asher et al., 2006; Tsvetkov et al., 2009a). Similarly, ornithine decarboxylase (Asher et al., 2005b), p21CIP1 (Touitou et al., 2001) and IκBa (Krappmann et al., 1996; Alvarez-Castelao and Castaño, 2005) are all poly-Ub but can also be degraded by 20S. This default degradation pathway might serve as a back-up to ensure timely removal of these biologically important proteins (Asher et al., 2006).
BIMEL, MCL-1 and the nanny model
We previously showed that phosphorylation of BIMEL promotes its dissociation from pro-survival BCL-2 proteins; indeed, BIMELΔBH3, which is defective for binding to pro-survival BCL-2 proteins, is turned over more rapidly than wild-type BIMEL suggesting that dissociation contributes to BIMEL turnover (Ewings et al., 2007). Interestingly, some IDPs are protected from 20S degradation by interactions with their partner proteins (Alvarez-Castelao and Castaño, 2005; Tsvetkov et al., 2008; Tsvetkov et al., 2009a) and this has led to the suggestion that such partner proteins serve as ‘nannies’ (Tsvetkov et al., 2009b). Our demonstration that binding of BIMEL to MCL-1 could protect it from degradation by 20S proteasomes (Fig. 4D,E) suggests that MCL-1 and presumably other pro-survival BCL-2 proteins serve as nannies for BIMEL. In the course of writing up this work, a study reported that MCL-1 is degraded by 20S proteasomes in a Ub-independent manner (Stewart et al., 2010); the authors speculated that this was dependent on the disordered N-terminus of MCL-1. There are remarkable parallels with our study on BIM; both proteins are subject to poly-Ub and turnover by the UD/26S pathway but both proteins can also be degraded by the UI/20S pathway when poly-Ub is prevented by mutation of all lysine residues. Studies have previously shown that the binding of BH3-only proteins can influence MCL-1 stability in cells; Noxa binding promoted MCL-1 degradation (Willis et al., 2005), whereas binding of BIM (Czabotar et al., 2007) or PUMA (Mei et al., 2005) promoted MCL-1 stabilization. Although binding of BH3-only proteins could stabilize MCL-1 by displacing the E3 ligase MULE (Warr et al., 2005; Zhong et al., 2005), these new studies suggest an additional explanation whereby BIMEL and MCL-1 serve as nannies for each other to prevent their degradation by 20S. The biological consequences of this are likely to be complex: dissociation of BIMEL from MCL-1 (Ewings et al., 2007) would facilitate degradation of BIMEL, which would support cell survival; conversely, this might also facilitate destruction of MCL-1 by the UD/26S or UI/20S pathways, which would tend to support cell death. The net effect is likely to be determined by the expression of other BCL-2 family proteins.
On the basis of these observations we propose that in addition to poly-Ub by β-TrCP (Dehan et al., 2009), phosphorylation-dependent dissociation from BCL-2 proteins represents a signal for targeting BIMEL to the 20S proteasome. This might be because ‘free’ BIMEL is intrinsically disordered or because dissociation of BIMEL unmasks disordered regions or favours a disordered conformation that is a prerequisite for access to 20S proteasomes (Baugh et al., 2009). Phosphorylation of BIMEL might contribute directly, because high net charge is often associated with disorder (Dyson and Wright, 2005). In this scenario, UD/26S would provide a rapid, targeted destruction mechanism, whereas the UI/20S pathway might operate to remove any excess or ‘free’ BIMEL and serve as a ‘failsafe’ to ensure the removal of BIMEL if ubiquitylation is disrupted. Quite how BIMEL accesses the 20S proteasome remains to be seen. Non-Ub p21CIP1 can bind to the REGγ 20S proteasome regulator (Li et al., 2007; Chen et al., 2007) or might be recognized by the C8α subunit of the 20S proteasome (Touitou et al., 2001). To date, we have not been able to detect specific binding of BIMEL to C8α (M.J., C.M.W. and S.J.C., unpublished observations).
In summary we have used the inherent linkage specificity of the UBA domains of Dsk2 and Mud1 as a diagnostic tool to demonstrate that BIMEL undergoes ERK1/2-driven, K48-linked poly-Ub. Despite this, poly-Ub is not a prerequisite for proteasomal degradation; we suggest that BIMEL can also be degraded in a Ub-independent fashion by 20S proteasomes following its dissociation from pro-survival BCL-2 proteins, by virtue of its intrinsic disorder. These different mechanisms of degradation reflect the biological imperative of destroying BIMEL in a timely fashion. BIM is one of the most toxic BH3-only proteins because of its ability to engage with all the pro-survival BCL-2 proteins (Chen et al., 2005). Consequently, its abundance must be tightly regulated by multiple mechanisms to ensure that cell death is only initiated under appropriate conditions.
Materials and Methods
Cells and cell culture
Culture of HR1 cells has been described previously (Ewings et al., 2007). SV40-immortalized MEFs from wild-type or Atg5−/− mice (Kuma et al., 2004), a kind gift from Noboru Mizushima (Tokyo Medical and Dental University, Tokyo, Japan), were provided by Aviva Tolkovky (Universtiy of Cambridge, Cambridge, UK).
