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The ~300 human Cullin-RING ligases (CRLs) are multisubunit E3s in which a RING protein, either RBX1 or RBX2, recruits an E2 to catalyze ubiquitination. RBX1-containing CRLs also can bind Glomulin (GLMN), which binds RBX1’s RING domain, regulates the RBX1-CUL1-containing SCFFBW7 complex, and is disrupted in the disease Glomuvenous Malformation. Here we report the crystal structure of a complex between GLMN, RBX1, and a fragment of CUL1. Structural and biochemical analyses reveal that GLMN adopts a HEAT-like repeat fold that tightly binds the E2-interacting surface of RBX1, inhibiting CRL-mediated chain formation by the E2 CDC34. The structure explains the basis for GLMN’s selectivity toward RBX1 over RBX2, and how disease-associated mutations disrupt GLMN-RBX1 interactions. Our study reveals a mechanism for RING E3 ligase regulation whereby an inhibitor blocks E2 access, and raises the possibility that other E3s are likewise controlled by cellular proteins that mask E2-binding surfaces to mediate inhibition.
Ubiquitin (Ub) ligation is highly regulated through the action of specific E1, E2, and E3 enzymes. Via a series of E1-dependent reactions, Ub’s C-terminus becomes covalently linked by a thioester bond to the E2’s active site cysteine. The resulting labile E2~Ub (~ denotes covalent bond) intermediate interacts with an E3 ligase, which recruits the substrate protein and promotes Ub transfer from the E2 to the substrate. Given the importance of ubiquitination in determining the fates of modified targets, it is of great interest to understand structural mechanisms by which E3 ligase activities are controlled.
Much knowledge of E3 regulation derives from investigations of the cullin-RING ligases (CRLs), which account for nearly half the predicted human E3s (Deshaies and Joazeiro, 2009). CRLs are multisubunit enzymes, nucleated by a cullin-RING catalytic core (Zimmerman et al., 2010). A cullin’s N-terminal domain (NTD) assembles with substrate-binding receptors (SRs), such as F-box proteins in the prototypic “SCF” (SKP1-CUL1-F-box protein) CRLs. Crystal structures have revealed how phosphorylation regulates binding of substrates such as CyclinE and p27 to SCFFBW7 and SCFSKP2-CKSHS1, respectively (Hao et al., 2007; Hao et al., 2005; Orlicky et al., 2003; Wu et al., 2003), and how other modifications or co-association with small molecules can activate substrate binding to other SRs (reviewed in (Duda et al., 2011)). Although mechanisms restraining substrate interactions with CRLs are less understood, structures revealed how the Mitotic Checkpoint Complex inhibitor blocks substrate binding to the highly divergent CRL, the Anaphase Promoting Complex (Buschhorn et al., 2011; Chao et al., 2012; da Fonseca et al., 2011; Herzog et al., 2009).
A cullin’s C-terminal domain (CTD) binds a catalytic RING-containing protein, which for cullins 1–4 is RBX1 and for CUL5 is RBX2 (Huang et al., 2009; Kamura et al., 1999b; Kamura et al., 2004; Ohta et al., 1999; Seol et al., 1999; Skowyra et al., 1999; Tan et al., 1999; Zheng et al., 2002b). The CTD comprises: a 4-helix bundle (4HB) linked to the cullin NTD; an α/β subdomain that forms an intermolecular β-sheet with a strand from RBX; and the extreme C-terminal WHB subdomain, which packs against and autoinhibits the RBX protein’s catalytic RING domain (Duda et al., 2008; Yamoah et al., 2008; Zheng et al., 2002b). CRLs are activated by autocatalytic covalent ligation of the Ub-like protein NEDD8 (Calabrese et al., 2011; Duda et al., 2008; Kamura et al., 1999a; Saha and Deshaies, 2008; Yamoah et al., 2008). This is thought to stimulate orientational flexibility of the RBX RING domain so that an associated Ub E2, such as CDC34, can be placed in multiple positions associated with processive polyubiquitination of substrate (Duda et al., 2008; Pierce et al., 2009; Saha and Deshaies, 2008; Wu et al., 2010; Yamoah et al., 2008). Notably, numerous factors have evolved to counteract neddylation: (1) the inhibitor CAND1 binds RBX1-CUL1 both at the SKP1/F-box protein-binding site and at the CUL1 WHB-RBX1 RING junction, simultaneously blocking both cullin-SR interactions and NEDD8 ligation (Goldenberg et al., 2004; Liu et al., 2002; Zheng et al., 2002a); (2) the deneddylating enzyme CSN removes NEDD8 thereby deactivating CRLs (Lyapina et al., 2001); and (3) the Gram-negative pathogenic bacterial effector Cif (cyclin inhibiting factor) inactivates neddylated cullins by deamidating Gln40 on NEDD8 (Cui et al., 2010; Jubelin et al., 2010; Morikawa et al., 2010).
