SENP

SENP1

SENP1 plays a key role in the hypoxic response by regulating HIF1α (hypoxia-inducible factor 1α) stability (Cheng et al., 2007) and mitochondrial biogenesis (Cai et al., 2012).

From: Advances in Protein Chemistry and Structural Biology, 2016

Related terms:

Ubiquitin

Protease

C-Terminus

Hypoxia-Inducible Factors

Apoptosis

SENP3

SENP2

Nested Gene

Sumoylation

Ubiquitination

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SENP1 and SENP2 Peptidases

Edward T.H. Yeh, Hong Dou, in Handbook of Proteolytic Enzymes (Third Edition), 2013

Preparation

SENP1 is widely distributed in mammalian tissues, such as testis, thymus, pancreas, spleen, liver, ovary and small intestine [29]; the tissue distribution of SENP2 is unknown. To prepare full-length proteins, SENP1 and SENP2 are first over-expressed in mammalian cells, and then isolated through the antibody-dependent affinity chromatograph [22]. In addition, bacterial strains, such as E. coli BL21 DE3, have been utilized to express the recombinant partial SENP1 and SENP2. The purification of these recombinant proteins is based on a combination of the affinity, gel filtration and ion-exchange chromatograph [6]. Commercial recombinant catalytic domains of SENP1, SENP2 are available.

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Hormones and Breast Cancer

Todd P. Knutson, Carol A. Lange, in Vitamins & Hormones, 2013

3.3.2 SENP

Like PIAS overexpression, SENP1 activity can also modulate SR SUMOylation. Despite the transcriptionally repressive effect of AR SUMOylation, transient overexpression of SENP1 augmented AR-mediated transactivation (Cheng, Wang, Wang, & Yeh, 2004). Paradoxically, transcriptional transactivation of the SUMO-deficient AR mutant was also augmented. This suggests that the effects of SENP1 overexpression were likely independent of AR deSUMOylation (Cheng et al., 2004). Interestingly, SENP1 expression levels are induced by ligand-dependent AR action and can be inhibited by treatment with AR antagonist, bicalutamide (Bawa-Khalfe, Cheng, Wang, & Yeh, 2007). Further, the authors identified an ARE in the core promoter sequences of SENP1, and ChIP assays at the SENP1 promoter confirmed AR recruitment (Bawa-Khalfe et al., 2007). Thus, it appears that AR drives the expression of SENP1, and SENP1 activity elevates AR transcriptional transactivation, however, not through direct AR deSUMOylation. Instead, other studies revealed that SENP1 deSUMOylates HDAC1, relieving its repressive effects on AR (further discussed below; Cheng et al., 2004).

Notably, SENP1 transiently overexpressed in breast cancer cells mediated PR deSUMOylation and increased PR transcriptional activity in both reporter transcription assays and at PR-target genes, as measured by increased HBEGF transcript levels (Abdel-Hafiz & Horwitz, 2012; Daniel et al., 2007). In reporter assays, SENP1 expression sensitizes WT PR to low concentrations of ligand, whereas SUMO-deficient PR transcriptional activity was not affected, indicating that SENP1 mediates PR hypersensitivity via deSUMOylation of the receptor (Abdel-Hafiz & Horwitz, 2012; Daniel et al., 2007). Catalytically inactive SENP1 expressed in breast cancer cells repressed PR-mediated gene expression through dominant negative action (Daniel et al., 2007). SENP2 did not affect PR-mediated transcriptional activity, possibly due to SENP2's more diffuse nuclear and cytoplasmic localization (Daniel et al., 2007).

Despite the variability of in vitro SENP overexpression studies, recent work has revealed that some prostate cancers express high levels of SENP1, which contributes to elevated AR-dependent gene expression (Bawa-Khalfe et al., 2010; Cheng et al., 2006). In the normal parental prostate epithelial cell line, RWPE1, endogenous levels of SENP1 mRNA expression levels are relatively low, whereas the transformed (RWPE2) and malignant (LNCaP) prostate cancer cells contain high levels of SENP1 mRNA expression (Bawa-Khalfe et al., 2007). Indeed, high levels of SENP1 expression in patient tumors were correlated with high Gleason scores and increased risk of prostate cancer recurrence (Wang et al., 2013). SENP1 action also contributes to tumor aggressiveness through the stabilization of HIF-1alpha (via its deSUMOylation), allowing the transcription factor to upregulate matrix metalloproteinases (e.g., MMP2, MMP9) that are important drivers of metastasis (Wang et al., 2013). These experiments revealed that SENP1-mediated deSUMOylation of important transcriptional mediators (e.g., HDAC1, PR, AR, HIF-1alpha) contributes to SR-driven malignancies.

