Structure and function of tripeptidyl peptidase II, a giant cytosolic protease

Biochimica et Biophysica Acta 1824 (2012) 237–245

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Review

Structure and function of tripeptidyl peptidase II, a giant cytosolic protease☆ Beate Rockel a,⁎, Klaus O. Kopec b, Andrei N. Lupas b, Wolfgang Baumeister a a b

Department of Molecular Structural Biology, Max Planck Institute of Biochemistry, Am Klopferspitz 18, D-82152 Martinsried, Germany Department I — Protein Evolution, Max Planck Institute for Developmental Biology, Spemannstr. 35, D-72076 Tübingen, Germany

a r t i c l e

i n f o

Article history: Received 28 April 2011 Received in revised form 29 June 2011 Accepted 1 July 2011 Available online 13 July 2011 Keywords: Tripeptidyl peptidase II Cytosolic proteolysis Hybrid structure Protein evolution

a b s t r a c t Tripeptidyl peptidase II is the largest known eukaryotic peptidase. It has been described as a multi-purpose peptidase, which, in addition to its house-keeping function in intracellular protein degradation, plays a role in several vital cellular processes such as antigen processing, apoptosis, or cell division, and is involved in diseases like muscle wasting, obesity, and in cancer. Biochemical studies and bioinformatics have identified TPPII as a subtilase, but its structure is very unusual: it forms a large homooligomeric complex (6 MDa) with a spindle-like shape. Recently, the high-resolution structure of TPPII homodimers (300 kDa) was solved and a hybrid structure of the holocomplex built of 20 dimers was obtained by docking it into the EM-density. Here, we summarize our current knowledge about TPPII with a focus on structural aspects. This article is part of a Special Issue entitled: Proteolysis 50 years after the discovery of lysosome. © 2011 Elsevier B.V. All rights reserved.

1. Introduction The ubiquitin–proteasome system constitutes the main pathway for protein degradation in eukaryotic cells [1]. Its most downstream element, the 26S proteasome, has been studied in great detail even though a high resolution structure of the holocomplex is still not available. The structure of its proteolytic core complex, the 20S proteasome, has long been solved and in conjunction with mutagenesis has clarified the role of 14 α- and 14 β-subunits in protein breakdown [2]. In recent years, the group of giant post-proteasomal proteases and especially tripeptidyl peptidase II (TPPII) have also come into focus [3]. The proposed cellular role of the latter is in cytosolic protein degradation downstream of the proteasome in conjunction with other exo- and endopeptidases [4]. Under conditions where the function of the proteasome is compromised, e. g. by inhibitors, but also in certain diseases, TPPII is upregulated and a number of studies have been performed to reveal its role in health and disease states (see [5,6] for recent reviews). TPPII has been described as a ‘multi-purpose peptidase’ [7], and indeed, many functions have been ascribed to it, but in many cases its substrates or reaction partners have remained obscure. Bioinformatic and biochemical studies had suggested early on that the N-terminal part of TPPII is homologous to subtilisin [8] but the function of the larger part of the polypeptide chain remained enigmatic. Based on its similarity to subtilisin, homology models of the ☆ This article is part of a Special Issue entitled: Proteolysis 50 years after the discovery of lysosome. ⁎ Corresponding author at: Max Planck Institute of Biochemistry, Department of Molecular Structural Biology, Am Klopferspitz 18, D-82152 Martinsried, Germany. Tel.: + 49 89 8578 2698; fax: + 49 89 8578 2641. E-mail addresses: [email protected] (B. Rockel), [email protected] (K.O. Kopec), [email protected] (A.N. Lupas), [email protected] (W. Baumeister). 1570-9639/$ – see front matter © 2011 Elsevier B.V. All rights reserved. doi:10.1016/j.bbapap.2011.07.002