Although structural data have provided important insights into mechanisms activating and inhibiting CRL-substrate binding, assembly, and neddylation (reviewed in (Duda et al., 2011; Zimmerman et al., 2010)), relatively less is known about structural mechanisms by which cellular proteins can attenuate Ub ligase activities of CRLs. We address this through studies of Glomulin (GLMN), a protein principally expressed in vascular smooth muscle cells (McIntyre et al., 2004). In humans, mutations of the GLMN gene cause the disorder Glomuvenous Malformation (GVM), which presents with nonmetastatic cutaneous vascular lesions resembling glomus tumors (Borroni et al., 2011; Brouillard et al., 2002; Brouillard et al., 2005). Prior work showed that the GLMN protein binds RBX1 but not RBX2-associated cullins in cells, that GLMN binds the isolated RBX1 RING domain, and that GLMN inhibits SCFFBW7-mediated ubiquitination of a CyclinE substrate in vitro (Tron et al., 2012). As with some other known CRL regulators, loss of GLMN influences only a subset of CRL activities in vivo (Arai et al., 2003; Tron et al., 2012). Knocking out GLMN in mice results in embryonic lethality due to severe developmental defects that partially mimic effects of ablating FBW7 (Tron et al., 2012). Decreased GLMN activity in cells leads to a striking CRL- and proteasome-dependent reduction in the level of FBW7, and a concomitant increase in levels of SCFFBW7 substrates CyclinE and c-Myc including in GVM patient samples (Tron et al., 2012). In order to gain insights into mechanisms by which GLMN can control SCF activities, we performed crystallographic, biochemical, and biophysical analyses of GLMN interactions with RBX1. Our data collectively reveal that GLMN binds and masks the E2 binding surface of RBX1’s RING domain to inhibit E3 ligase activity.
To understand the basis for GLMN-RBX1 interactions, we determined the 3.0 Å crystal structure of a complex between GLMN, RBX1, and a truncated form of CUL1 containing residues 411–690 that spans from the 4HB to the α/β subdomains (hereafter referred to as CUL1[4HB-α/β]), with two complexes per asymmetric unit (Table 1, HFig. 1, Fig. S1). The GLMN structure consists primarily of α-helices arranged as tandem antiparallel pairs, which form a double layer resembling HEAT (untingtin-Elongation-A subunit-TOR) repeats (Fig. 1A). Although HEAT-like repeats are often continuous and form superhelical solenoid structures (Groves and Barford, 1999), in GLMN the continuity is interrupted by a central helix lying roughly perpendicular to both adjacent repeats. As a result, the GLMN HEAT-like repeats are arranged into N-terminal and C-terminal domains, each containing 5 α-helical pairs plus some additional flanking helices. Despite no apparent sequence homology, GLMN’s two domains superimpose on each other with ~3 Å root-mean-square deviation. The connection between GLMN’s N- and C-terminal domains is somewhat flexible, as revealed by a ~10° difference in their relative orientations in the two molecules in the asymmetric unit (Fig. 1B). This agrees with our observations in solution using SAXS described below. Nonetheless, in both cases GLMN adopts a ~110 Å elongated structure, resembling a slightly bent finger.
The two RBX1-CUL1 portions of the asymmetric unit superimpose on each other. CUL1[4HB-α/β] and RBX1’s β-strand (residues 19–25), embedded in an intermolecular β-sheet within the CUL1 α/β-subdomain, superimpose on all prior CRL structures (Fig. S1H) (Angers et al., 2006; Calabrese et al., 2011; Duda et al., 2008; Fischer et al., 2011; Goldenberg et al., 2004; Zheng et al., 2002b). In isolation, RBX1’s RING domain also superimposes on all previous structures (Fig. S1H). However, in comparison to prior structures, the relative orientations differ for RBX1’s RING domain with respect to RBX1’s strand embedded in the CUL1 α/β-subdomain (Fig. S1I). This is consistent with the previously observed potential for RBX1 RING domain mobility (Calabrese et al., 2011; Duda et al., 2008; Fischer et al., 2011).