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Ion Channels as Therapeutic Targets, Part A

Hongmei Wu, ... Yitao Qi, in Advances in Protein Chemistry and Structural Biology, 2016

2.2 Reversal of SUMOylation by SUMO-Specific Proteases

SUMO conjugation is a dynamic process that is usually short lived, and SUMOylated proteins are rapidly deconjugated by SUMO-specific proteases (SENPs) (Mukhopadhyay & Dasso, 2007; Yeh, 2009).The initial step of SUMOylation requires the inactive SUMO to be cleaved at the carboxy terminus by the hydrolase activity of SENP. SENP exhibits isopeptidase activity to cleave the isopeptide bond between the glycine residue of SUMO and the lysine side chain of substrate (Yeh et al., 2000). The catalytic activity is maintained within a highly conserved 200 amino acid region in the C-terminus of the proteases. This enzymatic activity of the SENP exposes two glycine residues and generates active SUMO. Studies have shown that SENPs are important determinants of SUMO modification status in cells, and the deconjugation by SENPs plays a crucial role in determining a protein's SUMOylation status and activity (Cheng, Kang, Zhang, & Yeh, 2007; Jiang et al., 2012; Kang et al., 2010; Van Nguyen et al., 2012; Xu et al., 2010).

There are six mammalian SENPs with different subcellular locations and substrate specificities, and they exhibit different endopeptidase activities (Best et al., 2002; Gong, Millas, Maul, & Yeh, 2000; Hang & Dasso, 2002; Kim et al., 2000; Nishida, Kaneko, Kitagawa, & Yasuda, 2001; Nishida, Tanaka, & Yasuda, 2000; Yeh et al., 2000). These SENPs can deconjugate mono-SUMOylated proteins or disassemble polymeric SUMO side chains. SENP1, SENP2, SENP3, and SENP5 are more closely related to the yeast Ulp1, whereas SENP6 and SENP7 are related to Ulp2 (Yeh, 2009). SENP1 and SENP2 can deSUMOylate cellular substrates modified by any of the three SUMO isoforms, while the remaining four SENPs are more efficient at deconjugating SUMO2 and SUMO3 than SUMO1. SENP1, SENP6, and SENP7 are distributed in the nucleoplasm. SENP2 is more compartmentalized in the nuclear pore complex and SENP3 and SENP5 in the nucleolus. In addition, SENP1 and SENP2 express nuclear export sequences that allow these enzymes to shuttle in and out of the nucleus. Differences in the subcellular localization of the SENPs contribute to the selectivity of deSUMOylation for specific cellular proteins. Six SENPs can be divided into three independent subfamilies based on their sequence homology, substrate specificity, and cellular localization: family 1, SENP1 and SENP2; family 2, SENP3 and SENP5; and family 3, SENP6 and SENP7. SENP1 (Cheng et al., 2007), SENP2 (Kang et al., 2010), and SENP6 (unpublished data) knockout mouse embryos do not survive to birth, suggesting that these SENPs are not redundant and must have substrate specificity during development.

SENP1 and SENP2 process both C-terminal hydrolysis and isopeptidase activity in mammalian cells (Mukhopadhyay & Dasso, 2007; Yeh, 2009). SENP1 is a nuclear protease that deconjugates a large number of SUMOylated proteins (Gong et al., 2000) and is more efficient at processing both SUMO1 and SUMO2 than SUMO3 (Shen, Tatham, et al., 2006). There are two residues at the C-terminal side of the cleavage site that make significant contribution to SUMO process by SENP1 (Shen, Tatham, et al., 2006) and SENP2 (Reverter & Lima, 2006). SENP1-(415–643) deconjugates SUMO1- and SUMO2-conjugated RanGAP1 in the same manner in vitro (Shen, Tatham, et al., 2006), but it can discriminate between SUMO1-conjugated RanGAP1 and Sp100 (Shen, Dong, Liu, Naismith, & Hay, 2006), suggesting that the structure of its target proteins also impacts its isopeptidase activity. SENP1 plays a key role in the hypoxic response by regulating HIF1α (hypoxia-inducible factor 1α) stability (Cheng et al., 2007) and mitochondrial biogenesis (Cai et al., 2012).

