Solution Structures of UBA Domains Reveal a Conserved Hydrophobic Surface for Protein–Protein Interactions

doi: 10.1016/S0022-2836(02)00302-9 available online at http://www.idealibrary.com on

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J. Mol. Biol. (2002) 319, 1243–1255

Solution Structures of UBA Domains Reveal a Conserved Hydrophobic Surface for Protein –Protein Interactions Thomas D. Mueller and Juli Feigon* Department of Chemistry and Biochemistry, Molecular Biology Institute, University of California at Los Angeles, Los Angeles, CA 90095-1569, USA

UBA domains are a commonly occurring sequence motif of , 45 amino acid residues that are found in diverse proteins involved in the ubiquitin/ proteasome pathway, DNA excision-repair, and cell signaling via protein kinases. The human homologue of yeast Rad23A (HHR23A) is one example of a nucleotide excision-repair protein that contains both an internal and a C-terminal UBA domain. The solution structure of HHR23A UBA(2) showed that the domain forms a compact three-helix bundle. We report the structure of the internal UBA(1) domain of HHR23A. Comparison of the structures of UBA(1) and UBA(2) reveals that both form very similar folds and have a conserved large hydrophobic surface patch. The structural similarity between UBA(1) and UBA(2), in spite of their low level of sequence conservation, leads us to conclude that the structural variability of UBA domains in general is likely to be rather small. On the basis of the structural similarities as well as analysis of sequence conservation, we predict that this hydrophobic surface patch is a common protein-interacting surface present in diverse UBA domains. Furthermore, accumulating evidence that ubiquitin binds to UBA domains leads us to the prediction that the hydrophobic surface patch of UBA domains interacts with the hydrophobic surface on the five-stranded b-sheet of ubiquitin. Detailed comparison of the structures of the two UBA domains, combined with previous mutagenesis studies, indicates that the binding site of HIV-1 Vpr on UBA(2) does not completely overlap the ubiquitin binding site. q 2002 Elsevier Science Ltd. All rights reserved

*Corresponding author

Keywords: ubiquitin; NMR; HHR23A; Vpr; proteasome

Introduction The human homologue of yeast Rad23A (HHR23A) is an evolutionarily conserved protein involved in DNA nucleotide excision-repair.1 – 3 The protein, like all Rad23 homologues, has a modular structure that includes an N-terminal ubiquitin-like (Ubl) domain and two ubiquitinassociated (UBA) domains, located in the middle (UBA(1)) and at the C terminus (UBA(2)) of the protein. A region between the two UBA domains, which does not clearly exhibit all signs of a globular domain motif, is responsible for binding to the xeroderma pigmentosum C (XPC) protein (yeast homologue RAD4) which is part of the DNA repair Abbreviations used: UBA, ubiquitin-associated; XPC, xeroderma pigmentosum C. E-mail address of the corresponding author: [email protected]

complex.4,5 The Ubl domain has been shown to interact directly with the 26 S proteasome6,7 and is required for full DNA repair function in the yeast protein,1 suggesting a link between DNA repair and protein degradation via the ubiquitin-proteasome pathway. The UBA domain is a short sequence motif of , 45 amino acid residues that occurs frequently in proteins found in all eukaryotes.8 It is found in many enzymes of the ubiquitination pathway, and in UV excision repair proteins and protein kinases involved in cell-signaling pathways and cell-cycle control. On the basis of the frequent appearance of UBA domains in the ubiquitin/proteasome pathway, Hofmann & Bucher suggested that UBA domains might possibly bind ubiquitin, although no direct evidence leading to this conclusion was given.8 Subsequently, it was shown that p62, the phosphotyrosine independent ligand of p56lck, interacts with ubiquitin directly.9 The ubiquitin binding was

0022-2836/02/$ - see front matter q 2002 Elsevier Science Ltd. All rights reserved

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confined to the C-terminal 80 residues later identified as a UBA domain that is rather distant from the UBA domain family. Although the authors did not propose the interaction with ubiquitin as a general function for all UBA domains, it was speculated that UBA domains are involved in the recognition and binding of ubiquitin in protein degradation pathways for regulatory purposes. Very recently, it was shown by studies both in vitro and in vivo that both UBA domains of HHR23A,10 – 12 as well as another DNA damageinducible protein DDI1,10 bind to ubiquitin directly. The demonstrated interaction of members of the UBA domain family with ubiquitin along with the data linking HHR23A with the ubiquitin/ proteasome pathway led us to investigate the structures of UBA domains further in order to see if they had structural similarities that might suggest a common binding interface for ubiquitin or other proteins. In addition to the possible common function for UBA domains of binding ubiquitin, several other binding partners that bind to the C-terminal UBA domain of only HHR23A have been identified, suggesting that individual UBA domains may have a more specific function. For example, binding of HIV-1 Vpr,13 3-methyladenine DNA glycosylase (MPG),14 the deglycosylating enzyme N-glycanase Png1,15 as well as the transcription regulator p300/CBP,16 bind to HHR23A at UBA(2) only. Binding of the ubiquitin-ligase E6-AP to HHR23A might be mediated through UBA domains, although evidence for a direct interaction with the UBA domains was not shown. However, deletion of the N-terminal ubiquitin-like domain of HHR23A did not abrogate the binding, suggesting that the interaction of E6-AP with HHR23A must be confined to the C-terminal part.17 The solution structure of HHR23A UBA(2) revealed that the domain forms a compact threehelix bundle with an unusually large hydrophobic surface patch.18,19 The probable source for the specificity of the interaction of HIV-1 Vpr and UBA(2) was determined from the solution structure of a UBA(2) mutant protein P333E, which is deficient in binding to HIV-1 Vpr.19 Small but significant changes in the loop conformation introduced by the amino acid exchange as well as the presence of a negatively charged residue in the hydrophobic surface patch are probably responsible for the loss in binding. The UBA(1) domain has a glutamate residue instead of proline at the same position. It was therefore of interest to determine whether the loop in UBA(1) adopts a similar conformation to that seen in the UBA(2) mutant P333E. Here, we present the three-dimensional solution structure of UBA(1) of HHR23A solved by heteronuclear multidimensional NMR spectroscopy. Like UBA(2), UBA(1) forms a compact three-helix bundle that is remarkably similar to UBA(2), in spite of a low level of sequence homology in the hydrophobic core. Comparison of the structures of