A 25 Å × 35 Å surface roughly centered on Ile44 of RBX1’s RING domain is essentially entirely covered by a platform formed by one face of the three C-terminal helical repeats that constitute GLMN’s “fingertip” (Fig. 1C). This agrees with our systematic deletion mutational analysis of GLMN, which mapped this identical region as required for RBX1 RING-binding (Fig. S2). The interaction buries ~880 Å2 from RBX1’s RING domain and ~825 Å2 from GLMN.
In order to interpret functional roles of the GLMN-RBX1-CUL1 interactions, we wished to model GLMN bound to RBX1-CUL1 complexed with other components of SCF E3 complexes. However, in the crystals, GLMN not only contacts RBX1’s RING domain, but also makes some contacts with the linker between RBX1’s β-strand and RING, and also portions of CUL1 (Fig. 1C, S1J). It is not possible to model GLMN interacting with larger SCF complexes simultaneously via all the crystallographically-observed contacts due to differences in relative orientations of RBX1’s RING between prior structures and the complex with GLMN (Angers et al., 2006; Calabrese et al., 2011; Duda et al., 2008; Fischer et al., 2011; Goldenberg et al., 2004; Zheng et al., 2002b). Thus, to determine interactions in solution, we performed several experiments that reveal stoichiometric recruitment of GLMN to a variety of RBX1-containing complexes via direct interactions with the RING domain (Fig. 2).
First, we considered that RBX1’s RING domain favors different relative orientations in different SCF complexes. Two extremes of RBX1 RING conformation are observed for RBX1-CUL1-CAND1, in which CAND1 rigidifies RBX1’s RING domain and locks it against CUL1’s WHB subdomain, and RBX1-cullin~NEDD8, in which RBX1’s RING domain is apparently more flexibly tethered and rotating (Duda et al., 2008; Goldenberg et al., 2004). Analysis of complex formation by gel filtration chromatography showed that GLMN associates stoichiometrically with the two very distinctive RBX1-complexes that also contain the “split ‘n coexpress” (Zheng et al., 2002b) version of full-length CUL1 (Fig. 2A, 2B).
Second, in order to test dependence of GLMN-cullin interactions on RBX1’s RING domain, we performed gel filtration chromatography of 1:1 mixtures of GLMN with RBX1 complexed with CUL1’s isolated C-terminal domain, or a mutant complex deleted for RBX1’s RING domain. The data show that GLMN forms a stoichiometric complex with RBX1-CUL1CTD, and that the RBX1 RING is necessary for this interaction (Fig. 2C).
Third, we used enzymatic competition assays to qualitatively compare GLMN’s interactions with RBX1’s isolated RING domain versus RBX1 assembled into active CRL complexes. We previously showed that addition of excess GLMN inhibits in vitro ubiquitination of a CyclinE phosphopeptide by SCFFBW7 (Tron et al., 2012). To gain insights into the ratio of GLMN:SCFFBW7 required for inhibition, we performed a titration experiment. Some inhibition is observed at a 1:1 ratio of GLMN:SCF, with more GLMN nearly completely squelching ubiquitination (Fig. 2D). If GLMN functions by binding to RBX1’s RING domain, then in vitro, in the absence of any cellular factors influencing regulation, GLMN would be expected to exert similar effects on other SCFs. Indeed, addition of GLMN also inhibited in vitro SCFSKP2-CKSHS1-mediated ubiquitination of phospho-p27 (Fig. 2E). To probe whether access to RBX1’s RING is important for GLMN-mediated SCF inhibition, we added increasing amounts of the isolated RBX1 RING domain to the assays. This relieves the inhibition, presumably by competing GLMN away from SCFs (Fig. 2E, 2F). This raised the possibility that GLMN binds equally well to RBX1’s RING domain in isolation and in active SCF complexes.
