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Mardiyyah
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TAPT_Variant_FT

PMCID stringlengths 10 10	Title stringlengths 46 162	FullText stringlengths 17.4k 118k
PMC4802042	A conserved motif in JNK/p38-specific MAPK phosphatases as a determinant for JNK1 recognition and inactivation	Mitogen-activated protein kinases (MAPKs), important in a large array of signalling pathways, are tightly controlled by a cascade of protein kinases and by MAPK phosphatases (MKPs). MAPK signalling efficiency and specificity is modulated by protein–protein interactions between individual MAPKs and the docking motifs in cognate binding partners. Two types of docking interactions have been identified: D-motif-mediated interaction and FXF-docking interaction. Here we report the crystal structure of JNK1 bound to the catalytic domain of MKP7 at 2.4-Å resolution, providing high-resolution structural insight into the FXF-docking interaction. The FNFL segment in MKP7 directly binds to a hydrophobic site on JNK1 that is near the MAPK insertion and helix αG. Biochemical studies further reveal that this highly conserved structural motif is present in all members of the MKP family, and the interaction mode is universal and critical for the MKP-MAPK recognition and biological function.The mitogen-activated protein kinases (MAPKs) are central components of the signal-transduction pathways, which mediate the cellular response to a variety of extracellular stimuli, ranging from growth factors to environmental stresses123. The MAPK signalling pathways are evolutionally highly conserved. The basic assembly of MAPK pathways is a three-tier kinase module that establishes a sequential activation cascade: a MAPK kinase kinase activates a MAPK kinase, which in turn activates a MAPK. The three best-characterized MAPK signalling pathways are mediated by the kinases extracellular signal-regulated kinase (ERK), c-Jun N-terminal kinase (JNK) and p38. The ERK pathway is activated by various mitogens and phorbol esters, whereas the JNK and p38 pathways are stimulated mainly by environmental stress and inflammatory cytokines456. The MAPKs are activated by MAPK kinases that phosphorylate the MAPKs at conserved threonine and tyrosine residues within their activation loop. After activation, each MAPK phosphorylates a distinct set of protein substrates, which act as the critical effectors that enable cells to mount the appropriate responses to varied stimuli. MAPKs lie at the bottom of conserved three-component phosphorylation cascades and utilize docking interactions to link module components and bind substrates78. Two types of docking motifs have been identified in MAPK substrates and cognate proteins: kinase-interacting motif (D-motif) and FXF-motif (also called DEF motif, docking site for ERK FXF). The best-studied docking interactions are those between MAP kinases and ‘D-motifs', which consists of two or more basic residues followed by a short linker and a cluster of hydrophobic residues. The D-motif-docking site (D-site) in MAPKs is situated in a noncatalytic region opposite of the kinase catalytic pocket and is comprised of a highly acidic patch and a hydrophobic groove. D-motifs are found in many MAPK-interacting proteins, including substrates, activating kinases and inactivating phosphatases, as well as scaffolding proteins. A second docking motif for MAPKs consists of two Phe residues separated by one residue (FXF-motif). This motif has been observed in several MAPK substrates910111213. The FXF-motif-binding site of ERK2 has been mapped to a hydrophobic pocket formed between the P+1 site, αG helix and the MAPK insert14. However, the generality and mechanism of the FXF-mediated interaction is unclear. The physiological outcome of MAPK signalling depends on both the magnitude and the duration of kinase activation15. Downregulation of MAPK activity can be achieved through direct dephosphorylation of the phospho-threonine and/or tyrosine residues by various serine/threonine phosphatases, tyrosine phosphatases and dual-specificity phosphatases (DUSPs) termed MKPs. MKPs constitute a group of DUSPs that are characterized by their ability to dephosphorylate both phosphotyrosine and phosphoserine/phospho-threonine residues within a substrate1617. Dysregulated expression of MKPs has been associated with pathogenesis of various diseases, and understanding their precise recognition mechanism presents an important challenge and opportunity for drug development1819. Here, we present the crystal structure of JNK1 in complex with the catalytic domain of MKP7. This structure reveals the molecular mechanism underlying the docking interaction between MKP7 and JNK1. In the JNK1–MKP7 complex, a hydrophobic motif (FNFL) that initiates the helix α5 in the MKP7 catalytic domain directly binds to the FXF-motif-binding site on JNK1, providing the structural insight into the classic FXF-type docking interaction. Biochemical and modelling studies further demonstrate that the molecular interactions mediate this key element for substrate recognition are highly conserved among all MKP-family members. Thus, our study reveals a hitherto unrecognized interaction mode for encoding complex target specificity among MAPK isoforms. DUSPs belong to the protein-tyrosine phosphatases (PTPase) superfamily, which is defined by the PTPase-signature motif CXXGXXR20. MKPs represent a distinct subfamily within a larger group of DUSPs. In mammalian cells, the MKP subfamily includes 10 distinct catalytically active MKPs. All MKPs contain a highly conserved C-terminal catalytic domain (CD) and an N-terminal kinase-binding domain (KBD)1521. The KBD is homologous to the rhodanese family and contains an intervening cluster of basic amino acids, which has been suggested to be important for interacting with the target MAPKs. On the basis of sequence similarity, substrate specificity and predominant subcellular localization, the MKP family can be further divided into three groups (Fig. 1). Biochemical and structural studies have revealed that the KBD of MKPs is critical for MKP3 docking to ERK2, and MKP5 binding to p38α, although their binding mechanisms are completely different2223. However, it is unknown if other MAPKs can interact with the KBD of their cognate phosphatases in the same manner as observed for recognition of ERK2 and p38α by their MKPs, or whether they recognize distinct docking motifs of MKPs. MKP7, the biggest molecule in the MKP family, selectively inactivates JNK and p38 following stress activation24. In addition to the CD and KBD, MKP7 has a long C-terminal region that contains both nuclear localization and export sequences by which MKP7 shuttles between the nucleus and the cytoplasm (Fig. 2a). To quantitatively assess the contribution of the N-terminal domain to the MKP7-catalysed JNK1 dephosphorylation, we first measured the kinetic parameters of the C-terminal truncation of MKP7 (MKP7ΔC304, residues 5–303) and MKP7-CD (residues 156–301) towards phosphorylated JNK1 (pJNK1). Figure 2b shows the variation of initial rates of the MKP7ΔC304 and MKP7-CD-catalysed reaction with the concentration of phospho-JNK1. Because the concentrations of MKP7 and pJNK1 were comparable in the reaction, the assumption that the free-substrate concentration is equal to the total substrate concentration is not valid. Thus, the kinetic data were analysed using the general initial velocity equation, taking substrate depletion into account: The kcat and Km of the MKP7-CD (0.028 s and 0.26 μM) so determined were nearly identical to those of MKP7ΔC304 (0.029 s and 0.27 μM), indicating that the MKP7-KBD has no effect on enzyme catalysis. We next examined the interaction of JNK1 with the CD and KBD of MKP7 by gel filtration analysis. When 3 molar equivalents of CD were mixed with 1 molar equivalent of JNK1, a significant amount of CD co-migrated with JNK1 to earlier fractions, and the excess amount of CD was eluted from the size exclusion column as a monomer, indicating stable complex formation (Fig. 2c). In contrast, no KBD–JNK1 complex was detected when 3 molar equivalents of KBD were mixed with 1 molar equivalent of JNK1. To further confirm the JNK1–MKP7-CD interaction, we performed a pull-down assay using the purified proteins. As shown in Fig. 2d, the CD of MKP7 can be pulled down by JNK1, while the KBD failed to bind to the counterpart protein. Taken together, our data indicate that the CD of MKP7, but not the KBD domain, is responsible for JNK substrate-binding and enzymatic specificity. To