SENP2 has been reported to be tethered to the nuclear pore through binding to Nup153 nucleoporin (Hang & Dasso, 2002; Zhang, Saitoh, & Matunis, 2002). SENP2 is also localized in a yet undefined nuclear speckle that is distinct from the nuclear body (Best et al., 2002). Our laboratory has shown previously that SENP2 contains both nuclear import and export signals that can shuttle between the nucleus and cytoplasm (Itahana, Yeh, & Zhang, 2006). SENP2 deconjugates SUMO1-conjugated RanGAP1 more efficiently than SUMO2-conjugated RanGAP1 in vitro, in agreement with its higher hydrolytic activity in processing SUMO2 (Reverter & Lima, 2006). The isopeptidase activities of SENP1 and SENP2 seem to be related to the nature of enzyme's interface with SUMOs. A spliced isoform of mouse SENP2, called SuPr1, can alter the distribution of nuclear POD (promyelocytic leukemia protein oncogenic domain)-associated proteins, such as CBP (CREB-binding protein) and Daxx, and convert Sp3 to a strong activator with diffuse nuclear localization (Best et al., 2002; Ross, Best, Zon, & Gill, 2002). SENP2 is the key regulator of Pc2/CBX4 function, which is critical for embryonic heart development, through regulation of the protein's SUMOylation status and is involved in the binding of polycomb complex to H3K27me3 (Kang et al., 2010). SENP2 also regulates myostatin expression and myogenesis (Qi, Zuo, Yeh, & Cheng, 2014) and has been implicated in Kv7 potassium channel-related seizures and sudden death (Qi, Wang, et al., 2014).

Our laboratory has characterized SENP3 and SENP5 as members constituting a family of nucleolar SENPs with preference for SUMO2/3 (Gong & Yeh, 2006). SENP3 and SENP5 are more active in deconjugating SUMO2/3-conjugated targets than SUMO1-containing ones (Di Bacco et al., 2006; Gong & Yeh, 2006). SENP3 can shuttle from the nucleolus to the nucleus under mild oxidative stress (Huang et al., 2009). It also regulates the transcriptional activity of HIF1α via deSUMOylation of the coregulatory protein p300 (Huang et al., 2009). SENP5 processes the precursor of SUMO3 only when compared with SENP1 and SENP2 (Di Bacco et al., 2006; Gong & Yeh, 2006).

SENP6 and SENP7, which are members of Ulp2 family, have an additional insertion in the catalytic domain, are mainly localized in the nucleus (Shen, Geoffroy, Jaffray, & Hay, 2009), and show very low processing activity (Lima & Reverter, 2008; Mukhopadhyay et al., 2006). In yeast, Ulp2 deletion causes an accumulation of high-molecular-weight conjugated species of Smt3p. In mammalian cells, SENP6 and SENP7 show stronger activity on poly-SUMO2/3 chains than on single SUMO moieties and SUMO1 chains (Lima & Reverter, 2008; Mukhopadhyay et al., 2006). SENP6 regulation of RPA70 plays a critical role in the regulation of DNA repair through homologous recombination (Dou, Huang, Singh, Carpenter, & Yeh, 2010). The structure of SENP7 is unique among well-characterized Ulp family members such as SENP1, SENP2, and Ulp1. The deletion of a distinct SENP7 region, whose location appears to explain its specificity in interactions with an extended SUMO chain, had no effect on the activities of SENP7 (Lima & Reverter, 2008). Differential expression of SENP7 variants regulates epithelial–mesenchymal transition (Bawa-Khalfe et al., 2012).

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Ubiquitin and Ubiquitin-like Protein Modifiers

Paul P. Geurink, ... Huib Ovaa, in Methods in Enzymology, 2019

3.1 Probes based on a monoUb or Ubl recognition element

We first demonstrate how a typical ABP labeling experiment can be performed using a panel of Ub-Prg and Ubl-Prg probes in combination with their known proteases. The panel of ABPs consists of untagged constructs of human Ub, Nedd8, SUMO1, SUMO2, SUMO3, ISG15, and the C-terminal domain of ISG15 (see Table 1). The C-terminal glycine is replaced by propargylamine in all ABP reagents.

Table 1. Overview of Ub and Ubl ABPs used in this study

ProteinAbbreviationUniProt IDResiduesaMW (kDa)UbiquitinUbP0CG47b1-758.5Nedd8N8Q158431-758.5SUMO1S1P631651-9611.1SUMO2S2P619561-9210.6SUMO3S3P558541-9110.5ISG15 C-domainI15cP0516179–1568.9ISG15 (C78S)cI15P051611–15617.1

aThe C-terminal Gly residue is not included in this list.bUb is only listed as a polyubiquitin in UniProt; the UniProt ID refers to polyubiquitin-B (UBB).cThe C78S-mutation was introduced to solubilize the ISG15-protein.