active site region of TPPII have been created [9,10] and sequence comparisons have been used to pinpoint potential functional regions [11]. New opportunities for the functional analysis of TPPII have been opened by the recent determination of a high resolution structure of TPPII using a hybrid EM-X-ray crystallography approach, where the crystal structure of TPPII dimers was docked into the structure of the TPPII holocomplex obtained by cryo-electron microscopy [12]. In this review we will summarize our current knowledge about this giant protease. 2. Cellular functions of TPPII 2.1. TPPII in cytosolic proteolysis TPPII was discovered in 1983 in the extralysosomal fraction of rat liver during a search for peptidases with specificity to proteins phosphorylated by cyclic AMP-dependent protein kinase [13]. Subsequently it was found in many other tissues and also in red blood cells [14]. Its function – the removal of a tripeptide from the free N-terminus of longer peptides – had up to then only been observed for TPPI, a structurally unrelated lysosomal peptidase. In addition to exopeptidase activity, endopeptidase activity has also been ascribed to TPPII, but this activity is much lower than its exopeptidase activity [15,16]. So far only unfolded peptides have been reported to be cleaved by TPPII, the longest one, with a length of 41-residues, being Ova37–77 [15]. Based on the type of substrates degraded and in analogy to Tricorn protease [17], TPPII was assigned a role downstream of the proteasome in cellular protein degradation [4]; however, direct experimental evidence for this disassembly line is still lacking. In fact, the processing of proteasomal products is something TPPII has in common with other peptidases like leucine aminopeptidase LAP [18], thimet oligopeptidase TOP [19], bleomycin hydrolase BH [20,21], or

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puromycin-sensitive aminopeptidase PSA [21,22]. Nonetheless, TPPII appears to be the only peptidase capable of degrading peptides that are longer than 15 residues [23,24]. Among the cellular functions attributed to TPPII, its function as a neuropeptidase inactivating the satiety hormone CCK8 is probably characterized best [25]. The cleavage of CCK8 to CCK5 in rat brain is carried out by a membrane-associated version of TPPII, which was suggested to be anchored in the lipid bilayer by a covalent glycosyl phosphatidyl inositol link [25]. Its influence on satiety has made TPPII an interesting target for obesity-treatment and, indeed, TPPII-inhibition by treatment with the specific inhibitor butabindide was shown to reduce the food intake in rats [25]. A role in fat metabolism was also proposed for C. elegans TPPII, which surprisingly appeared to be independent of the presence of a functional proteolytic domain [26]. TPPII has been implicated in antigen processing, however the extent to which it is essential for the processing of antigenic peptides has remained controversial (for reviews see [5,6,27]). TPPII was reported to be involved in the generation of two viral epitopes [16,28], although a proteasome with an altered specificity could also be responsible for the creation of these MHC class-I peptides [29]. Altogether, the generation of most MHC class I-bound peptides appears to be independent of TPPII [27]. Nevertheless, the processing of peptides longer than 15 residues requires TPPII [23,24], but only a small fraction of the peptides released by the proteasome falls into that size range [24,30,31].

2.2. TPPII and its role in diseases TPPII-activity is increased in skeletal muscle during sepsis-induced muscle wasting [32] as well as during cancer cachexia [33]. Responsible for the accelerated proteolysis under such catabolic conditions is the ubiquitin proteasome-system [34,35], supporting the notion that TPPII functions downstream of the proteasome and is co-induced with it. TPPII is also upregulated in tumor cells, a finding that might have an impact on cancer therapy: EL4 thymoma or EL4 lymphoma cells adapted to proteasome-inhibition as well as Burkitt's lymphoma cells, where the proteasome appears to be functionally impaired, show increased TPPII activity [15,36,37]. From such observations it was concluded that TPPII may allow survival of these cells by compensating for a loss of proteasome function [15,36]. Indeed, it was shown that in Burkitt's lymphoma cells, protein turnover is unaffected and that ubiquitinated proteins do not accumulate [37] unless the cells are treated with the covalent serine protease inhibitor AAF-CMK, which inhibits TPPII [37,38]. However, AAF-CMK is not specific for TPPII, since it affects also other proteases like the proteasome [39]. In the presence of the specific TPPII inhibitor butabindide or siRNA against TPPII no such accumulation occurs, implying that TPPII cannot substitute for the proteasome in the cleavage of ubiquitinated proteins [38]. Burkitt's lymphoma cells are apoptosis-resistant, but apoptosis can be induced by TPPII-inhibition with AAF-CMK [37]. For EL-4 lymphoma cells adapted to proteasome-inhibition apoptosis-resistance and increased growth-rate was ascribed to an impaired degradation of inhibitors of apoptosis (IAP) and both features could be induced by TPPII-upregulation after TPPII-transfection [40]. HEK293 cells are yet another cell line for which apoptosis-resistance and accelerated growth upon overexpression of TPPII were shown. Such TPPII-overexpressing HEK293-cells could survive the effect of the spindle poison nocodazole and showed a higher degree of aneuploidy as well as more structural and numerical centrosome abnormalities than control cells [41,42]. Also the cell-division errors observed in Burkitt's lymphoma cells seem to depend on TPPII, since the observed c-MYC induced centriole overduplication can be avoided by TPPII inhibitors like butabindide or by siRNAmediated protein knock-down [43]. A participation of TPPII in cell division, as suggested by these experiments, might be the reason for its observed localization in the vicinity of daughter centrioles in late mitosis and between daughter and mother centrioles during G2 phase [43].