understand the molecular basis of JNK1 recognition by MKP7, we determined the crystal structure of unphosphorylated JNK1 in complex with the MKP7-CD (Fig. 3a, Supplementary Fig. 1a and Table 1). In the complex, JNK1 has its characteristic bilobal structure comprising an N-terminal lobe rich in β-sheet and a C-terminal lobe that is mostly α-helical. The overall folding of MKP7-CD is typical of DUSPs, with a central twisted five-stranded β-sheet surrounded by six α-helices. One side of the β-sheet is covered with two α-helices and the other is covered with four α-helices (Fig. 3b). The catalytic domain of MKP7 interacts with JNK1 through a contiguous surface area that is remote from the active site. MKP7-CD is positioned onto the JNK1 molecule so that the active site of the phosphatase faces towards the activation segment. In an alignment of the structure of MKP7-CD with that of VHR25, an atypical ‘MKP' consisting of only a catalytic domain, 119 of 147 MKP7-CD residues could be superimposed with a r.m.s.d. (root mean squared deviation) of 1.05 Å (Fig. 3c). The most striking difference is that helix α0 and loop α0–β1 of VHR are absent in MKP7-CD. Another region that cannot be aligned with VHR is found in loop β3–β4. This loop is shortened by nine residues in MKP7-CD compared with that in VHR. Since helix α0 and the following loop α0–β1 are known for a substrate-recognition motif of VHR and other phosphatases, the absence of these moieties implicates a different substrate-binding mode of MKP7. The active site of MKP7 consists of the phosphate-binding loop (P-loop, Cys244-Leu245-Ala246-Gly247-Ile248-Ser249-Arg250), and Asp213 in the general acid loop (Fig. 3b and Supplementary Fig. 1b). The MKP7-CD structure near the active site exhibits a typical active conformation as found in VHR and other PTPs25. The catalytic residue, Cys244, is located just after strand β5 and optimally positioned for nucleophilic attack262728. Asp213 in MKP7 also adopts a position similar to that of Asp92 in VHR (Supplementary Fig. 1c), indicating that Asp213 is likely to function as the general acid in MKP7. We also observed the binding of a chloride ion in the active site of MKP7-CD. It is located 3.36 Å from the Cys244 side chain and makes electrostatic interactions with the dipole moment of helix α3 and with several main-chain amide groups. The side chain of strictly conserved Arg250 is oriented towards the negatively charged chloride, similar to the canonical phosphate-coordinating conformation. Thus this chloride ion is a mimic for the phosphate group of the substrate, as revealed by a comparison with the structure of PTP1B in complex with phosphotyrosine29 (Supplementary Fig. 1d). Although the catalytically important residues in MKP7-CD are well aligned with those in VHR, the residues in the P-loop of MKP7 are smaller and have a more hydrophobic character than those of VHR (Cys124-Arg125-Glu126-Gly127-Tyr128-Gly129-Arg130; Fig. 3b,c). The difference in the polarity/hydrophobicity of the surface may also point to the origin of the differences in the substrate-recognition mechanism for these two phosphatases (Supplementary Fig. 1e,f). In the complex, MKP7-CD and JNK1 form extensive protein–protein interactions involving the C-terminal helices of MKP7-CD and C-lobe of JNK1 (Fig. 3d,e). As a result, the buried solvent-accessible surface area is ∼1,315 Å. In the C-terminal domain, JNK1 has an insertion after the helix αG. This insertion consists of two helices (α1L14 and α2L14) that are common to all members of the MAPK family. The interactive surface in JNK1, formed by the helices αG and α2L14, displays a hydrophobic region, centred at Trp234 (Fig. 3d). The MKP7-docking region includes two helices, α4 and α5, and the general acid loop. The aromatic ring of Phe285 on MKP7 α5-helix is nestled in a hydrophobic pocket on JNK1, formed by side chains of Ile197, Leu198, Ile231, Trp234, Val256, Tyr259, Val260 and the aliphatic portion of His230 (Fig. 3d,f and Supplementary Fig. 1g). In addition, there are hydrogen bonds between Ser282 and Asn286 of MKP7 and His230 and Thr255 of JNK1, and the main chain of Phe215 in the general acid loop of MKP7 is hydrogen-bonded to the side chain of Gln253 in JNK1. The second interactive area involves the α4 helix of MKP7 and charged/polar residues of JNK1 (Fig. 3e). The carboxylate of Asp268 in MKP7 forms a salt bridge with side chain of Arg263 in JNK1, and Lys275 of MKP7 forms a hydrogen bond and a salt bridge with Thr228 and Asp229 of JNK1, respectively. To assess the importance of the aforementioned interactions, we generated a series of point mutations on the MKP7-CD and examined their effect on the MKP7-catalysed JNK1 dephosphorylation (Fig. 4a). When the hydrophobic residues Phe285 and Phe287 on the α5 of MKP7-CD were replaced by Asp or Ala, their phosphatase activities for JNK1 dephosphorylation decreased ∼10-fold. In comparison, replacement of the other residues (Phe215, Asp268, Lys275, Ser282, Asn286 and Leu292) with an Ala or Asp individually led to a modest decrease in catalytic efficiencies, suggesting that this position may only affect some selectivity of MKP. Mutation of Leu288 markedly reduced its solubility when expressed in Escherichia coli, resulting in the insoluble aggregation of the mutant protein. Gel filtration analysis further confirmed the key role of Phe285 in the MKP7–JNK1 interaction: no F285D–JNK1 complex was detected when 3 molar equivalents of MKP7-CD (F285D) were mixed with 1 molar equivalent of JNK1 (Fig. 4b). Interestingly, mutation of Phe287 results in a considerable loss of activity against pJNK1 without altering the affinity of MKP7-CD for JNK1 (Supplementary Fig. 2a). We also generated a series of point mutations in the JNK1 and assessed the effect on JNK1 binding using the GST pull-down assay (Fig. 4c). Substitution at Asp229, Trp234, Thr255, Val256, Tyr259 and Val260 significantly reduced the binding affinity of MKP7-CD for JNK. To determine whether the deficiencies in their abilities to bind partner proteins or carry out catalytic function are owing to misfolding of the purified mutant proteins, we also examined the folding properties of the JNK1 and MKP7 mutants with circular dichroism. The spectra of these mutants are similar to the wild-type proteins, indicating that these mutants fold as well as the wild-type proteins (Fig. 4d,e). Taken together, these results are consistent with the present crystallographic model, which reveal the hydrophobic contacts between the MKP7 catalytic domain and JNK1 have a predominant role in the enzyme–substrate interaction, and hydrophobic residue Phe285 in the MKP7-CD is a key residue for its high-affinity binding to JNK1. It has previously been reported that several cytosolic and inducible nuclear MKPs undergo catalytic activation upon interaction with the MAPK substrates15. This allosteric activation of MKP3 has been well-documented in vitro using pNPP, a small-molecule phosphotyrosine analogue of its normal substrate3031. We then assayed pNPPase activities of MKP7ΔC304 and MKP7-CD in the presence of JNK1. Incubation of MKP7 with JNK1 did not markedly stimulate the phosphatase activity, which is consistent with previous results that MKP7 solely possesses the intrinsic activity (Supplementary Fig. 2b). The small pNPP molecule binds directly at the enzyme active site and can be used to probe the reaction mechanism of protein phosphatases. We therefore examined the effects of the MKP7-CD mutants on their pNPPase activities. As shown in Fig. 4f, all the mutants, except F287D/A, showed little or no activity change compared with the wild-type MKP7-CD. In the JNK1/MKP7-CD complex structure, Phe287 of MKP7 does not make contacts with JNK1 substrate. It penetrates into a pocket formed by residues from the P-loop and general acid loop and forms hydrophobic contacts with the aliphatic portions of side chains of Arg250, Glu217 and Ile219, suggesting that Phe287 in MKP7 would play a similar role to that of its structural counterpart in the PTPs (Gln266 in PTP1B) and VHR (Phe166 in VHR) in the precise alignment of active-site residues in MKP7 with respect to the substrate for efficient catalysis32333435 (Supplementary Fig. 2c). Kinase-associated phosphatase (KAP), a member of the DUSP family, plays a crucial role in cell cycle regulation by dephosphorylating the pThr160 residue of CDK2 (cyclin-dependent kinase 2). The crystal structure of the CDK2/KAP complex has been determined at 3.0 Å (Fig. 5a)36. The interface between these two proteins consists of three discontinuous