Fourth, to quantitatively compare binding between GLMN and RBX1’s RING domain either in isolation or when RBX1 is associated with a cullin, we developed a FRET-based binding assay, similar to that used previously to measure SCF binding to its cognate Ub E2, CDC34 (Kleiger et al., 2009). In this assay, GLMN-YFP binds to CFP-RBX1-CUL1CTD, CFP-RBX1-CUL1CTD~NEDD8, and CFP-RBX1RING with Kds of 36 ± 6 nM, 28 ± 4 nM, and 45 ± 10 nM, respectively (Fig. 2G). Thus, RBX1’s isolated RING domain is sufficient to bind GLMN with high-affinity, with similar interactions for RBX1-cullin complexes. Although the E2, CDC34, also interacts with RBX1-CUL1~NEDD8 with high affinity (Kleiger et al., 2009), the binding modes differ for GLMN, which requires RBX1’s RING domain (Fig. 2C), and CDC34, which does not (Fig. S3). While RBX1’s RING domain binds weakly to CDC34’s E2 core domain (Spratt et al., 2012), a CRL binds CDC34’s C-terminal acidic tail with high affinity even in the absence of CDC34’s E2 catalytic core domain and RBX1’s RING domain (Fig. S3).
Finally, we mutagenized GLMN residues mediating crystal contacts with CUL1 and observed no effect on inhibition (Fig. S1J, K). This is consistent with GLMN binding to RBX1’s RING domain in a range of orientations relative to CUL1, and with the notion that the observed contacts to CUL1 arose from packing during crystallization.
To gain insights into structures of GLMN-RBX1 complexes in solution in the absence of crystal contacts, we performed Small Angle X-ray Scattering (SAXS) studies (Putnam et al., 2007). The broadening in P(r) and Kratky plots indicate higher flexibility for GLMN alone as compared to in complex with RBX1’s RING domain (Fig. 3A, S4). Given the crystallographically-observed conformational flexibility of GLMN (Fig. 1), we reasoned that the coexistence of different GLMN conformations with a hinge between the N- and C-terminal domains could contribute to the experimental SAXS profile. Indeed, an ensemble of two GLMN conformers was required to successfully match the GLMN experimental data (Fig. 3A, 3B, and S4C) (Pelikan et al., 2009). For the GLMN complex with RBX1’s isolated RING domain, an ensemble predominantly corresponding to the GLMN-RBX1RING portion of the crystal and also containing 15% of free GLMN is sufficient to match experimental curve (Fig. 3A, 3B). These data are consistent with GLMN conformational flexibility (Fig. 1B) decreasing upon binding to the RBX1 RING. We also obtained SAXS data for GLMN bound to RBX1 complexes with unneddylated and neddylated “split ‘n coexpress” full-length CUL1 (Zheng et al., 2002b) and CUL1CTD. The data are consistent with GLMN binding the RBX1 RING domain as in our crystal structure, and with prior SAXS analyses of RBX1-CUL1CTD and RBX1-CUL1CTD~NEDD8, which indicated some conformational flexibility in the absence of neddylation, and substantial rotational flexibility of the RBX1 RING domain upon neddylation (Duda et al., 2008). For both unneddylated complexes the SAXS curves and envelopes matched with models obtained by docking GLMN onto the RING domain of RBX1 as in the original RBX1-CUL1 structures (Zheng et al., 2002b), in which the RBX1 RING approaches the CUL1 WHB subdomain (Fig. 2H, ,33 and S4). For the neddylated structures, the SAXS data are consistent with a flexible hinge linking the RBX1-RING and GLMN to RBX1’s N-terminal strand and the remainder of CUL1 (Fig. 3C, 3E, and S4).
Taken together, our assembly, competitive inhibition/relief, quantitative binding, and solution structural data indicate that RBX1’s RING domain is both necessary and sufficient to recruit GLMN to numerous SCF ubiquitin ligase assemblies (Fig. 2H).
The crystal structure reveals that GLMN-RBX1 RING interactions are anchored by two complementary hydrophobic surfaces from both proteins (Fig. 4A). One cluster involves GLMN’s Met472, Leu567, and Val571 packing against RBX1’s Ala43, Ile44, the aliphatic portion of Arg46, Trp87, Pro95, and Leu96, with support from Arg91. A second, smaller hydrophobic cluster lies at the distal end of the complex. Here, Tyr480 and Ile483 from the C-terminal end of the central helix in the GLMN tri-HEAT-like platform interacts with RBX1’s Ile54, Ala58, and hydrophobic portions of Glu55 and Gln57.