Solution Structures of UBA Domains

UBA(1) and UBA(2) reveals that both domains have a large hydrophobic surface patch. Analysis of the sequence conservation in this hydrophobic patch shows a region with a high level of sequence conservation that is not required to maintain the local structure and is therefore a likely interface for specific protein interactions. On the basis of these results, we predict that UBA domains might interact via their hydrophobic surface patch with the hydrophobic epitope found on the surface of the five-stranded b-sheet of ubiquitin.

Results and Discussion Preparation of UBA(1) for NMR spectroscopy For cloning, the boundaries for the UBA domain (residues 155 –204) were chosen on the basis of the sequence alignment in the PFAM database† and the predicted secondary structure using the PHD program. The purified glutathione-S-transferase fusion protein was homogeneous, with no signs of partial proteolytic degradation due to bacterial proteases. The overall yield of UBA(1) protein was about 10 mg per liter of bacterial culture (in rich media) and up to 7 mg per liter for minimal media (15N labeling using 6 g/l glucose). The use of Escherichia coli strain Bl21(DE3)Star increased the yield compared to normal Bl21(DE3) by a factor of 2. The final UBA(1) protein was very stable with no signs of secondary protease cleavage sites even if the incubation with thrombin was extended for more than 48 hours. Analysis of the line-width of the NMR signals indicated that there was no dimerization or aggregation even at a concentration of 3 mM and above for the isolated UBA(1) domain protein. However, Bertoleat et al., report that RAD23 forms homodimers as well as heterodimers with the protein DDI1 in vitro and in vivo.10 The UBA domains of either protein were shown to be necessary for this dimerization. An explanation for the discrepancy might be in the fact that no data were provided as to whether one or both UBA domains of RAD23 are required for the dimerization. It is therefore possible that the dimerization seen for RAD23 is a result of the interaction of UBA(1) with UBA(2). In that case, no aggregation or dimerization would be seen for an isolated UBA(1) or UBA(2) domain. NMR analysis Although UBA(1) exhibits good chemical shift dispersion in the homonuclear 2D nuclear Overhauser effect (NOESY) and total correlated (TOCSY) spectroscopy experiments, there is considerable overlap in the aliphatic region of the spectra, despite the small size of the protein, due to the mainly alpha-helical nature of the protein domain. We therefore used heteronuclear NMR † http://www.sanger.ac.uk/Software/Pfam

Solution Structures of UBA Domains

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Figure 1. The 2D 1H – 13C correlation strip pairs (left strip CBCA(CO)NH, right strip CBCANH) used for sequential assignment of residues Gly174 to Val 180. The strips are taken at the 15N chemical shift of the particular residue. The CBCA(CO)NH spectra contains only the Ca and Cb chemical shift information of residue i 2 1 correlated with the amide proton and nitrogen chemical shift of residue i. The corresponding CBCANH spectrum contains, in addition, the carbon chemical shift information for residue i. Red contours indicates negative levels, which are specific for Cb spins or Ca spins for glycine residues. Asterisks (p ) mark peaks missing due to low intensities.

techniques in order to obtain a high-resolution solution structure of UBA(1). Two triple-resonance NMR experiments, CBCA(CO)NH and CBCANH, were acquired for the sequential assignment of the backbone resonances.20 The processed data were striped using the software AURELIA (Bruker Karlsruhe) on the basis of a peak list generated from the 2D 15N – 1H heteronuclear single-quantum coherence (HSQC) spectra (Figure 1). This resulted in 47 1H – 13C correlation strips each for the CBCA(CO)NH and the CBCANH spectra. A starting strip pair was then chosen randomly and the database containing all the strips was searched for overlapping Ca and Cb carbon chemical shift information in both experiments. The interactive search tool in AURELIA allows these searches to be done very efficiently, with the advantage of visually inspecting possible hits for spectral artifacts. The HBHA(CO)NH spectra were used to generate additional proton chemical shift information for Ha/b for the interpretation of the HCCH-correlated (COSY) and HCCH-TOCSY spectra in order to obtain complete side-chain assignments.20 Complete resonance assignments were obtained for all residues from Ser155 to Gly204, with the following exceptions. No signal was detected for the N-terminal glycine residue, only proton chemical shift information was assigned for the aromatic protons, and no proton or carbon chemical shift information for Met166 methyl group was obtained. Structure of HHR23A UBA(1) The final structure calculations employed a total of 1692 NOE-derived distances, of which 1055 distances are non-redundant in that they do not