contact regions. Biochemical results suggested that the affinity and specificity between KAP and CDK2 results from the recognition site comprising CDK2 residues from the αG helix and L14 loop and the N-terminal helical region of KAP (Fig. 5b). There is a hydrogen bond between the main-chain nitrogen of Ile183 (KAP) and side chain oxygen of Glu208 (CDK2), and salt bridges between Lys184 of KAP and Asp235 of CDK2. Structural analysis and sequence alignment reveal that one of the few differences between MKP7-CD and KAP in the substrate-binding region is the presence of the motif FNFL in MKP7-CD, which corresponds to IKQY in KAP (Fig. 5c). The substitution of the two hydrophobic residues with charged/polar residues (F285I/N286K) seriously disrupts the hydrophobic interaction required for MKP7 binding on JNK1 (Fig. 4a). In addition, His230 and Val256 in JNK1 are replaced by the negatively charged residues Glu208 and Asp235 in CDK2 (Fig. 5d), and the charge distribution on the CDK2 interactive surface is quite different from that of JNK. These data indicated that a unique hydrophobic pocket formed between the MAPK insert and αG helix plays a major role in the substrate recognition by MKPs. JNK is activated following cellular exposure to a number of acute stimuli such as anisomycin, H2O2, ultraviolet light, sorbitol, DNA-damaging agents and several strong apoptosis inducers (etoposide, cisplatin and taxol)373839. To assess the effects of MKP7 and its mutants on the activation of endogenous JNK in vivo, HEK293T cells were transfected with blank vector or with HA-tagged constructs for full-length MKP7, MKP7ΔC304 and MKP7-CD or MKP7 mutants, and stimulated with ultraviolet or etoposide treatment. As shown in Fig. 6a–c, immunobloting showed similar expression levels for the different MKP7 constructs in all the cells. Overexpressed full-length MKP7, MKP7ΔC304 and MKP7-CD significantly reduced the endogenous level of phosphorylated JNK compared with vector-transfected cells. Parallel experiments showed clearly that the D-motif mutants (R56A/R57A and V63A/I65A) dephosphorylated JNK as did the wild type under the same conditions, further confirming that the MKP7-KBD is not required for the JNK inactivation in vivo. Consistent with the in vitro data, the level of phosphorylated JNK was not or little altered in MKP7 FXF-motif mutants (F285D, F287D and L288D)-transfected cells, and the MKP7 D268A and N286A mutants retained the ability to reduce the phosphorylation levels of JNK. We next tested in vivo interactions between JNK1 mutants and full-length MKP7 by coimmunoprecipitation experiments under unstimulated conditions. When co-expressed in HEK293T cells, wild-type (HA)-JNK1 was readily precipitated with (Myc)-MKP7 (Fig. 6d), indicating that MKP7 binds dephosphorylated JNK1 protein in vivo. In agreement with the in vitro pull-down results, the mutants D229A, W234D and Y259D were not co-precipitated with MKP7, and the I231D mutant had only little effect on the JNK1–MKP7 interaction (Fig. 6d and Supplementary Fig. 3a). Activation of the JNK signalling pathway is frequently associated with apoptotic cell death, and inhibition of JNK can prevent apoptotic death of multiple cells640414243. To examine whether the inhibition of JNK activity by MKP7 would provide protections against the apoptosis, we analysed the rate of apoptosis in ultraviolet-irradiated cells transfected with MKP7 (wild type or mutants) by flow cytometry. The results showed similar apoptotic rates between cells transfected with blank vector or with MKP7 (wild type or mutants) under unstimulated conditions (Supplementary Fig. 3b), while ultraviolet-irradiation significantly increased apoptotic rate in cells transfected with blank vector (Fig. 6e). Expressions of wild-type MKP7, MKP7ΔC304 and MKP7-CD significantly decreased the proportion of apoptotic cells after ultraviolet treatment. Moreover, treatment of cells expressing MKP7-KBD mutants (R56A/R57A and V63A/I65A) decreased the apoptosis rates to a similar extent as MKP7 wild type did. In contrast, cells transfected with the MKP7 FXF-motif mutants (F285D, F287D and L288D) showed little protective effect after ultraviolet treatment and similar levels of apoptosis rates were detected to cells transfected with control vectors (Fig. 6e,f). Taken together, our results suggested that FXF-motif-mediated, rather than KBD-mediated, interaction is essential for MKP7 to block ultraviolet-induced apoptosis. MKP5 belongs to the same subfamily as MKP7. MKP5 is unique among the MKPs in possessing an additional domain of unknown function at the N-terminus44 (Fig. 7a). The KBD of MKP5 interacts with the D-site of p38α to mediate the enzyme–substrate interaction. Deletion of the KBD in MKP5 leads to a 280-fold increase in Km for p38α substrate23. In contrast to p38α substrate, deletion of the MKP5-KBD had little effects on the kinetic parameters for the JNK1 dephosphorylation, indicating that the KBD of MKP5 is not required for the JNK1 dephosphorylation (Fig. 7b). The substrate specificity constant kcat /Km value for MKP5-CD was calculated as 1.0 × 10 M s, which is very close to that of MKP7-CD (1.07 × 10 M s). The crystal structure of human MKP5-CD has been determined45. Comparisons between catalytic domains structures of MKP5 and MKP7 reveal that the overall folds of the two proteins are highly similar, with only a few regions exhibiting small deviations (r.m.s.d. of 0.79 Å; Fig. 7c). Given the distinct interaction mode revealed by the crystal structure of JNK1–MKP7-CD, one obvious question is whether this is a general mechanism used by all members of the JNK-specific MKPs. To address this issue, we first examined the docking ability of JNK1 to the KBD and CD of MKP5 using gel filtration analysis and pull-down assays. It can be seen from gel filtration experiments that JNK1 can forms a stable heterodimer with MKP5-CD in solution, but no detectable interaction was found with the KBD domain (Fig. 7d). Pull-down assays also confirmed the protein–protein interactions observed above. The catalytic domain of MKP5, but not its KBD, was able to pull-down a detectable amount of JNK1 (Fig. 7e), implicating a different substrate-recognition mechanisms for p38 and JNK MAPKs. To further test our hypothesis, we generated forms of MKP5-CD bearing mutations corresponding to the changes we made on MKP7-CD on the basis of sequence and structural alignment and examined their effects on the phosphatase activity. As shown in Fig. 7f, the T432A and L449F MKP5 mutant showed little or no difference in phosphatase activity, whereas the other mutants showed reduced specific activities of MKP5. As in the case of MKP7, all the mutants, except F451D/A, showed no pNPPase activity changes compared with the wild-type MKP5-CD (Fig. 7g), and the point mutations in JNK1 also reduced the binding affinity of MKP5-CD for JNK1 (Fig. 7h). In addition, there were no significant differences in the CD spectra between wild-type and mutant proteins, indicating that the overall structures of these mutants did not change significantly from that of wild-type MKP5 protein (Supplementary Fig. 4a). Taken together, our results suggest that MKP5 binds JNK1 in a docking mode similar to that in the JNK1–MKP7 complex, and the detailed interaction model can be generated using molecular dynamics simulation based on the structure of JNK1–MKP7-CD complex (Supplementary Fig. 4b,c). In this model, the MKP5-CD adopts a conformation nearly identical to that in its unbound form, suggesting that the conformation of the catalytic domain undergoes little change, if any at all, upon JNK1 binding. In particular, Leu449 of MKP5, which is equivalent to the key residue Phe285 of MKP7, buried deeply within the hydrophobic pocket of JNK1 in the same way as Phe285 in the JNK1–MKP7-CD complex (Supplementary Fig. 4d). Despite the strong similarities between JNK1–MKP5-CD and JNK1–MKP7-CD, however, there are differences. The JNK1–MKP7-CD interaction is better and more extensive. Asp268 of MKP7-CD forms salt bridge with JNK1 Arg263, whereas the corresponding residue Thr432 in MKP5-CD may not interact with JNK1. In addition, the key interacting residues of MKP7-CD, Phe215, Leu267 and Leu288, are replaced by less hydrophobic residues, Asn379, Met431 and Met452 in MKP5-CD (Fig. 5c), respectively, which may result in weaker hydrophobic interactions between MKP5-CD and JNK1. This is consistent with the experimental observation showing that JNK1 binds to MKP7-CD much more tightly than MKP5-CD (Km value of MKP5-CD for pJNK1 substrate is ∼20-fold