In several malignant cell lines TPPII translocated into the nucleus upon γ-irradiation and the production of reactive oxygen species (ROS), which suggested a role for TPPII in DNA-repair [44,45]. However, this translocation as well as the accumulation of p53 remains controversial, since they were not observed in EL4 cells, COS cells, and transformed fibroblasts [46,47], a discrepancy that was attributed to different levels of ROS and sub-optimal cell densities [44].

2.3. TPPII-deficient species In order to investigate the importance of TPPII for cell survival, a number of TPPII-deficient species have been created. A T-DNA mutant of Arabidopsis defective in TPPII expression showed no phenotypic abnormalities [48] and likewise, a TPPII-knockout strain of S. pombe was viable and did not have any obvious growth defects [11,49]. Suppressing TPPII expression by siRNA in C. elegans resulted in decreased fat stores in adult worms; however, reduced CCK8degradation was not detectable and therefore no connection to satiety control could be established [26]. Divergent observations were reported for TPPII-deficient mice: McKay et al. [26] failed to obtain homozygotic TPPII-deficient mice due to early embryonic lethality. However, their tpp2 heterozygous mutants were lean compared with wild-type littermates, while their food intake was normal. Kawahara et al. [50] produced gene-trapped mice with an expression level of TPPII reduced by N90% compared to wild-type. These mice with a gene-trap disrupting tpp2 were viable, fertile, and normal in appearance and behavior. In contrast, Huai et al. [51] describe knockout mice homozygotic for tpp2−/−, which were viable but in which the TPPII-deficiency activated cell-type specific death programs. As a consequence the mice had a decreased life-span. Also, how TPPIIdeficiency affects Drosophila is not clear. In a screen of lethal mutants on the second chromosome of D. melanogaster for those that could enhance a weak Ras1 eggshell phenotype, one insertion disrupted two genes, Nrk, a neurospecific receptor tyrosine kinase and TPPII. Whether the lethality is attributable to either of the two disrupted genes alone or to the additive effect of both remains unclear [52].

Fig. 1. 3D structure of the TPPII holocomplex. A) 3D-reconstruction of DmTPPII, segment numbers are indicated for one strand. B) DmTPPII rotated about 90° around the longitudinal axis. Dimers in the strand on the left are highlighted in orange and red to visualize their stacking; the strand on the right was cut open in order to show the internal cavity system of TPPII.

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3. TPPII structure

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‘double-clamp’, a structural feature allowing reciprocal interactions between dimers positioned at the ends of the two strands [55].