contain any duplicate from either the 3D or 2D data. Of these 1055 distances, 407 are intraresidue, 212 are sequential, 226 are medium-range (li 2 jl # 4) and 210 are long-range (li 2 jl . 4) (Table 1). With an average of 21.5 NOE distance restraints per residue, the structures of UBA(1) are very well defined. Out of 100 calculated structures, ten were selected on the basis of their lowest overall and NOE energies. None of the selected structures exhibited any NOE violation greater than ˚ , and all structures showed good covalent 0.5 A geometry with no bond or angle violation. The f,c distribution reveals 94.8% of all residues are within the most favorable or additional allowed regions, and if only the structured regions (from residue 160– 201, see Figure 2(a)) are considered, only one residue out of ten structures is found in the disallowed region. A superposition of the backbone atoms of the ten lowest-energy structures is shown in Figure 2(b) and structural statistics are given in Table 1. The r.m.s. deviation from the mean structure for residues 162– 198, which form the well-defined ˚ for the backbone core of the protein, is 0.2 A ˚ for all heavy-atoms. UBA(1) atoms and 0.9 A forms a very compact three-helix bundle; helix 1 consists of residues 161– 172, helix 2 from 177 –185 and helix 3 from 192 –200. Helix 1 and helix 2 are connected by a highly ordered, four residue loop (loop 1), and helix 2 and helix 3 are connected by a six residue loop (loop 2). The side-chains of residues Tyr163, Ala166, Leu167, Ile170, Tyr175, Val180, Val181, Leu184, Ala194 and Leu198, which form the hydrophobic core that stabilizes the protein, are shown in Figure 2(b). Due to the compactness of the three-helical bundle, several of these residues are not buried completely in the

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Solution Structures of UBA Domains

Table 1. Input data for structural calculations and statistics for UBA(1) (ten structures) Distance restraints Total NOE-derived Intraresidue Sequential (li 2 jl ¼ 1) Medium-range (li 2 jl # 4) Long-range (li 2 jl . 4) Average NOE/residue Hydrogen bonds

1055 (1692) 407 212 226 210 21.5 24

Structure statistics ˚) RMSD from experimental distance restraints (A RMSD from idealized geometry ˚) Bonds (A Angles (deg.) Energies (kcal mol21) ENOE Ebond Eangle EvdW

0.044 ^ 0.0007 0.006 ^ 0.0002 0.721 ^ 0.009 165.6 ^ 5.7 23.6 ^ 1.4 109.9 ^ 2.7 40.2 ^ 1.3

˚) RMSD from average structure (A Backbone atoms for residues from 164–200 (ten submitted structures) All heavy-atoms for residues from 164– 200 (ten submitted structures) Backbone atoms for residues from 164–200 (all 100 structures) All heavy-atoms for residues from 164– 200 (all 100 structures) Backbone atoms for residues from 164–200 (50 structures with lowest energy) All heavy-atoms for residues from 164– 200 (50 structures with lowest energy)

0.21 ^ 0.05 0.86 ^ 0.12 0.33 ^ 0.08 1.01 ^ 0.10 0.29 ^ 0.08 0.96 ^ 0.09

PROCHECK analysis Residues in most favored region (%) Residues in additional allowed region (%) Residues in generously allowed region (%) Residues in disallowed region (%)

72.9 (77.9) 21.9 (18.2) 4.0 (3.7) 1.2 (0.8)

RMSD between average structures of UBA(1) and UBA(2) Residues 164 –199 (UBA(1)) and 321 to 356 (UBA(2)) backbone atoms only Three helices only

˚ 1.6 A ˚ 1.2 A

Inter-helical angles (deg.)

UBA(1)

UBA(2)

a2/a1 a3/a2 a1/a3 ˚) Helix to helix distance (A a2/a1 a3/a2 a1/a3

116 ^ 3 105 ^ 2 47 ^ 1

120 126 30

8 8 10

9 8 11

interior of the hydrophobic core and therefore display a large portion of their hydrophobic side-chain on the surface; only the side-chains of residues Leu167, Val180, Leu184 and Ala194 are shielded from the solvent completely. The N-terminal residues Ser155-Gly160 are disordered and seem to be flexible. These residues showed only sequential NOEs and had large TOCSY crosspeaks consistent with rapid internal motions. In contrast, the C terminus, which contains several hydrophobic residues, adopts an extended conformation. Several NOEs between residues Ile202 and Tyr197, and between Ala183 and Ala186 result in folding of the C terminus back onto the compact three-helix bundle. The angles between pairs of adjacent helices are 1168 for a1 and a2 (equivalent to 2 648) and 1058 for a2 and a3 (or 2 758). These values are reasonably close to the theoretical optimum of 528 for

tight “knob-in-hole” packing of side-chains, although slightly larger than those for UBA(2) (Table 1). Helix 1 follows the rules for helix capping,21 with the N-cap residue Glu162, which can serve as a possible hydrogen bond donor, followed by Tyr163 (N1), Glu164 (N2), Thr165 (N3) and Met166 (N4). The C-cap of helix 1 consists of a hydrogen bond between the carbonyl group of Met171 and the amide proton of Gly174 in loop 1. A glycine residue is required in this position in order to adopt the necessary conformation, since the f,c-dihedral angles for other amino acid residues would be in the disallowed region. Consistent with this, the sequence alignment for UBA domains clearly shows a high level of conservation of this glycine residue in loop 1 (see Figure 4(a)). The N-cap of helix 2 consists of the residues Glu176-Val180 (N-cap through N4), with Glu176 possibly forming a sidechain hydrogen bond to