higher than that of MKP7-CD). The MAPKs p38, ERK and JNK, are central to evolutionarily conserved signalling pathways that are present in all eukaryotic cells. Each MAPK cascade is activated in response to a diverse array of extracellular signals and culminates in the dual-phosphorylation of a threonine and a tyrosine residue in the MAPK-activation loop2. The propagation of MAPK signals is attenuated through the actions of the MKPs. Most studies have focused on the dephosphorylation of MAPKs by phosphatases containing the ‘kinase-interaction motif ' (D-motif), including a group of DUSPs (MKPs) and a distinct subfamily of tyrosine phosphatases (HePTP, STEP and PTP-SL)4647. Crystal structures of ERK2 bound with the D-motif sequences derived from MKP3 and HePTP have been reported2248. These structures revealed that linear docking motifs in interacting proteins bind to a common docking site on MAPKs outside the kinase active site. The particular amino acids and their spacing within D-motif sequences and amino acid composition of the docking sites on MAPKs appear to determine the specificity of D-motifs for individual MAPKs. Recently, the crystal structure of a complex between the KBD of MKP5 and p38α has been obtained23. This complex has revealed a distinct interaction mode for MKP5. The KBD of MKP5 binds to p38α in the opposite polypeptide direction compared with how the D-motif of MKP3 binds to ERK2. In contrast to the canonical D-motif-binding mode, separate helices, α2 and α3′, in the KBD of MKP5 engage the p38α-docking site. Further structural and biochemical studies indicate that KBD of MKP7 may interact with p38α in a similar manner to that of MKP5. In contrast to MKP5, removal of the KBD domain from MKP7 does not drastically affect enzyme catalysis, and the kinetic parameters of MKP7-CD for p38α substrate are very similar to those for JNK1 substrate23. Taken together, these results suggest that MKP7 utilizes a bipartite recognition mechanism to achieve the efficiency and fidelity of p38α signalling. The MKP7-KBD docks to the D-site located on the back side of the p38α catalytic pocket for high-affinity association, whereas the interaction of the MKP7-CD with another p38α structural region, which is close to the activation loop, may not only stabilize binding but also provide contacts crucial for organizing the MKP7 active site with respect to the phosphoreceptor in the p38α substrate for efficient dephosphorylation. In addition to the canonical D-site, the MAPK ERK2 contains a second binding site utilized by transcription factor substrates and phosphatases, the FXF-motif-binding site (also called F-site), that is exposed in active ERK2 and the D-motif peptide-induced conformation of MAPKs91049. This hydrophobic site was first identified by changes in deuterium exchange profiles, and is near the MAPK insertion and helix αG. Interestingly, many of the equivalent residues in JNK1, important for MKP7-CD recognition, are also used for substrate binding by ERK2 (ref. 14), indicating that this site is overlapped with the DEF-site previously identified in ERK2 (Fig. 5d). MKP3 is highly specific in dephosphorylating and inactivating ERK2, and the phosphatase activity of the MKP3-catalysed pNPP reaction can be markedly increased in the presence of ERK2 (refs 30, 31). Sequence alignment of all MKPs reveals a high degree of conservation of residues surrounding the interacting region observed in JNK1–MKP7-CD complex (Supplementary Fig. 5). Therefore, it is tempting to speculate that the catalytic domain of MKP3 may bind to ERK2 in a manner analogous to the way by which MKP7-CD binds to JNK1. A comprehensive examination of the molecular basis of the specific ERK2 recognition by MKP3 is underway. The ongoing work demonstrates that although the overall interaction modes are similar between the JNK1–MKP7-CD and ERK2–MKP3-CD complexes, the ERK2–MKP3-CD interaction is less extensive and helix α4 from MKP3-CD does not interact directly with ERK2. The FXF-motif-mediated interaction is critical for both pERK2 inactivation and ERK2-induced MKP3 activation (manuscript in preparation). In summary, we have resolved the structure of JNK1 in complex with the catalytic domain of MKP7. This structure reveals an FXF-docking interaction mode between MAPK and MKP. Results from biochemical characterization of the Phe285 and Phe287 MKP7 mutants combined with structural information support the conclusion that the two Phe residues serve different roles in the catalytic reaction. Phe285 is essential for JNK1 substrate binding, whereas Phe287 plays a role for the precise alignment of active-site residues, which are important for transition-state stabilization32. This newly identified FXF-type motif is present in all MKPs, except that the residue at the first position in MKP5 is an equivalent hydrophobic leucine residue (see also Fig. 7f,g), suggesting that these two Phe residues would play a similar role in facilitating substrate recognition and catalysis, respectively. An important feature of MKP–JNK1 interactions is that MKP7 or MKP5 only interact with the F-site of JNK1. One possible explanation is that JNK1 needs to use the D-site to interact with JIP-1, a scaffold protein for JNK signalling5051. The N-terminal JNK-binding domain of JIP-1 interacts with the D-site on JNK and this interaction is required for JIP-1-mediated enhancement of JNK activation5253. In addition, JIP-1 can also associate with MKP7 via the C-terminal region of MKP7 (ref. 54). When MKP7 is bound to JIP-1, it reduces JNK activation, leading to reduced phosphorylation of the JNK target c-Jun. Thus, our biochemical and structural data allow us to present a model for the JNK1–JIP-1–MKP7 ternary complex and provide an important insight into the assembly and function of JNK signalling modules (Supplementary Fig. 6). The cDNAs of human MKP7 and MKP5 were kindly provided by Dr Mathijs Baens (University of Leuven) and Dr Eisuke Nishida (Kyoto University), respectively. The cDNAs of human ASK1, MKK4, MKK7 and JNK1 were kindly provided by Dr Zhenguo Wu (Hong Kong University of Science and Technology). The catalytic domains of MKP7 (MKP7-CD, residues 156–301) and MKP5 (MKP5-CD, 320–467) and the full-length MKP5 were cloned into the pET15b vector, resulting in the N-terminal His-fusion proteins. The KBD domains of MKP7 (MKP7-KBD, 5–138) and MKP5 (MKP5-KBD, 139–287), and the C-terminal truncation of MKP7 (MKP7ΔC304, 5–303) were cloned into pET21b vector for generation of C-terminal His-tagged proteins. The human full-length JNK1, MKK4, MKK7 and the kinase domain of ASK1 (659–951) were cloned into pGEX4T-2, pET15b and/or pET21b vectors to produce a GST- or His-tagged protein. Mutations of MKP7-CD, MKP5-CD and JNK1 were generated by overlap PCR procedure. All constructs were verified by DNA sequencing. All proteins, overexpressed in BL21(DE3) cells at 20 °C, were first purified over Ni-NTA (Qiagen) or GS4B (GE Healthcare) columns, and then by ion exchange and gel filtration chromatography (Source-15Q/15S and Superdex-200, GE Healthcare) at 4 °C. The double-phosphorylated JNK1 (phospho-JNK1) was generated by mixing JNK1 with upstream kinases MKK4, MKK7 and ASK1 in buffer containing 10 mM MgCl2 and 2 mM ATP, and further purified by gel filtration chromatography (Superdex-200, GE Healthcare) at 4 °C (ref. 55). Proteins were stored at −80 °C, and stocks for phosphatase assays were supplemented with glycerol to a final concentration of 20% (v/v). Protein concentrations were determined spectrophotometrically using theoretical molar extinction coefficients at 280 nm (ref. 56). The mixture of unphosphorylated JNK1 and MKP7-CD at 1:1 molar ratio was subjected to crystallization trials. Crystals were grown by the vapor-diffusion technique in hanging drops, and the drops were prepared by mixing equal volumes of protein with reservoir solution containing 0.1 M HEPES, pH 7.0, 14% PEG3350, 0.2 M MgCl2, 6% 1,6-Hexanediol and 0.005 M EDTA at 21 °C. Crystals were cryo-protected in reservoir solutions supplemented with 10% glycerol and then flash frozen in liquid nitrogen. The diffraction data sets were collected at beamline 17U at Shanghai Synchrotron Radiation Facility and processed with the HKL2000 package57. The crystals belong to space group P1 and comprise eight molecules per asymmetric unit (four complexes). Structure was solved by molecular replacement using Phaser58 with JNK1 (PDB 1UKH) and MKP5-CD (PDB 1ZZW) as the search models. Standard refinement was performed with programs PHENIX59 and Coot60. The crystal structure of unphosphorylated JNK1 in complex with the catalytic domain of MKP7 was refined to 2.4 Å resolution. Initial structural refinement was performed with NCS restraints, and after several rounds the restraints were removed from the calculations. The final Rwork and Rfree were 21.7 and 23.9%, respectively. The crystallographic asymmetric unit contains four JNK1–MKP7-CD complexes. The four complexes are nearly identical with an r.m.s.d.