Antibodies that specifically recognize K48-linked or K63-linked Ub chains (Newton et al., 2008) were from Millipore. Other antibodies were obtained from the following vendors: cyclin E Ab-1 from Neomarkers; Cdc25A F6, cullin 1 D5 and anti-HA from Santa Cruz Biotechnology; p27KIP1 Ab-2 from Calbiochem; BIM from Chemicon; phosphorylated ERK1/2 (P-ERK1/2) and ERK1/2 from Cell Signaling Technology; anti-His from Amersham Biosciences; and anti-FLAG from Sigma.
His-tagged dominant-negative UBC12(C111S) in pcDNA3.1 was provided by Aaron Ciechanover (Technion, Haifa, Israel) (Amir et al., 2001). FLAG-tagged dominant-negative cullin 1 (dnCul1) was provided by Wade Harper (Harvard Medical School, Boston, MA) (Shirogane et al., 2005). The following UBD constructs were used: GST–Dsk2 UBA, wild-type (Ohno et al., 2005) and an inactive variant harbouring M342R and F344A point mutations; GST–Mud1 UBA, wild-type and the F330A mutant (Trempe et al., 2005); and GST–Tab2c NZF (Komander et al., 2009). GST–Dsk2 and GST–Mud1 fusions were expressed in DH5a cells, purified and immobilized on beads as described for GST–BIM (Ley et al., 2004). GST–Tab2c NZF (Komander et al., 2009) was expressed in BL21 cells and purified as described previously (Berrow et al., 2007). HA–BIMEL (rat sequence) has been described previously (Ewings et al., 2007); K3R and/or K108R mutations in BIMEL were introduced by site-directed mutagenesis.
Assay of BIMEL poly-Ub in cells
HR1 cells were transfected with the indicated plasmids. After 24 hours, cells were serum starved in the presence of U0126 to inactivate ERK1/2 or treated with 4-HT + MG132 to activate ERK1/2 and inhibit the proteasome. Following lysis in TG lysis buffer (Ley et al., 2003) cell extracts were retained (input) or incubated end-over-end with immobilized GST–UBDs at 4°C for 2 hours, washed and fractionated by SDS-PAGE as described for GST–BIM pulldowns (Ley et al., 2004).
Assay of BIMEL turnover in cells
HR1 cells were transfected with the indicated BIMEL plasmids. After 24 hours, cells were treated with emetine to block new protein synthesis and chased in serum-free medium or in the presence or absence of 4-HT, U0126 or MG132. Cell extracts were fractionated by SDS-PAGE and immunoblotted as described.
In vitro translation
In vitro translation was performed using the T&T Quick Coupled Transcription/Translation System (Promega, Madison, WI). The plasmids employed were pCDNA3-BIMEL, pCDNA3-BIMELΔKK, pCDNA3-BIMELΔBH3, pCDNA3-MCL-1, pIRES-p21CIP1 and pIRES-PCNA.
In vitro 20S proteasomal degradation assay
Purified 20S proteasomes were generated as described previously (Asher et al., 2005b). [35S]methionine-labelled proteins translated in vitro were incubated with 1 μg of purified 20S proteasomes in 150 mM NaCl, 50 mM Tris HCl, pH 7.5 at 37°C for the indicated times. Reactions were then resolved by SDS-PAGE, visualized by autoradiography and quantified by phosphorimaging (Tsvetkov et al., 2008).
RNAi against Cul1 and Rpn2
HR-1 cells (150,000 cells per well plated in a six-well plate) were plated in antibiotic-free media. Transfection complexes were prepared according to the manufacturer's instructions using Lipofectamine 2000 (Invitrogen). For Rpn2 knockdown, 200 pmol siRpn2-PT (5′-GCTCATATTGGGAATGCTTAT-3′) and 200 pmol siRpn2-MJ (5′-GGATACTTCTCCAGGATCA-3′) were mixed and 400 pmol of a control siRNA used (murine Bim1 5′-GGAGGAACCTGAAGATCTG-3′). Complexes were applied and cells incubated for 48 hours before being aspirated and fresh transfection complexes applied for a further 48 hours. Cells were then harvested immediately for western blot analysis, or an emetine chase experiment performed. For Cul1 knockdown, a pool of three siRNAs targeting Cul1 was used (Santa Cruz Biotechnology) alongside a human siRNA control (Santa Cruz Biotechnology). HR-1 cells were transfected for 24 hours, the transfection media was then aspirated and serum-free media applied. Following 16 hours of serum-free treatment, emetine (50 μM) was added to block protein synthesis followed 30 minutes later by addition of 4-HT for 8 hours.
We thank Rebecca Gilley and other members of the Cook laboratory for advice and encouragement and John Pascal (The Babraham Institute, Cambridge, UK) for initial advice on T&T reactions. We also thank Jane Endicott and Jean-Francois Trempe (University of Oxford, Oxford, UK), for providing the GST–Mud1 UBA constructs; Aaron Ciechanover, for dnUbc12 and J. Wade Harper, for dnCul1. This work was funded by core funding to the Babraham Institute and a response mode project grant from the BBSRC (BB/E02162X/1). C.M.W. was supported by a BBSRC PhD studentship and C.L.J. was supported by a BBSRC/AstraZeneca CASE studentship.