The GLMN-RBX1 complex is also stabilized by a vast network of electrostatic interactions, including between GLMN’s Gln422 and Lys425 and RBX1’s Asn47, between GLMN’s Lys425, Asn426, and Asp429 and RBX1’s Gln57, between GLMN’s Asn476 and the backbone oxygen and nitrogen respectively from RBX1’s Ile44 and Arg46, and between GLMN’s Arg479 and Arg574 and RBX1’s Glu55 (Fig. 4A).
We validated the functional importance of the structurally observed contacts between GLMN and RBX1 in four distinct assays. First, we measured binding between wild-type and mutant versions of GLMN and an isolated RBX1 RING domain using Biacore (Fig. 4B, S3). The Kd for wild-type GLMN binding to the RBX1 RING domain measured by Biacore (38.6 +/− 0.8 nM) is similar to that measured in our FRET assay. Ala subsitutions in place of key GLMN interface residues, either individually (Leu567 or Arg574), or pairwise substitutions in place of Asn476 and either Met472 or Asn479, impaired binding. Similarly, mutation of key contact residues from the RBX1 RING domain - Glu55, Gln57, or Arg91 - also decreased binding. Second, we tested the effects of mutations on the ability of HA-tagged GLMN to coimmunoprecipitate RBX1 when expressed in HEK293T cells. Indeed, the structure-based mutations diminished GLMN binding to endogenous RBX1 and its associated CUL1 in cells (Fig. 4C). As controls, the GVM disease-associated frameshift mutation with a 2-base deletion at position 1711 also eliminated the interaction (Tron et al., 2012), and Ala substitutions in GLMN at a variety of surface residues distal from the RBX1-binding surface (Glu458, Asn503, and Glu580) had no effect. Third, for our two different model Ub ligation systems, GLMN’s inhibitory activity was impaired by individual Ala substitutions for RBX1-contacting Lys425, Asn476, Leu567, and Arg574, or pairwise substitutions in place of Asn476 and either Met472 or Asn479 (Fig. 4D). Fourth, in our enzymatic competition assays, mutations in RBX1 that hinder binding to GLMN also impair relief from GLMN-inhibition of SCF E3 activities (Fig. 4E). Thus, the structurally-observed contacts mediate interactions between GLMN and the RBX1 RING domain in solution, and are critical for GLMN inhibition of RBX1-mediated E3 functions.
Validated for GLMN-RBX1 interactions, the crystal structure allowed us to address how GLMN inhibits SCF activation of CDC34-mediated formation of Ub chains on substrates. We made a model of an RBX1-CDC34 complex by docking based on a homologous RING-E2 structure (Fig. 5A) (Ceccarelli et al., 2011; Zheng et al., 2000). We confirmed the model by examining NMR chemical shift perturbation (csp) upon adding CDC34 to 15N-labeled RBX1 RING domain in a 15N-1H TROSY experiment. In agreement with results published during our manuscript revision (Spratt et al., 2012), resonances displaying significant csp map to the canonical E2-binding site on RBX1 (Fig. 5B, 5C).
Next, we superimposed our RBX1-GLMN structure onto the RBX1-CDC34 model. The comparison suggests that GLMN completely masks the E2-binding site on RBX1 (Fig. 5B). Because GLMN is a large protein for NMR experiments, we added substoichiometric amounts of a minimal RBX1-binding domain from GLMN, corresponding to residues 300-594, and observed csp for several of the same RBX1 resonances as upon addition of CDC34 (Fig. 5C, S3). Addition of stoichiometric GLMN300-594 led to severe line broadening and decreased signal intensity for RBX1 resonances, presumably due to the elongated high-affinity >40 kDa RBX1RING-GLMN300-594 complex tumbling slowly on the NMR timescale. This occurred even when 1 molar equivalent of GLMN300-594 was added to a 1:5.5 RBX1RING:CDC34 mixture (Fig. 5D). In a related vein, some GLMN inhibition is observed even with a 1:10 ratio of GLMN to CDC34 in our assays (Fig. 5E). Together, the results suggest that GLMN can outcompete excess CDC34 for binding to the RBX1 RING domain.