<1 Å for any complex pair in the asymmetric unit. Ramachandran analysis was carried out using PROCHECK61. Additional density at the active site of MKP7-CD was attributed to a chloride ion incorporated as a crystallizing agent, similar to those observed in the structures of MKP3-CD and MKP5-CD (refs 62, 63). The data collection and refinement statistics are summarized in Table 1. All structural representations in this paper were prepared with PYMOL (http://www.pymol.org). The activities of MKP7 and MKP5 was assayed using phospho-JNK1 as substrate in the coupled enzyme system containing 50 mM MOPS, pH 7.0, 100 mM NaCl, 0.1 mM EDTA, 50 μM MESG and 0.1 mg ml PNPase. This coupled assay uses PNPase and its chromogenic substrate MESG to monitor the production of inorganic phosphate64. The reactions were initiated by addition of 0.4 μM MKP7-CD and MKP7ΔC304 (or 0.1 μM MKP5 full-length and 0.15 μM MKP5-CD) for substrate phospho-JNK1. All experiments were carried out at 25 °C in 1.8 ml reaction mixtures, and the continuous absorbance changes were recorded with a PerkinElmer LAMBDA 45 spectrophotometer equipped with a magnetic stirrer in the cuvette holder. The quantification of inorganic phosphate produced was monitored at 360 nm with the extinction coefficient of 11,200 M cm (ref. 65). The initial rates were determined from the linear slope of the progress curves obtained. The activity of MKP7-CD or MKP5-CD mutants were assayed using pNPP or phospho-JNK1 as substrate. The phospho-JNK1 assay was performed as the same procedure mentioned above, and in the presence of wild type (as a control) or indicated mutants, and equal concentrations of phospho-JNK1. The pNPP assay was performed in the reaction mixture containg 50 mM MOPS, pH 7.0, 100 mM NaCl, 0.1 mM EDTA and 20 mM pNPP. The amount of the product p-nitrophenol was determined from the absorbance at 405 nm using a molar extinction coefficient of 18,000 M cm (ref. 31). The interactions of JNK1 with the CD and KBD domains of MKP7 and MKP5 were examined by gel filtration analyses using a Superdex-200 10/300 column on an ÄKTA FPLC (GE Healthcare). The column was equilibrated with a buffer containing 10 mM HEPES, pH 7.5, 150 mM NaCl and 2 mM dithiothreitol, and calibrated with molecular mass standards. Samples of individual proteins and indicated mixtures (500 μl each) were loaded to the Superdex-200 column and then eluted at a flow rate of 0.5 ml min. Fractions of 0.5 ml each were collected, and aliquots of relevant fractions were subjected to SDS-polyacrylamide gel electrophoresis (PAGE) followed Coomassie Blue staining. The interactions between various JNK1 mutants and MKP7-CD or MKP5-CD were assessed by GST-mediated pull-down assays at 4 °C. First, 0.5 ml GST-JNK1 proteins (6 μM) were loaded to 0.2 ml GS4B resin. The excess unbound JNK1 or other contaminants were removed by washing the column 5 times, each with 1.0 ml buffer containing 25 mM Tris-HCl, pH 8.0, 150 mM NaCl and 2 mM dithiothreitol. Then, 0.5 ml MKP7-CD or MKP5-CD (20 μM) was allowed to flow through the JNK1-bound column. After extensive washing, the bound proteins were eluted with 0.5 ml reduced glutathione (10 mM). The interactions of JNK1 with the CD and KBD domains of MKP7 and MKP5 were also examined by GST-mediated pull-down assays. The GST protein alone was used as a control. Aliquots of all eluates were subjected to SDS-PAGE, and proteins were visualized by Coomassie Blue staining. The uncropped gels are shown in Supplementary Fig. 7. The model of catalytic domain of MKP5 bound to JNK was constructed by superimposition of previous deposited structures of MKP5-CD (PDB 1ZZW) to the corresponding domains in the crystal structure of JNK1–MKP7-CD. The fractured loops in deposited structures were computationally generated using Modeller66. The program CHARMM22 (ref. 67) was then used to add hydrogen atoms, N- and C-terminal patches to the model. The model was then subjected to restrained energy minimization to optimize bonds and remove any nonbonded steric clashes. Refinement of the modelled complex was performed using NAMD2.9 package68 at 1 atm pressure and 300 K. The generated complex structure was solvated and neutralized in a box with TIP3P water at a minimum of 13 Å between the model and the wall of the box. The simulation was first set up with 1 fs time step under periodic boundary conditions. The particle mesh Ewald method was applied to model the electrostatics and the van der Waals interactions cutoff was set at 12 Å. The system was restrained for 5 ps minimization and 5 ps simulation, and followed by removing all the restraints and performing a minimization of 10 ps and an equilibration of 10 ns. Simulations were viewed using VMD69. The experiments were performed on a Chirascan-plus circular dichroism Spectrometer (Applied Photophysics, Surrey, UK) using 0.1 mm quartz cuvette. The protein sample were analysed at a concentration of 0.5 mg ml. Data were collected over a wavelength range from 260 to 190 nm with 1 nm intervals at room temperature, three scans were averaged, and the baseline spetrum of solution buffer containing 10 mM HEPES (pH 7.5) and 150 mM NaCl was subtracted. pcDNA3.3-Myc-MKP7, pCMV5-3HA-JNK1 and pBOBI-HA-MKP7 were generated with standard molecular techniques. Mutants with amino acid substitution and truncation constructs were generated through PCR-based site-directed mutagenesis method using Pfu polymerase (Stratagene). The authenticities of all constructs were confirmed by sequencing (Invitrogen, China). HEK293T and HeLa cells (ATCC) were maintained in DMEM supplemented with 10% fetal bovine serum, 100 IU penicillin, 100 mg ml streptomycin at 37 °C in a humidified incubator containing 5% CO2. Polyethylenimine (Polysciences, #23966) at a final concentration of 10 μM was used to transfect HEK293T cells. Total DNA for each plate was adjusted to the same amount by adding relevant blank vector. Lentiviruses for infection were packaged in HEK293T cells after transfection using Lipofectamine 2000 (Invitrogen, 11668-027). At 30 h post transfection, medium was collected for further infection. Cells were lysed in a lysis buffer containing 20 mM Tris-HCl (pH 7.4), 150 mM NaCl, 0.5% NP-40, 1 mM EDTA, 2 mM Na3VO4, 25 mM NaF, 1 mM phenylmethanesulfonyl fluoride, 1 μg ml leupeptin and 1 μg ml aprotinin. Cell lysates were incubated with respective antibodies overnight at 4 °C. Protein aggregates resulting from the overnight incubation were removed by centrifugation, and protein A/G beads (Santa-Cruz Biotechnology, Dallas, TX, USA) were then added into the lysates and incubated for another 3 h. After spinning and washing for three times with the lysis buffer, the beads were mixed with 2 × SDS sample buffer, boiled and subjected to 15% SDS/PAGE. The samples were transferred to PVDF membranes (Millipore), and immunoblotted with indicated antibodies. Levels of total proteins and the levels of phosphorylation of proteins were analysed on separate gels. The uncropped blots are shown in Supplementary Fig. 7. Antibodies used in this study: mouse anti-HA (1:100 for immunoprecipitation; F-7) and anti-JNK1 (1:1,000 for immunoblotting; F-3) antibodies were purchased from Santa-Cruz Biotechnology. Anti-c-Myc Agarose Affinity Gel antibody produced in rabbit (1:200 for immunoprecipitation; A7470) was purchased from Sigma. Rabbit anti-HA-tag (1:1,000 for immunoblotting; #3724), anti-phospho-JNK-T183/Y185 (1:1,000 for immunoblotting; #4668) and mouse anti-Myc-tag (1:1,000 for immunoblotting; #2276) antibodies were purchased from Cell Signaling Technology. Etoposide (E1383) was purchased from Sigma. HeLa cells were infected with lentiviruses expressing MKP7 or its mutants. At 36 h post infection, cells were irradiated with 25 J m ultraviolet light and collected at 6 h after irradiation. Cells were then stained with the Annexin-V-APC/PI double-staining solution (BD Biosciences) and analysed with a flow cytometer (BD LSRFortessa). The percentages of apoptotic cells were quantified with FlowJo 7.6.1 software. Accession codes: The coordinates and structure factors have been deposited in the Protein Data Bank with accession codes 4YR8 for the JNK1–MKP7-CD structure. How to cite this article: Liu, X. et al. A conserved motif in JNK/p38-specific MAPK phosphatases as a determinant for JNK1 recognition and inactivation. Nat. Commun. 7:10879 doi: 10.1038/ncomms10879 (2016).