Solution Structures of UBA Domains

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Figure 2. Stereoviews of the internal UBA domain UBA(1) of HHR23A (SWISS PROT code R23A_HUMAN) (human homologue of RAD23A) showing residues Thr156 to Gly204. (a) Ribbon representation. The three helices are labeled a1, a2, and a3. (b) Superposition of the ten lowest-energy structures. The backbone atoms are shown in black for carbon atoms, blue for nitrogen atoms, and the carbonyl oxygen atoms are omitted. The side-chains for residues forming the hydrophobic core are shown in green. The N-terminal residues 155– 160 are disordered and are not shown.

the backbone amide group of Glu178. C-capping of helix 2 is achieved by forming possible hydrogen bonds between the sidechain amide protons of Asn190 and the backbone carbonyl group of Tyr188 and Ser189. The high degree of conservation for this Asn residue in loop 2 indicates a possible role in structure stabilization. Finally, the Ncapping of helix 3 is formed by a proline residue, which induces the helix with its special f,c dihedral angle requirements. The residues Ser172 (i ) to Tyr175 (i þ 3) of loop 1 adopt a type IV b-turn conformation with a distance between the Ca atoms of ˚ . The six residue Ser172 and Tyr175 of 6.5 A (Arg185-Asn190) loop between helices 2 and 3 can be analyzed as two conjugated b-turns. The first

b-turn is formed by the residues Arg185 (i ) to Tyr188 (i þ 3) of loop 2. This turn conforms to the type I classification showing f,c-dihedral angles for Ala186 (i þ 1) and Ser187 (i þ 2) that are close to helical values (f , 2 60 to 2 908; c , 0 to 2 308). The second turn forms a type I0 conformation, in which the residues at position i þ 1 (Tyr188) and i þ 2 (Asn189) exhibit f,c-dihedral angles in agreement with a left-handed helix. UBA(1) has a large hydrophobic surface patch that includes several hydrophobic residues from the hydrophobic core, whose side-chains are partially exposed on the surface (Figure 3(a)). This hydrophobic surface patch includes residues Met171 in helix 1, Met173 through Tyr175 in loop 1, Val195,

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Solution Structures of UBA Domains

Figure 3. Stereoviews of superposition of UBA(1) (blue) and UBA(2) (red) (C-terminal UBA domain of HHR23A, PDB entry code 1DV0). The blue and red vectors represent the helix axes of UBA(1) and UBA(2), respectively. The proline residue 333 in UBA(2), which is critical for the binding of HIV-1 Vpr to the UBA(2) domain, is marked. UBA(1) does not have a proline residue at that position (instead a proline residue is found in the loop between helix 2 and helix 3). The loop conformations for both UBA domains are therefore slightly different on the C-terminal side of the proline residues.

Leu198 and Leu199 in helix 3, and Gly201 and Ile202 in the C-terminal extension. The surface area of the hydrophobic surface patch is about ˚ 2 (molecular surface), which corresponds to 470 A approximately 17% of the total surface area of ˚. about 2830 A

Comparison of HHR23A UBA(1) and UBA(2) reveals that the structural variability of UBA domains is small The overall fold of HHR23A UBA(1) and UBA(2) is very similar (Figure 3), in spite of the relatively Figure 4. A potential protein– protein binding interface of UBA domains is built from hydrophobic residues on the surface. (a) Surface representation of UBA(1) (left) using the following color coding: red, acidic amino acid residues Glu and Asp; blue, basic amino acid residues Arg and Lys, orange, polar amino acid residues Asn, Gln, His, Ser and Thr; white, hydrophobic residues Ala, Gly, Phe, Ile, Pro, Met, Leu, Tyr and Val. The major accessible residues on the hydrophobic surface, Met173, Gly174, Y175, L199 and I202, are marked. The size of the epitope is ˚ 2. The right approximately 470 A picture shows the orientation of the helical bundle with respect to the surface representation. The hydrophobic surface patch consists mainly of residues from loop 1 between helices 1 and 2 as well as residues from helix 3. (b) For comparison, the surface of UBA(2) is shown in the same orientation as UBA(1), revealing that the location of the hydrophobic epitope is indeed conserved and consists of identical or homologous residues. The C terminus of UBA(2) is not shown, due to its flexibility.

Solution Structures of UBA Domains

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Figure 5. (a) Sequence comparison of selected UBA domains (sequence alignment retrieved from PFAM database and edited to contain only sequences from the SWISS PROT database). The analysis was performed using the program Jalview‡ using the seed alignment for UBA domains in the PFAM database† (see Materials and Methods) accession code PF00627. The Taylor color-coding was used for primary coloring,40 in addition, the degree of conservation for each residue position was calculated and the brightness of the color was then correlated with the degree of conservation. Columns with very bright colors show the highest level of amino acid conservation. Arrows point to the amino acid residues that are shown labeled in the surface representations of (b) UBA(1) and (c) UBA(2). The surface representations are color-coded the same as used for the sequence alignment in (a). The amino acid residues occupying conserved positions on the hydrophobic surface patch are labeled, with the amino acid found in (b) UBA(1) and (c) UBA(2) underlined.

low level of sequence identity of 20% (see Figure 5(a)). The r.m.s. deviation between UBA(1) and UBA(2) for the backbone atoms of the three-helical ˚ , and if only the three helices bundle is about 1.6 A without the connecting loops are considered, the

˚ (see Table 1). Both r.m.s. deviation is about 1.2 A domains form compact, three helix bundles with helices of similar lengths, which are stabilized by a hydrophobic core and have an unusually large and conserved hydrophobic surface patch. The