Consistent with RBX1 using a common surface to bind GLMN and CDC34, an Ala substitution in place of Arg91 in this surface impairs SCF E3 activity, and the remaining activity is not completely eliminated upon adding GLMN (Fig. 5F). However, the structural data also suggest that RBX1’s Gln57 does not bind CDC34, but uniquely binds GLMN. Indeed, a Gln57Ala mutant RBX1 RING domain was impaired for binding to GLMN (Fig. 4B). In the context of a neddylated SCF complex, the RBX1 Gln57Ala mutant shows wild-type E3 activity in our assay, but is not inhibited by GLMN in agreement with the notion that this RBX1 surface binding to GLMN is essential for inhibition (Fig. 5F).
A key intrinsic E3 function of RBX1’s RING domain in the context of neddylated CUL1-RBX1 complexes is to stimulate CDC34’s innate ability to build a Ub chain even in the absence of substrate (Saha and Deshaies, 2008). Indeed, CDC34-mediated di-Ub chain formation is stimulated by addition of RBX1-CUL1CTD~NEDD8 in a pulse-chase assay, and GLMN inhibits this reaction (Fig. 5G). Inhibition by GLMN depends on binding to RBX1’s RING domain, because Ala substitutions in place of GLMN’s RBX1-binding residues hinder its inhibitory activity (Fig. 5G).
A prior study revealed GLMN as displaying absolute specificity for binding to RBX1 and its associated CUL1-CUL4, but not the nearly 50% identical RBX2 protein or its associated CUL5 in cells (Tron et al., 2012). Furthermore, domain swap experiments showed that GLMN binding in cells depends on the identity of the RBX RING domain. Indeed, we find that GLMN specifically recognizes the RBX1 RING domain, but not the RING domains from RBX2 or the more distal relative APC11 (Fig. S5). The GLMN-RBX1- CUL1[4HB-α/β] crystal structure shows key contact residues differing between RBX1 and RBX2, including RBX1’s Asn47, Glu55, and Arg91 (Fig. S5), which correspond to RBX2’s Val55, Arg63, and Asn96, respectively. Val55 and Asn96 would be unable to make the favorable contacts achieved by their RBX1 counterparts. Furthermore, RBX2’s Arg63 would repel GLMN’s Arg479 and Arg574, which make favorable contacts with RBX1’s Glu55.
The GLMN-RBX1-CUL1[4HB-α/β] structure provides a rationale for understanding the effects of mutations associated with Glomuvenous Malformation disease (Borroni et al., 2011; Brouillard et al., 2002; Brouillard et al., 2005). To date, no missense mutations have been found. Instead, most disease mutations result in premature truncations that delete the entire RBX1-binding region (Fig. S2C). The structure explains why RBX1 cannot bind even to the mutant with the most minimal truncation, which deletes GT at position 1711 and generates a frameshift altering the distal 3 ½ turns of the C-terminal helix in GLMN. This both removes key RBX1-binding residues and would also likely decrease stability due to loss of hydrophobic residues that face the adjacent HEAT-like repeat helices. In a related vein, the only in-frame deletion, of Asn393, would lead to destabilization of GLMN’s C-terminal domain, thereby impairing binding to RBX1.
At a rough level, GLMN may be considered to resemble the other CRL inhibitor CAND1, as both proteins are composed of HEAT/HEAT-like repeats (Fig. 1, ,2H).2H). However, the two inhibitors bind CRLs differently, and function through entirely distinct mechanisms. CAND1 targets unassembled, unneddylated CRLs at two points, and inhibits CRL assembly and activation in 3 ways: (1) by preventing binding to SR complexes, (2) by blocking the NEDD8 acceptor Lys, and (3) by locking RBX1 against the cullin to prevent rotation of RBX1’s RING domain required for neddylation (Goldenberg et al., 2004). In contrast, a larger RBX1 surface distinct from the CAND1-binding site but encompassing the E2-interacting region binds GLMN (Fig. 5, ,6,6, S6). Thus, unlike CAND1, GLMN binding is entirely compatible with CRL assembly and RBX1 RING domain conformational flexibility, but instead targets CRL catalytic function (Fig. 6).