PMC4968113	Structural diversity in a human antibody germline library	To support antibody therapeutic development, the crystal structures of a set of 16 germline variants composed of 4 different kappa light chains paired with 4 different heavy chains have been determined. All four heavy chains of the antigen-binding fragments (Fabs) have the same complementarity-determining region (CDR) H3 that was reported in an earlier Fab structure. The structure analyses include comparisons of the overall structures, canonical structures of the CDRs and the VH:VL packing interactions. The CDR conformations for the most part are tightly clustered, especially for the ones with shorter lengths. The longer CDRs with tandem glycines or serines have more conformational diversity than the others. CDR H3, despite having the same amino acid sequence, exhibits the largest conformational diversity. About half of the structures have CDR H3 conformations similar to that of the parent; the others diverge significantly. One conclusion is that the CDR H3 conformations are influenced by both their amino acid sequence and their structural environment determined by the heavy and light chain pairing. The stem regions of 14 of the variant pairs are in the ‘kinked’ conformation, and only 2 are in the extended conformation. The packing of the VH and VL domains is consistent with our knowledge of antibody structure, and the tilt angles between these domains cover a range of 11 degrees. Two of 16 structures showed particularly large variations in the tilt angles when compared with the other pairings. The structures and their analyses provide a rich foundation for future antibody modeling and engineering efforts.At present, therapeutic antibodies are the largest class of biotherapeutic proteins that are in clinical trials. The use of monoclonal antibodies as therapeutics began in the early 1980s, and their composition has transitioned from murine antibodies to generally less immunogenic humanized and human antibodies. The technologies currently used to obtain human antibodies include transgenic mice containing human antibody repertoires, cloning directly from human B cells, and in vitro selection from antibody libraries using various display technologies. Once a candidate antibody is identified, protein engineering is usually required to produce a molecule with the right biophysical and functional properties. All engineering efforts are guided by our understanding of the atomic structures of antibodies. In such efforts, the crystal structure of the specific antibody may not be available, but modeling can be used to guide the engineering efforts. Today's antibody modeling approaches, which normally focus on the variable region, are being developed by the application of structural principles and insights that are evolving as our knowledge of antibody structures continues to expand. Our current structural knowledge of antibodies is based on a multitude of studies that used many techniques to gain insight into the functional and structural properties of this class of macromolecule. Five different antibody isotypes occur, IgG, IgD, IgE, IgA and IgM, and each isotype has a unique role in the adaptive immune system. IgG, IgD and IgE isotypes are composed of 2 heavy chains (HCs) and 2 light chains (LCs) linked through disulfide bonds, while IgA and IgM are double and quintuple versions of antibodies, respectively. Isotypes IgG, IgD and IgA each have 4 domains, one variable (V) and 3 constant (C) domains, while IgE and IgM each have the same 4 domains along with an additional C domain. These multimeric forms are linked with an additional J chain. The LCs that associate with the HCs are divided into 2 functionally indistinguishable classes, κ and λ. Both κ and λ polypeptide chains are composed of a single V domain and a single C domain. The heavy and light chains are composed of structural domains that have ∼110 amino acid residues. These domains have a common folding pattern often referred to as the “immunoglobulin fold,” formed by the packing together of 2 anti-parallel β-sheets. All immunoglobulin chains have an N-terminal V domain followed by 1 to 4 C domains, depending upon the chain type. In antibodies, the heavy and light chain V domains pack together forming the antigen combining site. This site, which interacts with the antigen (or target), is the focus of current antibody modeling efforts. This interaction site is composed of 6 complementarity-determining regions (CDRs) that were identified in early antibody amino acid sequence analyses to be hypervariable in nature, and thus are responsible for the sequence and structural diversity of our antibody repertoire. The sequence diversity of the CDR regions presents a substantial challenge to antibody modeling. However, an initial structural analysis of the combining sites of the small set of structures of immunoglobulin fragments available in the 1980s found that 5 of the 6 hypervariable loops or CDRs had canonical structures (a limited set of main-chain conformations). A CDR canonical structure is defined by its length and conserved residues located in the hypervariable loop and framework residues (V-region residues that are not part of the CDRs). Furthermore, studies of antibody sequences revealed that the total number of canonical structures are limited for each CDR, indicating possibly that antigen recognition may be affected by structural restrictions at the antigen-binding site. Later studies found that the CDR loop length is the primary determining factor of antigen-binding site topography because it is the primary factor for determining a canonical structure. Additional efforts have led to our current understanding that the LC CDRs L1, L2, and L3 have preferred sets of canonical structures based on length and amino acid sequence composition. This was also found to be the case for the H1 and H2 CDRs. Classification schemes for the canonical structures of these 5 CDRs have emerged and evolved as the number of depositions in the Protein Data Bank of Fab fragments of antibodies grow. Recently, a comprehensive CDR classification scheme was reported identifying 72 clusters of conformations observed in antibody structures. The knowledge and predictability of these CDR canonical structures have greatly advanced antibody modeling efforts. In contrast to CDRs L1, L2, L3, H1 and H2, no canonical structures have been observed for CDR H3, which is the most variable in length and amino acid sequence. Some clustering of conformations was observed for the shortest lengths; however, for the longer loops, only the portions nearest the framework (torso, stem or anchor region) were found to have defined conformations. In the torso region, 2 primary groups could be identified, which led to sequence-based rules that can predict with some degree of reliability the conformation of the stem region. The “kinked” or “bulged” conformation is the most prevalent, but an “extended” or “non-bulged” conformation is also, but less frequently, observed. The cataloging and development of the rules for predicting the conformation of the anchor region of CDR H3 continue to be refined, producing new insight into the CDR H3 conformations and new tools for antibody engineering. Current antibody modeling approaches take advantage of the most recent advances in homology modeling, the evolving understanding of the CDR canonical structures, the emerging rules for CDR H3 modeling and the growing body of antibody structural data available from the PDB. Recent antibody modeling assessments show continued improvement in the quality of the models being generated by a variety of modeling methods. Although antibody modeling is improving, the latest assessment revealed a number of challenges that need to be overcome to provide accurate 3-dimensional models of antibody V regions, including accuracies in the modeling of CDR H3. The need for improvement in this area was also highlighted in a recent study reporting an approach and results that may influence future antibody modeling efforts. One important finding of the antibody modeling assessments was that errors in the structural templates that are used as the basis for homology models can propagate into the final models, producing inaccuracies that may negatively influence the