The reaction of a DUB or Ub-like protease with an ABP can be confirmed by incubation of the enzyme with the ABP followed by SDS-PAGE analysis. Fig. 3 shows the image of a typical ABP labeling experiment in which the ABPs were incubated with three proteases known to act on them: The DUB UCH-L3 is known to process Ub and Nedd8 (Gan-Erdene et al., 2003), SUMO protease SENP1 is active on all three SUMO proteins (Gong, Millas, Maul, & Yeh, 2000), and deISGylase USP18 targets ISG15 (Malakhov et al., 2002). The reaction between the enzyme and an ABP becomes apparent from the shift of a protein band to a higher molecular weight equal to the total mass of enzyme plus ABP. Here, the UCH-L3 band around 25 kDa shifts to a ~ 35 kDa band with either the Ub- or Nedd8 ABP. Similarly, the SENP1 band shifts from ~ 26 to ~ 40 kDa when incubated with either of the SUMO probes. It is to note here that SUMO proteins run somewhat higher than what would be expected from their mass. Finally, the USP18 corresponding band shifts from ~ 38 to ~ 43 kDa and ~ 50 kDa upon incubation with the truncated ISG15c ABP or the full-length ISG15, ABP respectively. From this result it follows that USP18 does not require full-length ISG15 for proper binding but that the C-terminal domain alone is enough, which corroborates earlier published results (Basters et al., 2017).

Fig. 3. Profiling of proteases UCH-L3, SENP1, and USP18 against Ub(l)-Prg ABPs.

Upon closer examination of the gel image in Fig. 3, it can be seen that in most cases where the enzyme is incubated with the ABP, a small protein band remains at the molecular weight corresponding to the unbound enzyme. This indicates that not all enzyme reacted with the ABP and that most likely the enzyme is not 100% active. Quantification of the band intensities will give an estimate of the active fraction of the enzyme.

An experiment as shown in Fig. 3 can also be used to validate the properties of an ABP that was constructed and purified, by incubation of the ABP with its known protease target. An appropriately folded and active ABP will result in a proper reaction with its protease, which can be checked and quantified by SDS-PAGE analysis.

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Ulp2 SUMO Protease

Jennifer Gillies, ... Mark Hochstrasser, in Handbook of Proteolytic Enzymes (Third Edition), 2013

Distinguishing Features and Related Peptidases

Ulp2 belongs to the protease family C48, which is a novel class of cysteine proteases distinct from, for example, the ubiquitin-specific processing proteases (UBP) in family C19. The SUMO-deconjugating enzymes in family C48 share similarity specifically in the catalytic domain of ~200 amino acids. The two yeast SUMO proteases, Ulp1 and Ulp2, are ~27% identical in this region [2]. Six human SUMO proteases can be classified into two branches: SENP1, SENP2, SENP3 and SENP5 belong to the Ulp1-like branch, while SENP6 and SENP7 are in the Ulp2-like branch [11]. Ulp1 and some Ulp1 branch members not only hydrolyze the peptide bond following the diglycine motif in SUMO precursors, but also cleave the isopeptide bond between SUMO and the lysine side chains of substrates. Ulp2 and the Ulp2-like SUMO proteases from other organisms appear to be specialized for SUMO-chain disassembly in vivo. A third, much more divergent branch in the C48 family, which contains human SENP8, acts on Nedd8-precursor, and possibly on Nedd8-conjugated substrates rather than sumoylated proteins. Nedd8 is a distinct ubiquitin-like protein. C48 family members are present in all eukaryotic species examined. Distantly related family members, also in the CE clan, include certain cysteine proteases in viruses and bacteria. So far, no specific antiserum against Ulp2 has been reported.

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SENP6 and SENP7 Peptidases

David Reverter, in Handbook of Proteolytic Enzymes (Third Edition), 2013

Structural Chemistry

The only structural information of the SENP6 and SENP7 members is the crystal structure of the catalytic domain of SENP7 [9] (Figure 531.1). SENP7 crystal structure revealed its relationship to the C48 SENP/ULP peptidase family as well as to the other members of the clan CE of cysteine peptidases. Despite the similar secondary structure elements forming the ULP/SENP fold, unique features are revealed from the structure of SENP7: (1) The absence of an N-terminal α-helix found in the structures of ULP1, SENP1 and SENP2; (2) four amino acid insertions varying in length between secondary structure elements; and (3) unique or extended secondary structure elements in SENP7 in comparison to SENP1 or SENP2 [15–18].

Figure 531.1. Ribbon representation of the crystal structure of the catalytic domain of SENP7. Active site catalytic residues are depicted in stick representation at the center of the molecule, namely Cys926, His794, Asp873 and Gln920. The insertion elements (Loop-1, Loop-2 and Loop-3) are labeled. Loop-1 can be clearly observed in the electron density maps, whereas Loop-2 and Loop-3 are disordered in the structure. N-terminal and C-terminal ends of the structure are also labeled with Nt and Ct, respectively.

Despite these differences, the SENP7 crystal structure shows a regular C48 fold, including all the signatures required for the SUMO peptidase activity. These signatures include the conserved catalytic residues forming the cysteine peptidase active site, Cys926, His794, Asp873 and Gln920, as well as the residues shaping the tunnel for the correct interaction with the C-terminal di-glycine motif of SUMO, Trp773 and Phe709. The latter phenylalanine is a tryptophan in all the other members of the family; however, activity assays on the single point mutant (F709W) indicate that phenylalanine is equally capable to place correctly the SUMO C-terminal tail on the active site of SENP7 [9].