Despite their different modes of inhibition, CAND1 and GLMN have some common properties. Both appear to exert broad biochemical effects on multiple RBX1-associated cullins: the presence of CAND1 leads to decreased cullin neddylation in vitro and in vivo (Bennett et al., 2010; Liu et al., 2002; Zheng et al., 2002a). Similarly, the presence of GLMN leads to decreased CRL-mediated neddylation and ubiquitination in vitro, and absence of GLMN leads to diminished levels of cullins 1–4 and RBX1 in vivo (Fig. 2 and (Tron et al., 2012)). Strikingly, loss of either CAND1 or GLMN in vivo appears to affect only selected CRL substrates in terms of biological function. The levels of many well-studied CRL targets appear uninfluenced by loss of CAND1 in either mammalian cells or worms (Bennett et al., 2010; Bosu et al., 2010; Chua et al., 2011). Indeed, only a small fraction of the total cellular pool of CRLs is CAND1-associated, and phenotypes upon CAND1 gene mutation in C. elegans reflect deregulation of SCFLIN-23, but not other CRLs (Bennett et al., 2010; Bosu et al., 2010; Chua et al., 2011). Similarly, GLMN binds only ~7 and 0.2 percent of cellular RBX1 and CUL1, respectively (Fig. S7), and GLMN’s known biological effects seem particularly directed toward SCFFBW7, whose levels are decreased in a CRL- and proteasome dependent manner in the absence of GLMN (Tron et al., 2012). At this point, the reasons why among F-box proteins FBW7 is particularly sensitive to loss of GLMN remain unknown. It is also possible that other F-box proteins are co-regulated by presently unidentified inhibitors that can compensate for loss of GLMN. Future studies will be required to expand our understanding of GLMN’s functions in vivo, whether other CRL substrates are also regulated by GLMN under various cellular settings, and how GLMN-mediated inhibition is overcome.
Although few other cellular inhibitors of E3 ligases have been described, there are many mechanisms maintaining E3s in an “off-state”. For example, a secondary substrate binding site in the RING E3, Ubr1p, is occluded until activation by peptide binding to a primary substrate binding site (Du et al., 2002). Also, both cIAP1 and c-CBL make intramolecular interactions that prevent dimerization and/or E2 binding by their RING domains, with c-CBL being activated by phosphorylation (Dou et al., 2012; Dueber et al., 2011; Kobashigawa et al., 2011; Lopez et al., 2011). Autoinhibition also regulates several HECT E3s, which function by a distinct catalytic mechanism from RING E3s. HECT E3s form a covalent, thioester-linked HECT~ubiquitin intermediate via a catalytic Cys from which Ub is transferred (reviewed in (Kee and Huibregtse, 2007)). Accessibility of the HECT domain active site Cys can be blocked by intra-E3 interactions with domains that mediate cellular localization or substrate binding for active forms of HECT ligases (Gallagher et al., 2006; Wiesner et al., 2007). Finally, in parkin, which is a member of the recently described RING-HECT hybrid class of E3 that contains both a RING domain for E2 binding and a catalytic Cys, an internal Ub-like domain binds near the parkin RING domain and prevents interaction with Ub (Chaugule et al., 2011; Wenzel et al., 2011). Thus, as with CRL autoinhibition that is overcome by neddylation, representative members of all three of the major known cellular E3 classes appear to normally exist in inhibited states, awaiting activation.
This wide-range of mechanisms mediating E3 autoinhibition suggests numerous potential means for inhibition of CRLs and other E3s by trans-acting cellular factors (Fig. 6). We anticipate that many other E3s will eventually be discovered to have modes of regulation that parallel GLMN binding to RBX1, with cellular proteins masking the E2 binding sites on other RING or HECT E3s (Fig. 6). Furthermore, it is an exciting possibility that GLMN may represent the tip of the iceberg in terms of E3 ligase inhibitors mutated in diseases. Recent reports of several small molecules that ultimately inhibit CRLs by diverse mechanisms - through blocking NEDD8 ligation, substrate binding, or CDC34 activity (Ceccarelli et al., 2011; Orlicky et al., 2010; Soucy et al., 2009) - inspire confidence that it may in the future be possible to therapeutically restrain CRL activities that are mis-activated due to mutation of cellular E3 inhibitors in diseases.
We obtained an initial electron density map from crystals of SeMet-labeled GLMN complexed with RBX1-CUL1CTD by MR-SAD after merging 9 different SAD datasets to obtain data nearly complete to 3.5 Å resolution. In addition to 11 of 14 SeMets in the GLMN sequence visible in the SAD map, data from 19 more site-specific SeMets guided structure building. CUL1’s helix 29 and WHB subdomain, which correspond to regions we previously found undergo rotations in CRLs Duda et al., 2008), were not observed in the electron density maps, and thus were deleted in the construct used for final structure refinement at 3 Å resolution. Details are in Supplemental Information.