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hydrophobic patch for UBA(2) is comprised of the residues Leu327, Leu330 through Pro333, Leu336, Ala340, Ala352, Leu355 and Leu356, and measures ˚ 2 out of a total surface area of about about 440 A ˚ 2 (Figure 4(b)). The major differences 2910 A between UBA(1) and UBA(2) are the angles between the axes of helix 1 and helix 3, which differ by about 178, and helix 2 and helix 3, which differ by about 218. The difference in the interhelical angle between helix 1 and helix 2 is less than 58 and, therefore, within the limit of error of the structure determination by NMR (standard deviation for the ten structures of UBA(1) is 38). The low level of sequence identity for residues located inside the hydrophobic core might explain the differences in the inter-helical angles (Table 1 and Figure 2). Between UBA(1) and UBA(2), only a few of the residues in the hydrophobic core are identical, i.e. Val180/Val337, Ala183/Ala340, Ala194/Ala351 and Leu198/Leu355. For the other positions in the hydrophobic core, residue types are swapped between the helices, so that the overall space requirement yields a similar packing, e.g. the triplet Met166/Leu184/Ala194 occupies a similar amount of space in UBA(1) as the triplet Ala323/Tyr341/Ala351 in UBA(2). This results in almost identical helix to helix distances for both proteins (Table 1), showing that the overall packing efficiency is the same. Also, the loop conformations for UBA(2) are very similar to those of UBA(1). Loop 1 of UBA(2) between helix 1 and helix 2 adopts a type IV b-turn with f,c-dihedral angles almost identical with those seen for UBA(1). Loop 2 of UBA(2) is likewise composed of two b-turns; residues Phe342-Glu345 form the first type I b-turn but the second b-turn for the residues Cys344-Asn347 is type IV instead of I0 as seen for UBA(1). The difference is significant, but an inspection of the f,c-angles in the Ramachandran plot reveals that the i þ 1 and i þ 2 residues of the second b-turn of both UBA domains adopt conformations that are close to a left-handed a-helix. The equivalent residues in loop 2 of UBA(1) adopt backbone torsion angles that fall into this category. The structural similarity between UBA(1) and UBA(2), in spite of their low level of sequence conservation, leads us to conclude that the structural variability of UBA domains in general is likely to be rather small. Significance of the hydrophobic patch Large hydrophobic surface patches on proteins are not that common and when present are often binding sites for other proteins. The striking similarity between the large hydrophobic surface patches of UBA(1) and UBA(2) might indicate that this is a common binding interface for all UBA domains. In the absence of other data, it could be argued that the hydrophobic surface patch is simply a consequence of the compactness of the three helix bundle. In contrast to four helix bundles, the side-chains of the bulkier residues

Solution Structures of UBA Domains

involved in forming the hydrophobic core of three helix bundles are partially accessible at the surface of the protein. However, analysis of the sequence alignment for UBA domains (PFAM database†) (Figure 5(a)) in conjunction with the structure information for UBA(1) and UBA(2) (Figure 4) reveals that several conserved residues in this hydrophobic patch (Figure 2) are not absolutely required for structure stabilization. The program Jalview‡ was used together with the “seed” sequence alignment for UBA domains as deposited in the PFAM database to analyze the degree of conservation of individual residue positions throughout the UBA family (Figure 5(a)). In addition to residues located inside the hydrophobic core, which are well conserved for structural stability reasons, several residues located mostly on the surface are among the most conserved positions. Several residues, especially in loop 1, are much more conserved than would be necessary simply to adopt the local structure. Specifically, loop 1 (Met/Leu-Gly-Phe/Tyr), loop 2 (Asn-X-polar), and two positions on helix 3, which are solventaccessible, are among the most highly conserved residues in the UBA domain family. The motif Met/Leu-Gly-Tyr/Phe (UBA(1) loop 1 residues 173 –175) is even more conserved than residues inside the interior of the helix bundle. Several residues on helix 2 and helix 3, namely Ala183/ Ala340, Leu198/Leu355 and Leu199/Leu356, are also highly conserved and form part of the hydrophobic surface patch. These most conserved residues form a consecutive area on the solventaccessible surface comprised of a subset of the residues that form the hydrophobic surface patches on UBA(1) and UBA(2), as shown in Figure 5(b) and (c). This conserved hydrophobic region likely indicates a common function, e.g. a common binding interface for all UBA domains. Other highly conserved residues, for example the asparagine residue located in loop 2 of the UBA domains, are either important for structural reasons (Asn191/ Asn348 form the C-cap for helix 2) or are close to the hydrophobic patch (His192 UBA(1) and Asn350 UBA(2)) and might be part of the binding interface. Thus, we propose that the conserved region of the hydrophobic surface on UBA(1) and UBA(2) represents a common binding surface for interacting with a specific protein(s). Is the hydrophobic patch a conserved binding site for ubiquitin? What sort of proteins might these be? As discussed, several UBA domains, specifically those found in p62, HHR23A, DDI1 and Rph23, have been shown to interact directly with ubiquitin.9 – 12 The binding affinities for the HHR23A UBA(1), HHR23A UBA(2), and DDI1 UBA domains with † http://www.sanger.ac.uk/Software/Pfam ‡ http://www.ebi.ac.uk/~michele/jalview/