Activity assays revealed that two of the sequence insertions, Loop-2 and Loop-3, are not required for the proteolytic activity in all substrates tested for SENP7 and SENP6 ([9] and D. Reverter, unpublished results). Loop-2 and Loop-3 insertions are disordered in the crystal structure of SENP7. However, deletion of Loop-1, with a highly conserved amino acid sequence between SENP6 and SENP7, shows major proteolytic defects in the cleavage of all substrates tested ([9] and D. Reverter, unpublished results). Interestingly Loop-1 is structured in the crystal structure of SENP7 and forms an extended hairpin loop with a stretch of four proline residues between two β-strands. Preliminary data indicate a role of the Loop-1 insertion in the preferential peptidase activity of SENP6 and SENP7 toward the SUMO2/3 isoforms (D. Reverter, unpublished results).

There is no crystal structure of the complex between SUMO2 with either SENP6 or SENP7. Based on structural comparison with other complexes, a similar extended quilt-like interface is expected to occur in SENP6 and SENP7 in the complex with SUMO2/3 [15–18]. Such interface can be divided in two different regions: the C-terminal tail with the di-Gly motif of SUMO that interacts with the active site of the protease, and an extended face of SUMO establishing several electrostatic contacts with the protease. In the model based on the crystal structure of SENP7, the Loop-1 insertion is in close proximity to SUMO2 and they can presumably establish relevant contacts [9].

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Recent Advances in the Discovery of Deubiquitinating Enzyme Inhibitors

Mark Kemp, in Progress in Medicinal Chemistry, 2016

4.3.4 USP8

In addition to compound 18 (USP8 IC50 = 96 nM), Hybrigenics have described several related fused tri- and tetra-cyclic series as USP8 inhibitors for treating a wide range of diseases [83,96,97]. Interestingly, the oxime derivative 26 retained the sub-μM USP8 potency of 18 but was inactive (IC50 > 100 μM) against USP7. The oxime group is responsible for this selectivity, since the parent ketone 27 has low μM USP7 potency (Figure 17). Another sub-μM USP8 oxime 28 was profiled more widely and found to have an IC50 of more than 100 μM against USP5, USP7, UCHL1 and SENP1. Byan et al. reported efficacy after administering compound 26 by i.p. injection at 0.2 and 1 mg/kg for 5 days per week in a non-small cell lung cancer mouse xenograft model [98].

Figure 17. Structures of USP8 inhibitors from Hybrigenics.

In 2014, Kathman et al. described how they synthesised a 100-member library of amidomethyl methyl acrylates and screened them against four proteases using an MS assay [99]. They identified compound 29 as a binding hit (Figure 18) albeit extremely weak (30% labelling of USP8 at 100 μM). This is interesting since at least the Michael acceptor group, which may be acting as a covalent warhead, has some clinical precedent from other cysteine proteases. Rupintrivir (30), an inhibitor of rhinovirus 3C protease, was progressed to Phase II trials by Agouron (now Pfizer) [100]. In addition, GSK recently reported some Phase I data on another structurally related compound which is a covalent irreversible cathepsin C inhibitor GSK'660 (31) [101].

Figure 18. Structures of a USP8 binder and two inhibitors of non-DUB cysteine proteases which advanced to clinical trials.

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SENP3 and SENP5 Peptidases

Long-Sheng Lu, Edward T.H. Yeh, in Handbook of Proteolytic Enzymes (Third Edition), 2013

Structural Chemistry

Human SENP3 consists of 574 amino acids and is a 65 kD protein. Human SENP5 consists of 755 amino acids and is an 87 kD protein. For both proteins, the characteristic C48 peptidase domain (~160 amino acids) localizes to C-terminus. Based on protein sequence, they are 80% similar to each other and are less so (~60%) to SENP1 and SENP2. The catalytic domain contains a catalytic triad of Cys, His and Asp. Based on crystal structure of SENP2 catalytic domain, His and Asp acid line up on the opposite sides of the active cleft [9]. Cys532 of SENP3 and Cys713 of SENP5 are key nucleophiles attacking isopeptide bond between terminal Gly of SUMO and Lys of a substrate. Mutation of this Cys to Ala results in complete loss of isopeptidase activity. In addition to the catalytic domain, SENP3 and SENP5 share a ~70 homologous amino acid segment that precedes the catalytic domain and characterizes this SENP subfamily. The N-terminal region of SENP3 and SENP5 dictates their subcellular distribution. SENP3 is a chromatin associated protein. Amino acids 86–153 contain a highly acidic stretch and absence of this segment results in redistribution from nucleolus to nucleoplasm. Overexpressing the catalytic domain of SENP3 selectively depletes SUMO2/3 conjugates in mammalian cells [10]. Similar features hold true for SENP5. The first 168 amino acids are required for its nucleolar localization [3]. The ectopically expressed catalytic domain of SENP5 also localizes to the nucleoplasm and depletes SUMO3 conjugates. A unique structural feature of SENP3 is a redox sensor between amino acids 163 to 320 [11]. Reactive oxygen species-mediated oxidation of cysteine thiols in this region blocks degradation of exogenous SENP3 in HEK293 cells in response to oxidative stress.