3.1. Quaternary structure of TPPII 3.2. Crystal structure of TPPII dimers The first electron micrographs of negatively stained TPPII particles isolated from human erythrocytes were already published in 1996 and showed that the subunits assemble into a large oligomeric structure [53]. Also Drosophila TPPII (DmTPPII) is a 6 MDa complex with a spindle-like shape. It is composed of two 60 nm long twisted strands build of 10 stacked dimers, where each strand encloses a central cavity system that is accessible through lateral openings [54,55] (Fig. 1). TPPII dimers possess only 10% of the specific activity of the spindle [56,57] but upon assembly their specific activity increases with each newly formed interface, suggesting a contact-induced activation mechanism [57]. Compared to long, single strands the spindles are thermodynamically stabilized. This is accomplished by a

The concentration-dependent size of TPPII oligomers induces a polymorphism, which is not conducive to crystallization [57]. However, TPPII spindles can be dissociated by cold-treatment and reassembly of the dimers can be prevented by the presence of detergents such as octyl glucoside, which made it possible to obtain crystals [12]. TPPII monomers are 128–150 kDa in size, dependent on the species. The 150-kDa monomer of DmTPPII can be divided into three basic domains (Fig. 2A): The N-terminal domain represents the subtilisin domain, which in TPPII is interrupted by a long insertion between the catalytic D44 and H272 residues. The central domain is mainly composed of β-strands. Together, these two domains form a ring-structure with a central hole, which

Fig. 2. High resolution structure of DmTPPII. A) Domain composition of DmTPPII and comparison with subtilisin. Upper bar: Yellow: subtilisin-like domain of TPPII (residues 1–522), active sites D44, H272, S462 are shown as red asterisks; orange: insertion within the active site (residues 75–266); green: central domain (residues 523–1098); blue: C-terminal domain (residues 1099–1354); gray blocks: Loops that are not present in the crystal structure of DmTPPII (PDB ID: 3LXU). Lower bar: yellow: subtilisin Carlsberg aligned to dmtppII, active sites D32, H64, S221 are shown as red asterisks. B) DmTPPII monomer shown in ribbon representation. Domain colors are as described in A). C) DmTPPII dimer shown in two orientations. Domain colors are as described above, one of the monomers is shown in ribbon — the other in surface representation. D) Overlay of the high resolution structure of TPPII (yellow) with subtilisin Carlsberg (gray) (PDB ID: 1CSE). Active site residues D44, H272, S462, as well as the helix connected to S462 and loop L2 of TPPII are shown in red, active site residues D32, H64, S221 and the helix connected to S221 and loop “L” of subtilisin are shown in blue. E, F) Conformation of the helix connected to the active site serine in subtilisin (E) and TPPII (F). G) Active-site region of TPPII showing the loop L2 residues bound to the active site as well as the location of the double-Glu motif (E312, E343). The N-terminal continuation of L2 is indicated by a dotted red line next to L457 (P3).

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contains the active site on one side (Fig. 2B). The insertion connects the ring-structure with the C-terminal domain, which is mainly α-helical. In a TPPII-dimer, two monomers are arranged in a C2-symmetry related position (Fig. 2C) and through this arrangement the holes in the centers of the rings are covered reciprocally and the C-terminal domains assemble into a handle-like structure. The active site residues D44 and H272 of DmTPPII are in a conformation very similar to that of the corresponding residues D32 and H64 of subtilisin Carlsberg, but in contrast to S221 in subtilisin, S462 of TPPII is pointing away from the other two residues of the catalytic triad (Fig. 2D). This difference in orientation is correlated with a difference in the position of the two serine residues: S221 in subtilisin is located at the N-terminal end of an α-helix. S462 in TPPII is also close to the N-terminal end of a helix but – unlike S221 in subtilisin – it is not part of the helix (Fig. 2E and F). At the N-terminal side of S221 in subtilisin there is a stretch comprising ~20 residues that form a rather rigid structure composed of two antiparallel β-sheets (loop “L” in Fig. 2D). In contrast, most of the 20 residues at the N-terminal end of S462 in TPPII belong to the flexible loop L2 with no defined density in the crystal structure (Fig. 2D); only residues L457-N459 are visible; they occupy the substrate-binding cleft of TPPII (Fig. 2D and G). The combination of a displaced active-site serine residue and internal loop-residues bound to the substrate-binding cleft is believed to be the reason for the observed low activity of TPPII dimers [56,57]. The main activity of TPPII is that of a tripeptidyl peptidase. The structural realization of the molecular ruler ensuring the separation of exactly a tripeptide is a ‘double-Glu’ motif, which had been ascribed to TPPII already earlier [10] and which is also found in prolyl tripeptidyl peptidase [58], dipeptidyl peptidase IV [59], tricorn-interacting aminopeptidase F1 factor [60], and aminopeptidase N [61]. In TPPII, the two glutamate residues E312 and E343 of the double-Glu motif form a ridge that prevents the binding of a fourth residue to the substrate-binding site thus limiting access to three residues only. At the same time they interact with the positively charged N-terminus of the substrate. Since TPPII can also act as an endopeptidase, longer peptides must be able to bind to the peptide binding cleft as well. The conformation of the three residues of the internal loop L2 that are bound to the substrate-binding cleft in the crystal structure may indicate how an endopeptidase substrate can be accommodated: the P3-residue L457 does not possess a free N-terminus and cannot built a