predictive nature of the V region model. To support antibody engineering and therapeutic development efforts, a phage library was designed and constructed based on a limited number of scaffolds built with frequently used human germ-line IGV and IGJ gene segments that encode antigen combining sites suitable for recognition of peptides and proteins. This Fab library is composed of 3 HC germlines, IGHV1-69 (H1-69), IGHV3-23 (H3-23) and IGHV5-51(H5-51), and 4 LC germlines (all κ), IGKV1-39 (L1-39), IGKV3-11 (L3-11), IGKV3-20 (L3-20) and IGKV4-1 (L4-1). Selection of these genes was based on the high frequency of their use and their cognate canonical structures that were found binding to peptides and proteins, as well as their ability to be expressed in bacteria and displayed on filamentous phage. The implementation of the library involves the diversification of the human germline genes to mimic that found in natural human libraries. The crystal structure determinations and structural analyses of all germline Fabs in the library described above along with the structures of a fourth HC germline, IGHV3-53 (H3-53), paired with the 4 LCs of the library have been carried out to support antibody therapeutic development. All 16 HCs of the Fabs have the same CDR H3 that was reported in an earlier Fab structure. This is the first systematic study of the same VH and VL structures in the context of different pairings. The structure analyses include comparisons of the overall structures, canonical structures of the L1, L2, L3, H1 and H2 CDRs, the structures of all CDR H3s, and the VH:VL packing interactions. The structures and their analyses provide a foundation for future antibody engineering and structure determination efforts. The crystal structures of a germline library composed of 16 Fabs generated by combining 4 HCs (H1-69, H3-23, H3-53 and H5-51) and 4 LCs (L1-39, L3-11, L3-20 and L4-1) have been determined. The Fab heavy and light chain sequences for the variants numbered according to Chothia are shown in Fig. S1. The four different HCs all have the same CDR H3 sequence, ARYDGIYGELDF. Crystallization of the 16 Fabs was previously reported. Three sets of the crystals were isomorphous with nearly identical unit cells (Table 1). These include (1) H3-23:L3-11 and H3-23:L4-1 in P212121, (2) H3-53:L1-39, H3-53:L3-11 and H3-53:L3-20 in P6522, and (3) H5-51:L1-39, H5-51:L3-11 and H5-51:L3-20 in P212121. Crystallization conditions for the 3 groups are also similar, but not identical (Table 1). Variations occur in the pH (buffer) and the additives, and, in group 3, PEG 3350 is the precipitant for one variants while ammonium sulfate is the precipitant for the other two. The similarity in the crystal forms is attributed in part to cross-seeding using the microseed matrix screening for groups 2 and 3.Table 1.Crystal data, X-ray data, and refinement statistics.FabH1-69:L1-39H1-69:L3-11H1-69:L3-20H1-69:L4-1PDB identifier5I155I165I175I18Crystal Data Crystallization Solution Buffer, pH0.1 M MES- pH 6.50.1 M MES pH 6.50.1 M MES, pH 6.50.1 M HEPES, pH 7.5 Precipitant5 M Na Formate25% PEG 33502.0 M Amm Sulfate10% PEG 8000 Additive 0.2 M Na Formate5% MPD8% EG Space GroupP3121C2P422P4212 Molecules/AU1221 Unit Cell a(Å)129.2212.0152.5120.0 b(Å)129.255.1152.5120.0 c(Å)91.880.3123.464.2 β(°)90.097.890.090.0 γ(°)120.090.090.090.0 Vm (Å/Da)4.672.443.772.39 Solvent Content (%)74506748X-Ray Data Resolution (Å)30-2.6 (2.7-2.6)30.0-1.9 (1.95-1.9)30.0-3.3 (3.4-3.3)30-1.9 (2.0-1.9) Measured Reflections136,745 (8,650)241,145 (16,580)237,504 (15,007)801,080 (19,309) Unique Reflections27,349 (1,730)71,932 (5,198)22,379 (1,590)35,965 (2,194) Completeness (%)99.3 (98.7)99.0 (97.3)99.5 (96.8)98.5 (82.8) Redundancy5.0 (5.0)3.4 (3.2)10.6 (9.4)22.3 (8.8) Rmerge0.048 (0.522)0.044 (0.245)0.086 (0.536)0.093 (0.231) < I/σ >21.2 (3.9)17.8 (4.7)25.5 (4.5)29.2 (8.1) B-factor (Å)60.533.261.019.6Refinement Resolution (Å)15-2.615-1.915-3.315-1.9 Number of Reflections26,23870,34621,19734,850 Number of All Atoms3,2246,9756,3983,695 Number of Waters24720399 R-factor (%)20.519.220.216.7 R-free (%)24.122.224.721.3RMSD Bond Lengths (Å)0.0060.0050.0050.008 Bond Angles (°)1.21.11.01.1 Mean B-factor (Å)65.334.480.120.0Ramachandran Plot (%) Outliers0.00.00.90.0 Favored92.396.993.196.91Abbreviations: Amm, ammonium;EG, ethylene glycol; PEG, polyethylene glycol.2Values for high-resolution shell are in parentheses.Table 1.(Continued) Crystal data, X-ray data, and refinement statistics.FabH3-23:L1-39H3-23:L3-11H3-23:L3-20H3-23:L4-1 PDB identifier5I195I1A5I1C5I1DCrystal Data Crystallization Solution Buffer, pHNo Buffer0.1 M Na Acetate, pH 4.50.1 M MES, pH 6.50.1 M HEPES, pH 7.5 Precipitant20% PEG 33502.0 M Amm Sulfate16% PEG 33502.0 M Amm Sulfate Additive0.2 M Li Citrate5% PEG 4000.2 M Amm Acetate2% PEG 400 Space GroupP41212P212121P6222P212121 Molecules/AU1212Unit Cell a(Å)96.660.9121.562.7 b(Å)96.6110.6121.5111.0 c(Å)105.4158.9160.4160.0 β(°)90909090 Vm (Å/Da)2.602.823.602.90 Solvent Content (%)53566657X-Ray Data Resolution (Å)30-2.8 (2.9-2.8)30-2.0 (2.1-2.0)30-2.25 (2.3-2.25)30-2.0 (2.1-2.0) Measured Reflections177,681 (12,072)351,312 (8,634)887,349 (59,919)873,523 (49,118) Unique Reflections12,678 (899)58,989 (2,870)32,572 (2,300)75,540 (5,343) Completeness (%)99.5 (97.4)80.9 (54.2)96.9 (94.8)99.7 (96.9) Redundancy14.0 (13.4)6.0 (3.0)27.2 (26.1)11.6 (9.2) Rmerge0.091 (0.594)0.066 (0.204)0.086 (0.478)0.094 (0.488) < I/σ >31.2 (5.1)20.4 (4.6)37.0 (10.4)21.6 (5.0) B-factor (Å)42.827.133.729.4Refinement Resolution (Å)15-2.815-2.015-2.2515-2.0 Number of Reflections11,97257,59931,41174,238 Number of All Atoms3,2346,9483,4727,210 Number of Waters0416222635 R-factor (%)23.920.522.021.6 R-free (%)31.525.526.625.1RMSD Bond Lengths (Å)0.0090.0100.0050.008 Bond Angles (°)1.31.31.01.1 Mean B-factor (Å)48.436.747.746.4Ramachandran Plot (%) Outliers0.00.00.00.0 Favored92.396.897.597.61Abbreviations: Amm, ammonium; PEG, polyethylene glycol.2Values for high-resolution shell are in parentheses.Table 1.(Continued) Crystal data, X-ray data, and refinement statistics.FabH3-53:L1-39H3-53:L3-11H3-53:L3-20H3-53:L4-1PDB indentifier5I1E5I1G5I1H5I1ICrystal Data Crystallization Solution Buffer, pHNo buffer0.1 M Na Acetate pH 4.50.1 M Na Acetate pH 4.50.1M MES, pH 6.5 Precipitant16% PEG 335025% PEG 335019% PEG 400017% PEG 3350 Additive0.2 M Amm Sulfate 5% Dioxane0.2 M Li2SO40.2 M Amm Sulfate0.2 M Na Formate, 5% MPD Space GroupP6522P6522P6522P31 Molecules/AU1111Unit Cell a(Å)89.488.189.468.1 b(Å)89.488.189.468.1 c(Å)212.4219.6211.795.6 β(°)90909090 γ(°)120120120120 Vm (Å/Da)2.572.642.572.64 Solvent Content (%)52535253X-Ray Data Resolution (Å)30-2.7 (2.8-2.7)30-2.3 (2.4-2.3)30-2.2 (2.3-2.0)30-2.5 (2.6-2.5) Measured Reflections297,367 (19,369)333,739 (8,008)381,125 (1,591)137,992 (9,883) Unique Reflections14,402 (1,003)21,683 (1,135)24,323 (964)16,727 (1,227) Completeness (%)99.6 (96.8)93.8 (68.4)95.3 (52.0)98.6 (98.1) Redundancy20.6 (19.3)15.4 (7.1)15.7 (1.7)8.2 (8.1) Rmerge0.095 (0.451)0.057 (0.324)0.062 (0.406)0.047 (0.445) < I/σ >38.3 (8.1)36.7 (5.5)36.2 (1.6)31.6 (5.6) B-factor (Å)33.237.333.754.8Refinement Resolution (Å)15-2.715-2.315-2.215-2.5 Number of Reflections13,58320,25524,96215,811 Number of All Atoms3,3353,2713,2983,239 Number of Waters88707121 R-factor (%)19.129.822.825.0 R-free (%)26.438.326.633.7RMSD Bond Lengths (Å)0.0080.0050.0050.006 Bond Angles (°)1.21.01.01.1 Mean B-factor (Å)49.146.351.788.9Ramachandran Plot (%) Outliers0.20.20.21.2 Favored96.797.196.590.91Abbreviations: Amm, ammonium; PEG, polyethylene glycol.2Values for high-resolution shell are in parentheses.Table 1.