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Ubiquitin and Ubiquitin-like Protein Modifiers

Jonathan N. Pruneda, David Komander, in Methods in Enzymology, 2019

3.2.2 Visualizing DUB activity

As with any structural biology project, it is difficult to provide a certain recipe for success. Instead, we will discuss the techniques and examples that have proven successful in the past in order to provide a framework for what can be done with the tools available in the field. Well-defined substrate-bound DUB complexes can be studied with solution and crystallographic structural methods. Depending on size and behavior, NMR can be a useful tool for understanding DUB dynamics and regulation in solution, as shown for the DUBs AMSH and Cezanne (Hologne et al., 2016; Mevissen et al., 2016) as well as the UBL protease SENP1 (Ambaye, Chen, Khanna, Li, & Chen, 2018). Hydrogen-deuterium exchange mass spectrometry can also be used to monitor conformational changes associated with substrate recognition (Gersch et al., 2017; Mevissen et al., 2016). Finally, single-molecule techniques can inform on more global structural parameters, such as polyUb chain conformation following DUB binding (Ye et al., 2012). Crystallography and potentially cryo-electron microscopy can provide the highest-resolution information on substrate recognition and combined with covalent ABPs can offer the fastest route to understanding DUB specificity and mechanism.

Ub ABPs have enabled structural characterization and mechanistic understanding of multiple layers of DUB specificity. We have used monoUb/UBL ABPs extensively to characterize the role of the S1 site in substrate recognition. Cross-specific DUBs that possess Ub and UBL protease activities are particularly interesting cases, and ABPs have allowed us to explain the Ub/ISG15 cross-reactivity of the Crimean Congo hemorrhagic fever virus vOTU (Akutsu, Ye, Virdee, Chin, & Komander, 2011; James et al., 2011), as well as the Ub/tomato SUMO (tSUMO) cross-reactivity of Xanthomonas campestris XopD (Pruneda et al., 2016). In the case of XopD, we found that the S1 site is malleable, allowing it to recognize the structurally similar but sequence-divergent Ub and tSUMO substrates (Fig. 3A). The propargyl amide warhead was used for both substrates in this case, and provided a nice mimetic in the XopD active site (Fig. 3B). S1–S1′ diUb ABPs have also successfully trapped and allowed the crystallization of the Cezanne-K11 diUb complex (Mevissen et al., 2016) and the OTULIN-Met1 diUb complex (Weber et al., 2017). In the case of OTULIN, the noncovalent complex with the inactive Cys-to-Ala variant DUB had been crystallized previously (Keusekotten et al., 2013; Rivkin et al., 2013), and the diUb ABP could be confirmed as a suitable mimetic (Weber et al., 2017). The S1–S2 diUb ABP has also proven effective in the crystallization of the SARS coronavirus papain-like protease, explaining its di-distributive behavior of cleaving K48-linked polyUb (Békés et al., 2016).

Fig. 3. Visualizing DUB activity. Covalent complexes of the Ub (left) and tomato SUMO (tSUMO, right) carboxy-termini linked to the active site of X. campestris effector protein XopD. Ub and tSUMO ABPs were prepared from intein constructs (Borodovsky et al., 2002) with the propargyl amide warhead (Ekkebus et al., 2013). (A) Crystal structures of Ub (red) and tSUMO (yellow) bound in the XopD (green) S1 site, with the carboxy-termini threading into the active site. Conformational changes that accommodate the two substrates are highlighted. (B) Zoom-in of the XopD active site showing the full catalytic triad (Cys, His, Asp), the oxyanion hole (Gln), and the covalent linkage to substrate. 2 | Fo | − | Fc | electron density contoured at 1σ is shown for the relevant components of the active site.