salt-bridge with the double-Glu motif, the P4 residue cannot bind to the blocked S4 position and as a consequence P3 would have to bend out of the cleft (Fig. 2G; [12]). 3.3. Hybrid structure of the TPPII spindle The hybrid structure of TPPII revealed the location of the domains of TPPII within the spindle architecture of the complex (Fig. 3A) and showed that the N-terminal and central domains are located at the inner, concave side of the spindle, whereas the C-terminal domains constitute the outer, convex side [12]. Through the stacking of the dimers into strands, the active sites, which are located on the surface in dimers (see Fig. 2), are sequestered inside a large cavity system traversing the strands. Substrates must enter through an opening at the handles into the foyer and proceed through the antechamber to reach either of the two catalytic chambers sandwiched between each two dimers (Fig. 3B and [12]). Constrictions at entry and exit sites of the antechambers are approx. 30 Å by 15 Å restricting access to unfolded polypeptides. In addition to compartmentalizing the active sites, the stacking of dimers into strands also induces activation, during which the S462 is moved to its catalytically active position and the L2 residues bound to the substrate-binding cleft are displaced. Most likely, the interaction of the flexible loop L2 with the neighboring dimer triggers this rearrangement (Fig. 3C and [12]). 4. Functional regions in TPPII Whereas the N-terminal third of the TPPII sequence had been identified as its proteolytic domain long time ago [62], the role of its Cterminal two-thirds remained enigmatic. From the hybrid structure of TPPII it is obvious that a large portion of the polypeptide chain is involved in the formation of the unique cavity system of the complex (see Fig. 2C). Whether parts of the sequence are also involved in other functions like substrate-binding or interaction with co-factors, is currently unknown. For some parts of the sequences there is structural and functional information based on sequence analysis, mutations and biochemical evidence [11] (Fig. 4). The residues of the active site [62,63] and the double-Glu motif [10,12] have been confirmed by biochemical studies and site-directed mutagenesis. The

Fig. 3. Hybrid structure of the TPPII spindle. A) Crystal structure of the DmTPPII-dimer docked into the EM-map of the DmTPPII spindle (mesh representation). TPPII dimers are shown in ribbon representation. The color code corresponds to Fig. 2A. B) Schematic drawing of the cavity system that is created through the stacking of the dimers into strands. Red dots: Locations of the active sites. H: Handles, CC: Catalytic chamber, AC: Antechamber, F: Foyer. Arrows label the entrances into the cavity system at either site of the handles of a dimer. C) Activation-scheme of TPPII: The stacking of the dimers leads to the formation of the cavity system, the reorientation of loop L2 and also to the correct placement of the active-site serine residue S462.

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insertion within the catalytic site, which has been reported to be involved in complex formation [64], is indeed located at the dimer– dimer interface in spindles. A special role in complex formation was attributed to G252, a residue within the insertion, since the mutation G252R prevented the assembly of dimers into spindles [64], even though at high protein concentrations assembly does proceed [57]. A role in the assembly-dependent activation of DmTPPII has been proposed for residues L457-N459 and L603-R610. Residues L457N459, which are connected to a flexible loop, are bound to the substrate-binding cleft in dimers [12]. It has been hypothesized that during assembly they are removed from the cleft due to the interaction of the flexible loop with residues L603-R610 of the adjacent dimer [12]. The region around K1219 appears to be conserved in arthropods [11] and the corresponding region in HsTPPII contains the first 18 residues of a 20 kDa fragment, which can be produced by chymotryptic cleavage of HsTPPII [65]. When the cleavage is carried out in the presence of CaEGTA, the 20 kDa fragment is not observed; instead, a 30 kDa fragment is produced,