Solution Structures of UBA Domains

monomeric ubiquitin were reported to be in the low micromolar range, i.e. about 10 mM for HHR23A UBA domains and 8 mM for the DDI1 UBA domain.10 In order to define the binding site on the UBA domains for ubiquitin, a mutagenesis study was performed for the UBA domains of HHR23A and DDI1.10 Two identical positions in both UBA(1) and UBA(2) were chosen and mutated to alanine. The first residue is Glu176 in UBA(1) and Leu336 in UBA(2), which did not abrogate the binding to ubiquitin upon mutation to alanine. The second residue position was Leu198 in UBA(1) and Leu355 in UBA(2), which resulted in both cases in a loss of binding. Since the first position is not part of the conserved surface patch, we suggest that this residue position is not involved in the binding of ubiquitin. However, the second residue is a highly conserved leucine residue, buried completely in the interior of the three helix bundle core. Therefore, we conclude that the observed loss of binding for this mutation is due to loss of structural integrity and does not provide any information on the binding interface. For the UBA domain of DDI1, additional point mutations were analyzed.10 On the basis of our structural analysis, the additional mutants of DDI1 were clustered into two groups. In the first group, point mutations were introduced at positions in the hydrophobic core that our structural analysis indicates would disrupt the tertiary structure, and therefore the observed decrease in binding of ubiquitin is likely to be an indirect effect. In the second group, the mutations are all located in loop 2 and helix 2 of the three helix motif. Residue Lys413 of DDI1 (equivalent to Arg185 in UBA(1) and Phe342 in UBA(2)) located in helix 2 shows a significant change in binding when mutated to isoleucine. This position is located in a second hydrophobic surface patch, which is smaller and for which residues are less highly conserved throughout the UBA domain family. Another position leading to a significant loss of binding is residue Gly417 (Asn189 UBA(1) and Lys346 UBA(2)) located in loop 2. This position is close to the proposed binding site and could be part of the proposed binding site. However, this position does not show a high degree of conservation (amino acid residues range from Gly, Asn, Lys, Tyr to Trp) (Figure 5(a)). In summary, for the second group of variants for which a loss of structure is not likely, two positions that are non-conserved in the UBA family do show some alteration in binding. In a very recently published study, binding of the UBA domains of the RAD23 homologue of fission yeast, Rhp23, was examined by surface plasmon resonance and site-directed mutagenesis.12 In contrast to the previous reports, the authors report very tight binding of the UBA domains to tetrameric ubiquitin in the range of 30 nM as compared to 10 mM found for the interaction of RAD23 and monomeric ubiquitin. In addition, in their BIAcore binding experiment, no affinity (detection limit . 1 mM) to monomeric ubiquitin was observed.

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This finding is surprising in light of other reports, which showed that UBA domains prevent multiubiquitylation of target proteins.11,22 The number of attached ubiquitin moieties was decreased efficiently from more than four to a single ubiquitin moiety. Therefore, it seems plausible, that binding of the UBA domains to a single attached ubiquitin moiety is the key step of this inhibition mechanism. In the mutagenesis study, the highly conserved glycine residue in loop 1, which is a central residue in our proposed ubiquitin interaction epitope, was exchanged to a proline residue in both Rhp23 UBA domains. When only one UBA domain was mutated, binding to tetrameric ubiquitin was still detected, although at reduced levels, thereby indicating that the binding of the UBA domains to tetrameric ubiquitin is independent and not cooperative. The double mutant did not retain any binding to ubiquitin, suggesting that the glycine residue in the conserved loop 1 is indeed in the center of the binding interface. Ubiquitin has a hydrophobic surface patch across its five-stranded b-sheet where several residues that have been identified as the major binding determinants for the interaction with the proteasomal subunit S5a are located. Crystallographic studies of di and tetra-ubiquitin reveal that the hydrophobic area is extended upon formation of multi-ubiquitin chains and thereby provides insight into how the affinity is increased with increasing numbers of ubiquitin moieties.23,24 This hydrophobic area on ubiquitin is utilized also in the interaction of monomeric ubiquitin with enzymes of the ubiquitin/proteasome pathway. NMR mapping studies of the interaction of ubiquitin with several ubiquitin ligases (E2) indicate that this epitope is also a general binding interface used in the generation of polyubiquitin chains.25 – 29 Using this information, we suggest that the interaction of UBA and ubiquitin takes place between the two hydrophobic surface patches of these proteins. Consequently, the stabilization of proteins through UBA domains could be a result of a competitive inhibition of the binding to the proteasome. Considering that the binding affinity of Rhp23 UBA domains to tetrameric ubiquitin is of the same order of magnitude as that reported for the in vitro binding of tetrameric ubiquitin to the S5a subunit,12,30 a blockage of the targeting of polyubiquitylated proteins to the proteasome seems to be a possible mechanism. Although this is in contrast to the observation that prevention of protein degradation is due to inhibition of polyubiquitin chain extension by UBA domains,11,22 both mechanisms might be possible, depending on the kind of ubiquitin linkage. The amino acid sequence of loop1 of UBA(2) is critical for specific binding by HIV-1 Vpr The C-terminal UBA domain of HHR23A UBA(2) binds specifically to HIV-1 Vpr, whereas UBA(1) and other UBA domains do not.13,18 We