Beyond polyUb chains, some DUBs preferentially recognize the most proximal, substrate-attached Ub linkage for hydrolysis. DUBs encoding this level of substrate- and site-specificity are likely few in number (as the number of ubiquitination sites outweighs the number of regulatory DUBs by ~ 500 fold) but critical for regulating fundamental cellular processes. Proteases responsible for regulating the UBL modifier SUMO must not only recognize polySUMO chains but also process the precursor SUMO translation product and remove SUMO from target proteins. The latter two processes have been captured in noncovalent complexes and crystallized to reveal the details of how both peptide- and isopeptide-linked SUMO are coordinated into the active site for hydrolysis (Reverter & Lima, 2006). The major role of the UBL modifier NEDD8 is in the regulation of cullin-RING Ub ligases, and this modification is in turn regulated by a dedicated ~ 350 kDa complex termed the COP9 signalosome (Lingaraju et al., 2014). While higher-resolution studies are eagerly awaited, low-resolution electron microscopy studies show large conformational changes associated with NEDDylated cullin binding to the COP9 signalosome, placing the NEDD8 carboxy-terminus near the catalytic subunit, CSN5 (Lingaraju et al., 2014). Lastly, a recent crystal structure has captured how a module from the SAGA transcriptional coactivator complex deubiquitinates monoubiquitinated histone H2B (Morgan et al., 2016). Each of these studies needed to overcome major obstacles in both enzyme and substrate preparation with the reported structures revealing remarkable insights into DUB mechanism and biology.

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Protein Kinases in Development and Disease

Jessica A. Blaquiere, Esther M. Verheyen, in Current Topics in Developmental Biology, 2017

4 Regulation of Hipk Activity

Peak activation of Hipk activity can be triggered by diverse morphogenetic signals, DNA damage, or cell stress. Unlike many mitogen-activated protein kinases (MAPKs), Hipk proteins seem to exist constitutively in a partially active state via autophosphorylation. Therefore, regulation of Hipk activity has evolved to focus on how to deactivate Hipk. Most notably, Hipk proteins are extensively PTM, and these modifications can be correlated with altered stability, localization, and consequently activity. In this next section, we briefly review these modes of regulation and how they affect the role of Hipk proteins in development and disease. For an excellent recent review on this topic, we also refer the reader to Saul and Schmitz (2013).

4.1 Phosphorylation

Hipks are dual specificity serine/threonine and tyrosine kinases. Hipk2 can autophosphorylate on conserved tyrosine residue in the kinase activation loop (Hipk2 Y354) (Saul et al., 2013; Siepi, Gatti, Camerini, Crescenzi, & Soddu, 2013). The activation loop is defined in earlier studies as the bold amino acids STYLQS [S352, Y354] (Rinaldo, Prodosmo, Mancini, et al., 2007; Rinaldo, Prodosmo, Siepi, et al., 2007), while later studies propose that phosphorylation of the activation loop occurs on STYLQS [Y354, S357] (Saul & Schmitz, 2013). Autophosphorylation is proposed to affect the degree of kinase activity and in turn the affinity for substrates, although a Y354F phospho-resistant mutant retains intermediate kinase activity (Saul et al., 2013; Siepi et al., 2013) and Y354 phosphorylation alone is not sufficient for full Hipk2 activity (Saul & Schmitz, 2013; Schmitz et al., 2014). Hipk3 also undergoes autophosphorylation (Rochat-Steiner et al., 2000). Hipk2 can also be modified by transphosphorylation at other sites outside of the kinase domain. It is likely that differential phosphorylation of Hipk2 provides a mechanism for controlling the signal output of the kinase. Of particular interest are Hipk2 phosphorylation sites that are not autophosphorylated, suggesting that other, as yet unknown, kinases are responsible for phosphorylating these sites. One such kinase may be Tak1 which is proposed to activate Hipk2 (Kanei-Ishii et al., 2004).

4.2 SUMO Modification

Hipk2 can be modified through the addition of the small ubiquitin-like modifier (SUMO) by SUMO1 (Kim et al., 1999), and SUMO modification of Hipk2 at K25 was shown to regulate its stability and activity (Gresko, Moller, Roscic, & Schmitz, 2005; Hofmann et al., 2005; Sung et al., 2005). Hipk2 recruitment to PML nuclear bodies requires the SIM of Hipk2 (Sung et al., 2011). Mutation of these residues results in localization of Hipk2 to the entire cell and reduced interaction with PML, although Hipk2 retained the ability to phosphorylate PML (de la Vega et al., 2011). Deletion of the SIM in Hipk2ΔSIM impaired the ability of Hipk2 to induce p53-dependent transcription, while enhancing its ability to phosphorylate its cytosolic substrate Siah2 (de la Vega et al., 2011). The K25 residue is conserved across vertebrates (human, Gallus gallus, Fugu rubripes, Xenopus) and can also be SUMO-modified in Hipk1 and Hipk3 (Gresko et al., 2005; Li et al., 2005). Removal of the SUMO moiety by the SUMO-specific protease SENP1 causes Hipk2 dissociation from PML bodies (Kim et al., 2005). dHipk is also SUMOylated, though the exact site is not known (Huang et al., 2011). In Drosophila tissues lacking SUMO (smt3 RNAi-expressing cells), JNK signaling becomes triggered, leading to ectopic apoptosis and compensatory proliferation. Loss of smt3 also causes exogenous dHipk to become delocalized from the nucleus, and enriched in the cytoplasm which allows it to stimulate JNK signaling (Huang et al., 2011). It is possible that the sumoylation status of dHipk can regulate its interaction with nuclear vs cytoplasmic partners. While some studies observed that the nuclear localization of Hipk2 and dHipk is dependent on its SUMOylation state (Huang et al., 2011; Kim et al., 1999), others found that adding or removing SUMO from Hipk2 does not influence its localization to PML nuclear bodies (Hofmann et al., 2005). Changes to the SUMOylation state of Hipk's can also alter the binding between Gro and Hipk2 (Sung et al., 2005).