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which corresponds to residues 939–956 in HsTPPII. This suggests that the corresponding regions can undergo conformational changes [8,11], which is quite interesting, since they reside in different domains of the complex. The region corresponding to residues 1137– 1157 in HsTPPII was described as hypervariable in mammals [11,66] and in the crystal structure of DmTPPII dimers it represents a loop region connecting two helices of the C-terminal domain. In HEK293 cells the cDNA of a potential splicing variant of TPPII has been identified and expressed. The additional 13 residues located between residues 983 and 984 in HsTPPII seem to lead to the formation of a larger TPPII complex or to aggregation [66,67]. In DmTPPII the corresponding residues are located at the dimer–dimer contact region, where it is quite likely that they influence complex formation. 5. Evolution of TPPII Although TPPII is thought to be specific to eukaryotes, it is clearly a member of the subtilisin family and thus homologous to many

Fig. 4. Functional regions in TPPII. Functional regions in the TPPII sequence (amino acid sequence of the expressed Drosophila TPPII, http://uniprot.org/uniprot/Q9V6K1 Isoform 2, or Flybase ID FBpp0086888) are highlighted (and numbered) in both the sequence (A) and in the crystal structure (B–C) of DmTPPII. The underlined residues belong to regions that are missing in the crystal structure: loops L1, L2 and L3, as well as 17 residues at the beginning of the sequence and a stretch of 14 residues in the insertion between D44 and H272. B) Highresolution structure of DmTPPII-monomers in ribbon-representation, the locations of the functional regions highlighted in A) are indicated. C) High-resolution structure of DmTPPIImonomers in surface representation in two orientations. Red (1): active site residues (D44, H272, and S462) and double-Glu motif (E312, E343) [12], Crème (2): insertion within active site residues D44 and H272, residues Y69-L263, corresponding to residues 68–255 in HsTPPII (DH-insert) [64]. Magenta: G260 (3), corresponding to G252 in HsTPPII, which was described to be critical for complex formation [64]. Orange (4): Residues L457-N459 are bound to the active site in the crystal structure of DmTPPII [12]. Olive (5): Residues L603-R610 supposedly involved in the activation of DmTPPII [12]. Bright green (7): R1012-V1013, in a splicing variant of TPPII, 13 residues are inserted between the corresponding residues (HsTPPII 983–984) in chordate and are affecting complex formation [66,67]. Dark blue (8): K1219-N1238, corresponding region in HsTPPII represents the first 18 residues of a 20 kDa fragment, which can be produced if TPPII is cleaved by chymotrypsin in the absence of CaEGTA [11,65]. Dark green (6): S967-T973, corresponding region in HsTPPII 939–946 is produced instead of K1219-N1238 after chymotrypsin cleavage of TPPII in the presence of CaEGTA [8,11], Cyan (9): L1257-K1262, corresponding region in HsTPPII (1137–1157) has been described as hypervariable in mammals [11,66]. For all functional regions – except the DH-insert – residues are shown in stick representation for better visibility.