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proposed previously that the hydrophobic surface on UBA(2) could be a binding site for HIV-1 Vpr, and showed that mutation of Pro333 to Glu in the hydrophobic loop 1 completely abolishes binding of HIV-1 Vpr to UBA(2).19 The structure of UBA(2) P333E, a Vpr-binding-deficient mutant of UBA(2), showed that exchange of the proline residue with a glutamate residue results in a small but significant change in the loop conformation around the mutation site. The largest changes in backbone torsion angles were observed for the mutated residue Pro333 and the preceding residue Phe332. The c angle of residue 342 of UBA(2) changes by about 358 and the c angle of residue 333 also differs by 358 upon mutation of proline to glutamate. Since in UBA(1) a glutamate residue (Glu176) occupies the same position, could this explain why UBA(1) does not bind to HIV-1 Vpr? A superposition of the structures of UBA(1) with either wild-type UBA(2) or mutant P333E was analyzed with regard to whether the backbone torsion angles of the residues Tyr175 and Glu176 of UBA(1) are closer to those seen for the mutant P333E than for the wild-type UBA(2). If so, this would provide an indication that the loop conformation as well as the composition of loop 1 is very important for the binding of HIV-1 Vpr. However, much to our surprise, analysis of the f,c-angles of the residues in loop 1 revealed that the backbone conformation of loop 1 of UBA(1) is not similar to either wild-type UBA(2) or the mutant P333E. The c angle of Tyr175 (equivalent to Phe332 in UBA(2)) is ,908, whereas the c angles for UBA(2) and its mutant P333E are between 21758 and 21408. The c-angle of the following residue, Glu176 (equivalent to either Pro 333 (wild type UBA(2)) or Glu333 (UBA(2)P333E), also differs significantly, with a value of 2408 for Glu176 of UBA(1) and 21608 for Glu333 for UBA(2). Consequently, the carbonyl oxygen atom of UBA(1) Tyr175 points into the interior of the three-helix bundle, whereas for the two UBA(2) structures, the carbonyl oxygen atom of the equivalent residue 333 is oriented toward the solvent-accessible surface. This finding may be explained by differences in the amino acid sequence following residue Glu176 and Pro333 in loop 1 of UBA(1) and UBA(2), which results in slightly different packing for individual residues in this loop. For example, in UBA(2) Leu336 folds into the hydrophobic core, whereas the side-chain of the equivalent residue in UBA(1), Arg179, points into the solvent. As a consequence, we conclude that the amino acid composition of loop 1 is critical for the binding of HIV-1 Vpr, with Pro333 in the center of the binding epitope forming an enlarged hydrophobic patch not present in either UBA(1) or the mutant UBA(2)P333E.

Conclusions We have shown that the overall structure of the UBA domains is highly conserved and that the

Solution Structures of UBA Domains

UBA domains exhibit a conserved hydrophobic patch on the solvent-accessible surface, suggesting a common binding interface. This is the likely binding interface for ubiquitin, and suggests a common regulatory function for UBA domains in the ubiquitin/proteasome pathway. The functional consequence of the UBA –ubiquitin interaction might be to inhibit polyubiquitin chain formation,22,31 thereby stabilizing proteins, as shown for pds1 – 128,31 which is otherwise degraded rapidly by the proteasome. We have shown that the binding of HIV-1 Vpr to UBA(2) is determined by the amino acid composition of the hydrophobic loop 1 and is more specific than the binding of ubiquitin to UBA domains. The binding epitope for HIV-1 Vpr is centered around Pro333 and therefore is not identical with the proposed binding interface with ubiquitin. However, since both epitopes are in close proximity, they might partially overlap, suggesting that each would interfere with the binding of the other.

Materials and Methods Sample preparation The gene encoding residues 155– 204 for the UBA(1) domain of HHR23A was amplified using PCR from a human QuickClone library (Clontech). The cDNA was cloned into the pGEX-2T expression vector (Pharmacia) using the Bam HI and Eco RI restriction sites. The plasmid was then transferred into the E. coli strain Bl21(DE3)Star (Invitrogen). The correctness of the cDNA was confirmed by DNA sequencing using the Dyedeoxy-Terminator method (Perkin – Elmer). A typical protein purification involved growing transformed cells at 37 8C until the absorbance at 550 nm reached 0.6 to 0.7, induction of overexpression was achieved by adding IPTG to a final concentration of 1 mM. The temperature was lowered to 30 8C during expression to maximize the yield of soluble protein. The cells were lysed using BugBuster/Benzonase (Novagen) and the supernatant was applied to a glutathione – Sepharose 4B column. Fusion protein was dialyzed against 20 mM Tris (pH 8.0), 150 mM sodium chloride, 2.5 mM calcium chloride and subsequently cleaved using biotinylated thrombin (Novagen) at a ratio of 50 units per 100 A280 nm. UBA(1) protein was then separated by a second glutathione – Sepharose 4B chromatography step. Thrombin was removed by quenching activity with protease inhibitor cocktail (Calbiochem) and binding it to avidin-agarose. The final purification step was performed using either gel-filtration on a Sephacryl 200 column or by reversed phase chromatography using RPC-30 (Pharmacia). Pooled fractions containing pure UBA(1) were dialyzed against 50 mM sodium phosphate (pH 6.5), 100 mM sodium chloride, 0.1% (w/v) sodium azide and concentrated for NMR using ultrafiltration. The final UBA(1) protein comprises residues Ser155-Gly204 plus a Gly residue at the N terminus resulting from the thrombin cleavage site. Protein homogeneity and purity were checked by SDSPAGE. Uniformly 15N-labeled and 13C,15N-labeled UBA(1) were obtained by growing the bacteria in M9 minimal medium containing 0.5 g/l of 15N-labeled ammonium

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Solution Structures of UBA Domains

chloride or 0.5 g/l of 15N-labeled ammonium chloride and 4 g/l of [13C6]glucose (Isotec). Typical yields were about 10 mg per liter of culture for final purified protein from rich medium (LB) to 7 mg per liter from M9 medium. NMR samples of UBA(1) were prepared by dialyzing the protein after the final gel-filtration step into the NMR buffer and concentrating the protein solution, using ultrafiltration, to 1 to 3 mM.