Detailed experimental procedures are in Supplemental Information. Briefly, to measure binding by FRET, equilibrium dissociation constants were obtained from titrations monitoring YFP quenching of a donor CFP signal. Interactions were examined between a C-terminal YFP fusion to GLMN and CFP-RBX1 purified alone, and in complexes with unneddylated and neddylated CUL1CTD, and between C-terminal YFP fusions to CDC34 or CDC34TAIL and RBX1 or RBX1 ΔRING complexed with split- ‘n-coexpress CUL1 C-terminally fused to CFP. The FRET efficiency (E) was determined by taking the maximum fluorescence intensity for the donor (CFP-conjugate) at 475 nm in the presence (FDA) and absence (FD) of acceptor (GLMN- or CDC34- or CDC34TAIL-YFP fusion protein) and solving for E=1−(FDA/FD). Kd was estimated by fitting of the titration curves to a hyperbolic equation assuming a single-site binding model.
For Biacore assays, RBX1’s RING domain was injected over minimally biotinylated GLMN (or mutant) captured on a CM-4 chip coated with covalently immobilized streptavidin. Triplicate injections were made for each concentration of RBX1’s RING domain (or mutant), and the data were processed, double-referenced and analyzed with the software package Scrubber2 (version 2.0b, BioLogic Software). The kinetic rate constants and/or equilibrium dissociation constants were determined by fitting the data to a 1:1 (Langmuir) interaction model.
In enzyme assays, GLMN was incubated with RBX1-CUL1~NEDD8 for 10 minutes prior to mixing with other SCF components and initiating reactions. Polyubiquitination assays were adapted from those described previously (Duda et al., 2008; Jubelin et al., 2010), with either 5 μM of a biotinylated CyclinE phosphopeptide or 0.2 μM phospho-p27 (complexed with CyclinA-CDK2) as substrates, 200 nM neddylated SCFFBW7 or SCFSKP2-CKSHS1, and 500 nM CDC34 for all figures except 5E, where CDC34 concentrations range from 500 nM-6 μM. GLMN was 50 nM-1 μM for Fig. 2D, 200 or 600 nM (1x or 3x relative to SCF) for Fig. 5E, and 600 nM in all other polyubiquitiation assays. SCFSKP2-CKSHS1 polyubiquitination assays also contained 250 nM UbcH5B. In the competition assays, 600 nM GLMN was incubated for 10 min on ice with 200 (1x in Fig. 2E and F) or 600 (3x in Fig. 2E and F and Fig. 4E) nM of the indicated version of the isolated RBX1RING, prior to incubation with RBX1-CUL1~NEDD8.
For pulse-chase di-Ub chain synthesis assays, a charging reaction containing 10 μM CDC34, 20 μM fluorescein-ubiquitin K0 and 150 nM UBA1 was quenched by 5-fold dilution into buffer containing 50 mM EDTA. This was diluted 10-fold into chase mixes containing 500 μM unlabeled native ubiquitin (SIGMA), and where indicated 500 nM RBX1-CUL1~NEDD8 and 2 μM GLMN. CDC34’s ability to synthesize a di-Ub chain was monitored by detecting transfer of fluorescein-K0 Ub from CDC34 to the free Ub.
This work was supported by ALSAC, PHS grants 5P30CA021765, R01GM069530 (BAS), RO1CA93804, R01CA63113 and P01CA050661 (JAD), and fellowships from the Pew Charitable Trust and the International Human Frontier Science Program (AET). BAS is an Investigator of the Howard Hughes Medical Institute. We are grateful to DC Scott, DW Miller, DJ Miller Polson, N Brown, P Soule, CO Rock, JW Harper, and EJ Enemark for advice and/or reagents; I Kurinov and C Ralston for synchrotron assistance; P Rodrigues and R Cassell for peptide synthesis and purification; S Bozeman and J Bollinger for administrative/computational support; and R Clubb for providing the sortase clone. NECAT is supported by NIH NCRR RR-15301, APS by US DOE W-31-109-ENG-38, ALS by US DOE DE-AC02-05CH11231. SAXS at the SIBYLS beamline BL12.3.1 of ALS is supported in part by the US DOE Integrated Diffraction Analysis Technologies (IDAT).
Accession code: Coordinates and structural data have been deposited in the Protein Data Bank with accession code 4F52.
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