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prokaryotic proteins. To explore this relationship, we performed a bioinformatic analysis using structural (DALI [68]) and sequence comparisons (HHpred [69]), as well as clustering (CLANS [70]) methods (Fig. 5). As a first step, we compared the sequences of all subtilisins in the non-redundant database and clustered them by the significance of their pairwise matches. The resulting map (Fig. 5A and B) shows a compact cluster for eukaryotic TPPII proteins (green in Fig. 5A and B), with two embedded bacterial proteases that presumably originated by lateral transfer (pink): a protein from Blastopirellula marina, annotated incorrectly as pyrolysin but previously reported by Eriksson et al. as a TPPII homolog [11], and a protein from Planctomyces brasiliensis. Closely connected to eukaryotic TPPII are a cluster of actinobacterial proteases (yellow) and the pyrolysins of euryarchaea (red) – which can thus be considered the bacterial and archaeal orthologs of TPPII, respectively – as well as a few peripheral sequences from environmental microorganisms (Symbiobacterium, Thermincola, Planctomyces, Chloroherpeton). The STABLE protease of crenarchaea (stalk-associated archaebacterial endoprotease [71]) is a divergent satellite cluster to TPPII, branching off from the pyrolysins. The TPPII group is connected to the main subtilisin groups via a large cluster of sequences found mainly in actinobacteria (blue in Fig. 5A and C), which we will refer to in the following as the anchor group (the second group of eukaryotic proteases originating from this anchor group – light blue in Fig. 5A and C – is membrane-bound transcription factor site-1 protease). The broad range of organisms containing TPPII-like proteins suggests that a primitive form of TPPII was already present in the last universal common ancestor (LUCA). In order to gain an understanding of the domain structure of this early form of TPPII and its evolution to the giant, multi-domain protease found in eukaryotes, we performed a domain analysis of the groups of TPPII relatives identified by clustering (colored clusters in Fig. 5A). TPPII was previously described by sequence analysis as composed of three parts: (i) the subtilisin domain, including the insertion; (ii) a group of domains consisting of β-sheets; and (iii) a helical C-terminal part [12]. For the present analysis, we used a more precise, structure-derived division of TPPII into eight domains, which we named A–H. The domains are arranged along the sequence from N- to C-terminus as A′–B′–C–B″–A″–D–E–F′–G–F″–H (Fig. 5D and E). In case of domain insertions, letters with single and double primes indicate the N- and C-terminal parts of one domain, respectively. This new partition offers a finer-grained view than the previously used one, but the two correspond directly: part (i) contains A, B, and C, part (ii) comprises the four β-sheet domains D, E, F, and G, and part (iii) is synonymous for H. The N-terminal domain A is the subtilisin-like domain and universally present. Within the catalytic triad of A, domain B is inserted and domain C is again inserted inside domain B, which yields the rather unusual telescopic arrangement A′–B′–C–B″–A″. Insert domain B adopts an unusual fold so far not captured in SCOP or CATH. Its closest structural relative appears to be another insert domain, which protrudes from the catalytic domain of oligosaccharyltransferase (OST), a multidomain enzyme that catalyzes the co-translational transfer of an oligosaccharide from a lipid donor to an asparagine residue in nascent polypeptide chains [72]. We were unable to detect homologues of domain B by sequence comparisons outside the TPPII group and no other proteins of the same fold were found using DALI. All proteins of the TPPII group have domain B (Fig. 5C), including the bacterial and archaeal orthologs, and the satellite clusters (STABLE, Thermaerobacter). Domain C is not so much a domain as an elongated, structured protrusion within domain B; it is difficult to detect by sequence comparisons due to its small size and poor conservation, so not much can be concluded from its apparent absence in many proteins containing domain B. The four domains D, E, F, and G in the central part (ii) of TPPII are all β-domains, with G inserted into domain F. Domains D and F are