4.3 Ubiquitin and Proteasomal Degradation

Hipk2 levels in unstressed cells are kept low through ubiquitin-targeted degradation mediated by proteins such as the E3-ubiquitin ligases Mdm2, Siah-1, Siah-2, Fbx3, and WSB-1 (Calzado, de la Vega, Möller, Bowtell, & Schmitz, 2009; Calzado, De La Vega, Muñoz, & Schmitz, 2009; Choi et al., 2008; Rinaldo, Prodosmo, Mancini, et al., 2007; Rinaldo, Prodosmo, Siepi, et al., 2007; Shima et al., 2008; Winter et al., 2008). The role of Mdm2 has already been described above in the p53 signaling section. Following treatments that induce severe DNA damage, Hipk2 degradation induced by Siah1 and WSB-1 ceases, resulting in elevated Hipk2 levels. In contrast, mild genotoxic stress does not block Siah-1, and Hipk2 is rapidly degraded. The p53-inducible ligase Siah-1L targets Hipk2 and Hipk3 for degradation, thus preventing p53 activation and apoptosis (Calzado, de la Vega, Möller, et al., 2009; Calzado, De La Vega, Muñoz, et al., 2009). Zyxin promotes Hipk2 stability by interfering with Siah-1 function (Crone et al., 2011). The XIAP-associated factor 1 (XAF1) can also block Hipk2 degradation by interfering with the Siah2–Hipk2 interaction (Lee et al., 2014), while hypoxia can trigger the interaction between Hipk and Siah2 (Calzado, de la Vega, Möller, et al., 2009; Calzado, De La Vega, Muñoz, et al., 2009).

4.4 Acetylation

Hipks 1–3 can be acetylated on lysines in the presence of CBP, although this occurs at very low levels in normal cells (Aikawa et al., 2006; de la Vega et al., 2012; Hofmann et al., 2002). Acetylation causes delocalization of Hipk2 from speckles to the nucleoplasm and cytoplasm (de la Vega et al., 2012). Mutation of Hipk2 lysine residues does not affect its ability to phosphorylate its substrate Siah-2, suggesting that acetylation may not affect all cellular functions of Hipk2. The ability of Hipk2 to induce cell death is regulated by acetylation and levels of ROS (de la Vega et al., 2012). Inhibition of acetylation through mutation of lysine residues causes Hipk2 to have enhanced cell killing properties, suggesting acetylation normally protects against such death. SUMOylation of Hipk2 promotes its deacetylation by HDAC3 and favors cell survival. Under physiological conditions, acetylation provides a protective effect by delocalizing Hipk2 from sites where it interacts with p53, and could trigger cell death.

4.5 Regulation of Hipk Gene Expression

As described above, Hipk protein levels are tightly regulated by numerous PTMs that affect stability, localization, and function. Hipk gene expression levels are also controlled through microRNA-mediated posttranscriptional regulation (reviewed in Conte & Pierantoni, 2015). For example, in renal tubular epithelial cells, miR-141 downregulates expression of Hipk2 (Huang et al., 2015). Currently, the transcriptional regulation of Hipk family genes is an understudied area, which may yet reveal additional modes of ensuring adequate Hipk function.

4.6 Drug-Based Inhibition of Hipk

While numerous generic protein kinase inhibitors can influence Hipk, only a few have been described to be specific to Hipk. The drug D-115893 has dramatic effects causing Hipk delocalization and distribution throughout the cell, in addition to blocking p53 serine 46 phosphorylation (de la Vega et al., 2011). Recently a very specific Hipk2 inhibitor named TBID was identified that can block phosphorylation of both p53 and a generic substrate by endogenous Hipk2 in human T-lymphoblastoid cells (Cozza et al., 2014). TBID is also capable of inhibiting Hipk1 and Hipk3 at a lower efficiency.

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