NMR experiments and structure calculations All NMR experiments were performed at 27 8C on a Bruker DRX-500 or Bruker DRX-600 instrument equipped with a triple-resonance/triple-axis gradient probe. Quadrature detection in the indirect dimensions of three-dimensional experiments was achieved by STATES-time-proportional phase incementation (TPPI) phase cycling. For two-dimensional NOESY experiments, TPPI was used for quadrature detection. All NMR experiments were processed using XWINNMR version 2.5 (Bruker). For 3D experiments, linear prediction was usually applied in both indirect dimensions to double the size of the acquired points. Spectra were transformed using a Gaussian-to-Lorentzian window function for the time domain and a 608 shifted, squared sine bell for the indirect dimensions. All 1H dimensions were referenced to external 3,3,3-trimethylsilylpropionate (TSP). 13C and 15N dimensions were calibrated using the gyromagnetic ratios 15N/1H ¼ 0.101329118 and 13C/1H ¼ 0.251449530.32 All data analysis, i.e. peak picking and integration, was performed using the software AURELIA2.8 (Bruker). For the assignment of the backbone atom chemical shifts, three triple-resonance experiments, CBCA(CO)NH, CBCANH and HBHA(CO)NH, were acquired.20 Sequential assignment was achieved by striping the 3D spectra (based on a chemical shift list derived from a 2D 15N– 1H HSQC) and searching for sequential pairs in the CBCA(CO)NH and CBCANH derived 2D strips using AURELIA’s database search tool. The side-chain assignments were then obtained from 3D HCCH-COSY and HCCH-TOCSY spectra starting from the chemical shift information already derived from the triple-resonance experiments. NOE for distance restraints were assigned using a 13C-edited 3D NOESYHSQC (tmix ¼ 300 ms) of UBA(1) in 95% H2O/5% 2H2O using presaturation for water signal suppression. Additional restraints were obtained from two 15N-edited 3D experiments, e.g. a 15N HSQC-NOESY and a 15NHSQC-NOE-HSQC (tmix ¼ 300 ms). An initial set of NOE restraints was obtained from the 15N-edited and 13 C-edited 3D NOESY spectra. NOE information of ˚ ), these two spectra was classified as weak ($ 5 A ˚ ) or strong ($ 3 A ˚ ) on the basis of their medium ($4 A intensities. Distances derived from NOE signals from 3D experiments were used with large error boundaries ˚ ) plus additional pseudo atom corrections for (^2 A ˚ ) and aromatic NOEs involving methyl groups (þ 1 A ˚ ). Converged structures from strucring systems (þ 2.2 A ture calculations using this limited set of NOE-derived distances (823 NOE derived distances) were then used to interpret the 2D NOESY data. NOE from these experiments (tmix ¼ 80 and 200 ms) were integrated using AURELIA and calibrated using reference distances with known geometry (side-chain amide protons of Asn, aromatic protons in Tyr and Ha to Hb cross-peak in Ala). Lower boundary errors were applied, based on statistical analysis of the reference NOE, and upper

boundary corrections were used according to Wu¨thrich.33 A total of 1692 distance restraints were obtained from all 3D and 2D NOESY data sets. Due to duplicates either from symmetrical peaks in the 2D or 3D data, or due to peaks that were assigned in both the 2D and the 3D data sets, 1055 of the 1692 distance restraints were unique. Structures for UBA(1) were calculated using XPLOR3.1.34 In all, 100 structures were obtained using a simulated annealing protocol employing the all-hydrogen force-field parallhgd.pro version 4.05. NOE-derived distances were treated as point-to-point distances (center) using a soft potential function with a force con˚ 22 (1 cal ¼ 4.184 J).35,36 The simustant of 50 kcal mol21 A lated annealing procedure started from random coordinates and consisted of 30 ps of high-temperature dynamics at 2000 K followed by a slow cooling to 100 K in 30 ps. Then 10% of the calculated structures were selected on the basis of their lowest overall energy (cutoff 370 kcal mol21) and lowest NOE energy (cutoff 180 kcal mol21). For the final minimization, structures that did not exhibit good geometry (bond length and angles, van der Waals contacts) due to the limitations in the parallhdg force-field were subjected to a restrained energy minimization using the Adopted Raphson– Newton algorithm employing the CHARMM24 forcefield37 in the Quanta98 software package (MSI/Biosym). None of the selected structures exhibits NOE violations ˚ and all structures show good covalent greater than 0.5 A geometry and packing (Table 1). The quality of the structures was assessed using the programs PROCHECK38 and Quanta98. Analysis of helix and turn geometry was performed using the program PROMOTIF.39 Protein Data Bank accession code Coordinates for the ten lowest-energy structures have been deposited in the RCSB Protein Data Bank with entry code 1IFY.

Acknowledgments We thank M. Evan Feinstein for manuscript and Figure preparation, and Dr Dara E. Gilbert for acquisition of some of the NMR spectra. This work was supported by NIH grant AI43190 to I. S. Y. Chen and J.F.

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Edited by M. F. Summers (Received 21 December 2001; received in revised form 28 March 2002; accepted 1 April 2002)