typical IG folds of the PapD superfamily, E is a jelly-roll β-sandwich of unclear origin, and G is homologous to the collagen-binding domain of class 1 collagenase by structure and sequence. Domain F may be involved in dimerization, since in TPPII dimers it localized at the monomer–monomer interface. Clearly, domain D is present in all TPPII relatives analyzed here, including the actinobacterial anchor cluster (blue in Fig. 5A and C), and distinguishes these proteins from other subtilisins. Domain E is present in the TPPII group, including its satellites, and domains F and G are only reliably detected in the eukaryotic TPPII core cluster. Most proteins analyzed here contain other, seemingly unrelated domains following domains D and E. The C-terminal part (iii) of TPPII, i.e. domain H, is a solenoid of five hairpins with strong similarity to tetratricopeptide repeats (TPR), which are often involved in protein–protein-interactions [73,74]. This is suggestive, since domain H is the region flanking the entrance to the cavity system of TPPII. Indeed, the last two hairpins are clearly identified as TPR and the domain as a whole is recognized by HMM–HMM sequence comparisons as a TPR-like solenoid. Domain H seems to be specific for the eukaryotic TPPII cluster, however we note that pyrolysins also seem to contain a much shorter version of domain H consisting of only two TPR hairpins, which is not necessarily of homologous origin, as it could have arisen by an independent fusion event. Except for TPPII, which is intracellular, all other TPPII-related proteases have signal peptides (as predicted with SignalP, [75]; Fig. 5C). In addition, most of them have a propeptide located N-terminally to the subtilisin domain, which appears to be a subtilisin-specific intramolecular chaperone [76]. Its cleavage leads to activation of the subtilases [76], which may explain its absence in TPPII, where activation is coupled to assembly [57]. Indeed, this is one of several indications that higher oligomeric assembly might be specific to eukaryotic TPPII. Thus, domains F (which may mediate dimerization) and H (which interacts with the insert domain C to give TPPII monomers their peculiar shape for spindle assembly) are also specific to these proteins. These analyses suggest that the earliest form of a TPPII-like protease was an extracellular protease containing domains A and D. The TPPII branch proper subsequently acquired domains B and E, and the eukaryotic form of the protein finally lost the signal sequence and propeptide, while adding domains F, G, and H. It is attractive to consider that the re-localization of the protease to the cytosol following loss of the signal sequence provided the evolutionary pressure for segregating the active sites by self-compartmentalization and thus for recruiting domains F and H. 6. Conclusions In recent years, substantial progress has been made in elucidating the structure and function of TPPII. Structural studies have revealed an intriguing architecture but why TPPII forms an assembly of such extraordinary size remains a mystery. The high-resolution structure of TPPII obtained by using a hybrid EM-X-ray crystallography approach has revealed an extensive network of internal cavities and channels, which sequester the active sites from the cytosolic environment and control access to them. The structure suggests a mechanism by which the essentially inactive dimers are converted into active ones upon assembly; activation follows sequestration of the active sites [12] — a common theme among self-compartmentalizing proteases. A doubleGlu motif in the catalytic domain provides a molecular ruler for the cleavage of tripeptides, as verified by mutational studies [10,12]. A putative endoproteolytic activity and its functional significance await further clarification. The list of physiological functions ascribed to TPPII in health and disease is long but in almost all cases the exact role of TPPII remained enigmatic. The grand challenge for the years to come is to firmly establish the role of TPPII in cellular processes. In the case of the proteasome the dissection of its physiological functions was greatly facilitated by the availability of specific proteasome inhibitors [77,78].

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Fig. 5. Analysis of TPPII-like proteins: Domain composition and cluster map. A, B) Cluster maps. We used the buildali.pl script of HHpred with the sequence of TPPII as query to search the non-redundant database of NCBI for TPPII homologs. The script performs iterative PSI-BLAST runs and contains heuristics to reduce false positive matches caused by nonhomologous sequence segments at the end of PSI-BLAST matches. The obtained sequences were clustered to equilibrium in CLANS [70] at a P-value cutoff of e-25 using default settings. In the cluster maps, dots represent proteins, while the gray lines represent BLAST p-values — the darker a line, the more significant the p-value. Both panels show the same map but with different color schemes. In A), proteins are colored according to their group membership, while in B they are colored by superkingdom (blue = Archaea, yellow = Bacteria, green = Eukaryota). Subtilisin Carlsberg is marked in black as reference outgroup. Proteins shown in white are distantly related to TPPII and were not analyzed in this study. The blow-up shows a magnified view of the TPPII cluster. C) Table of domain compositions of TPPII-like proteins. Rows correspond to the different groups (color-coded as in A), while columns correspond to the constituent domains, detected by SignalP [75] and HHpred [69]. D) Eight-domain composition of TPPII. Domain borders for the basic domains are indicated below the colored bar, domain borders for the inserted domains are indicated above the colored bar. A′ denotes the N-terminal, A″ denotes the C-terminal part of domain A, the same code is used for the divided domains B and F. The dashed line between domains F″ and H represents the missing loop L3. E) Ribbon representation of a TPPII monomer (left) and octamer (right) color coded as described in D. The dashed line between domains F and H corresponds to loop L3.

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