(Continued) Crystal data, X-ray data, and refinement statistics.FabH5-51:L1-39H5-51:L3-11H5-51:L3-20H5-51:L4-1 PDB identifier4KMT5I1J5I1K5I1LCrystal Data Crystallization Solution Buffer, pH0.1 M CHES, pH 9.50.1 M Tris, pH 8.50.1 M CHES, pH 9.50.1 M Tris, pH 8.5 Precipitant1.8 M Amm Sulfate25% PEG 33501.0 M Amm Sulfate24% PEG 3350 Additive5% dioxane0.2 M MgCl2 0.2 M Amm Sulfate Space GroupP212121P212121P212121P21 Molecules/AU1112Unit Cell a(Å)63.764.163.8106.0 b(Å)73.873.874.138.0 c(Å)103.1103.0103.0112.3 β(°)909090100.4 Vm (Å/Da)2.532.562.542.28 Solvent Content (%)51525146X-Ray Data Resolution (Å)30-2.1 (2.2-2.1)30-2.5 (2.6-2.5)30-1.65 (1.7-1.65)30-1.95 (2.0-1.95) Measured Reflections131,839 (6,655)120,521 (7,988)246,750 (4,142)320,324 (12,119) Unique Reflections27,026 (1,885)17,286 (1,236)53,058 (2,141)61,554 (3,243) Completeness (%)93.6 (89.8)99.7 (97.3)89.8 (49.8)94.4 (67.1) Redundancy4.9 (3.5)7.0 (6.5)4.7 (1.9)5.2 (3.7) Rmerge0.079 (0.278)0.080 (0.281)0.034 (0.131)0.060 (0.395) < I/σ >16.8 (5.7)21.1(6.9)27.5 (5.8)19.7 (3.1) B-factor (Å)26.027.021.631.4Refinement Resolution (Å)15-2.115-2.515-1.6515-1.95 Number of Reflections25,85716,32851,88260,181 Number of All Atoms3,6763,4543,8147,175 Number of Waters302196527445 R-factor (%)17.117.717.219.4 R-free (%)22.025.819.725.8RMSD Bond Lengths (Å)0.0060.0090.0050.009 Bond Angles (°)1.01.31.31.3 Mean B-factor (Å)25.238.220.019.5Ramachandran Plot (%) Outliers0.00.00.00.0 Favored98.497.998.198.01Abbreviations: Amm, ammonium; PEG, polyethylene glycol.2Values for high-resolution shell are in parentheses. Crystal data, X-ray data, and refinement statistics. Abbreviations: Amm, ammonium;EG, ethylene glycol; PEG, polyethylene glycol. Values for high-resolution shell are in parentheses. (Continued) Crystal data, X-ray data, and refinement statistics. Abbreviations: Amm, ammonium; PEG, polyethylene glycol. Values for high-resolution shell are in parentheses. (Continued) Crystal data, X-ray data, and refinement statistics. Abbreviations: Amm, ammonium; PEG, polyethylene glycol. Values for high-resolution shell are in parentheses. (Continued) Crystal data, X-ray data, and refinement statistics. Abbreviations: Amm, ammonium; PEG, polyethylene glycol. Values for high-resolution shell are in parentheses. The crystal structures of the 16 Fabs have been determined at resolutions ranging from 3.3 Å to 1.65 Å (Table 1). The number of Fab molecules in the crystallographic asymmetric unit varies from 1 (for 12 Fabs) to 2 (for 4 Fabs). Overall the structures are fairly complete, and, as can be expected, the models for the higher resolution structures are more complete than those for the lower resolution structures (Table S1). Invariably, the HCs have more disorder than the LCs. For the LC, the disorder is observed at 2 of the C-terminal residues with few exceptions. Apart from the C-terminus, only a few surface residues in LC are disordered. The HCs feature the largest number of disordered residues, with the lower resolution structures having the most. The C-terminal residues including the 6xHis tags are disordered in all 16 structures. In addition to these, 2 primary disordered stretches of residues are observed in a number of structures (Table S1). One involves the loop connecting the first 2 β-strands of the constant domain (in all Fabs except H3-23:L1-39, H3-23:L3-11 and H3-53:L1-39). The other is located in CDR H3 (in H5-51:L3-11, H5-51:L3-20 and in one of 2 copies of H3-23:L4-1). CDR H1 and CDR H2 also show some degree of disorder, but to a lesser extent. Several CDR definitions have evolved over decades of antibody research. Depending on the focus of the study, the CDR boundaries differ slightly between various definitions. In this work, we use the CDR definition of North et al., which is similar to that of Martin with the following exceptions: 1) CDRs H1 and H3 begin immediately after the Cys; and 2) CDR L2 includes an additional residue at the N-terminal side, typically Tyr. The four HCs feature CDR H1 of the same length, and their sequences are highly similar (Table 2). The CDR H1 backbone conformations for all variants for each of the HCs are shown in Fig. 1. Three of the HCs, H3-23, H3-53 and H5-51, have the same canonical structure, H1-13-1, and the backbone conformations are tightly clustered for each set of Fab structures as reflected in the rmsd values (Fig. 1B-D). Some deviation is observed for H3-53, mostly due to H3-53:L4-1, which exhibits a significant degree of disorder in CDR H1. The electron density for the backbone is weak and discontinuous, and completely missing for several side chains. Figure 1.The superposition of CDR H1 backbones for all HC:LC pairs with heavy chains: (A) H1-69, (B) H3-23, (C) H3-53 and (D) H5-51. Table 2.Canonical structures.PairsPDBCDR H1CDR H2CDR H3H1-69 KASGGTFSSYAISGIIPIFGTANARYDGIYGELDFH1-69:L1-395I15H1-13-4H2-10-1H3-12-1H1-69:L3-115I16H1-13-1/H1-13-1H2-10-1/H2-10-1H3-12-1/H3-12-1H1-69:L3-205I17H1-13-3/H1-13-6H2-10-1/NAH3-12-1/H3-12-1H1-69:L4-15I18H1-13-10H2-10-1H3-12-1H3-23 AASGFTFSSYAMSAISGSGGSTYAKYDGIYDGIYGELDFH3-23:L1-395I19H1-13-1H2-10-2H3-12-1H3-23:L3-115I1AH1-13-1/H1-13-1H2-10-2/H2-10-2H3-12-1/H3-12-1H3-23:L3-205I1CH1-13-1H2-10-2H3-12-1H3-23:L4-15I1DH1-13-1/H1-13-1H2-10-2/H2-10-2H3-12-1/NAH3-53 AASGFTVSSNYMSVIYSGGSTYARYDGIYGELDFH3-53:L1-395I1EH1-13-1H2-9-3H3-12-1H3-53:L3-115I1GH1-13-1H2-9-3H3-12-1H3-53:L3-205I1HH1-13-1H2-9-3H3-12-1H3-53:L4-15I1IH1-13-1H2-9-3NAH5-51 KGSGYSFTSYWIGIIYPGDSDTRARYDGIYGELDFH5-51:L1-394KMTH1-13-1H2-10-1H3-12-1H5-51:L3-115I1JH1-13-1H2-10-1NAH5-51:L3-205I1KH1-13-1H2-10-1NAH5-51:L4-15I1LH1-13-1/H1-13-1H2-10-1/H2-10-1H3-12-1/H3-12-1 CDR L1CDR L2CDR L3L1-39 RASQSISSYLNYAASSLQSQQSYSTPLTH1-69:L1-395I15L1-11-1L2-8-1L3-9-cis7-1H3-23:L1-395I19L1-11-1L2-8-1L3-9-cis7-1H3-53:L1-395I1EL1-11-1L2-8-1L3-9-cis7-1H5-51:L1-394KMTL1-11-1L2-8-1L3-9-cis7-1L3-11 RASQSVSSYLAYDASNRATQQRSNWPLTH1-69:L3-115I16L1-11-1/L1-11-1L2-8-1/L2-8-1L3-9-cis7-1/L3-9-cis7-1H3-23:L3-115I1AL1-11-1/L1-11-1L2-8-1/L2-8-1L3-9-cis7-1/L3-9-cis7-1H3-53:L3-115I1GL1-11-1L2-8-1L3-9-cis7-1H5-51:L3-115I1JL1-11-1L2-8-1L3-9-cis7-1L3-20 RASQSVSSSYLAYGASSRATQQYGSSPLTH1-69:L3-205I17L1-12-2/L1-12-1L2-8-1/L2-8-1L3-9-cis7-1/L3-9-cis7-1H3-23:L3-205I1CL1-12-2L2-8-1L3-9-cis7-1H3-53:L3-205I1HL1-12-1L2-8-1L3-9-cis7-1H5-51:L3-205I1KL1-12-1L2-8-1L3-9-cis7-1L4-1 KSSQSVLYSSNNKNYLAYWASTRESQQYYSTPLTH1-69:L4-15I18L1-17-1L2-8-1L3-9-cis7-1H3-23:L4-15I1DL1-17-1/L1-17-1L2-8-1/L2-8-1L3-9-cis7-1/L3-9-cis7-1H3-53:L4-15I1IL1-17-1L2-8-1L3-9-cis7-1H5-51:L4-15I1LL1-17-1/L1-17-1L2-8-1/L2-8-1L3-9-cis7-1/L3-9-cis7-11CDRs are defined using the Dunbrack convention . Assignments for 2 copies of the Fab in the asymmetric unit are given for 5 structures. No assignment (NA) for CDRs with missing residues. The superposition of CDR H1 backbones for all HC:LC pairs with heavy chains: (A) H1-69, (B) H3-23, (C) H3-53 and (D) H5-51. Canonical structures. CDRs are defined using the Dunbrack convention . Assignments for 2 copies of the Fab in the asymmetric unit are given for 5 structures. No assignment (NA) for CDRs with missing residues. The CDR H1 structures with H1-69 shown in Fig. 1A are quite variable, both for the structures with different LCs and for the copies of the same Fab in the asymmetric unit, H1-69:L3-11 and H1-69:L3-20. In total, 6 independent Fab structures produce 5 different canonical structures, namely H1-13-1, H1-13-3, H1-13-4, H1-13-6 and H1-13-10. A major difference of H1-69 from the other germlines in the experimental data set is the presence of Gly instead of Phe or Tyr at position 27 (residue 5 of 13 in CDR H1). Glycine introduces the possibility of a higher degree of conformational flexibility that undoubtedly translates to the differences observed, and contributes to the elevated thermal parameters for the atoms in the amino acid residues in this region. The canonical structures of CDR H2 have fairly consistent conformations (Table 2, Fig. 2). Each of the 4 HCs adopts only one canonical structure regardless of the pairing LC. Germlines H1-69 and H5-51 have the same canonical structure assignment H2-10-1, H3-23 has H2-10-2, and H3-53 has H2-9-3. The conformations for all of these CDR H2s are tightly clustered (Fig. 2). In one case, in the second Fab of H1-69:L3-20, CDR H2 is partially disordered (Δ55-60). Figure 2.The superposition of CDR H2 backbones for all HC:LC pairs with heavy chains: (A) H1-69, (B) H3-23, (C) H3-53 and (D) H5-51. The superposition of CDR H2 backbones for all HC:LC pairs with heavy chains: (A) H1-69, (B) H3-23, (C) H3-53 and (D) H5-51. Although three of the germlines have CDR H2 of the same length, 10 residues, they adopt 2 distinctively different conformations depending mostly on the residue at position 71 from the so-called CDR H4. Arg71 in H3-23 fills the space between CDRs H2 and H4, and defines the conformation of the tip of CDR H2 so that residue 54 points away from the antigen binding site. Germlines H1-69 and H5-51 are unique in the human repertoire in having an Ala at position 71 that leaves enough space for H-Pro52a to pack deeper against CDR H4 so that the following residues 53 and 54 point toward the putative antigen. Conformations of CDR H2 in H1-69 and H5-51, both of which have canonical structure H2-10-1, show little deviation within each set of 4 structures. However, there is a significant shift of the CDR as a rigid body when the 2 sets are superimposed. Most likely this is the result of interaction of CDR H2 with CDR H1, namely with the residue at position 33 (residue 11 of 13 in CDR H1). Germline H1-69 has Ala at position 33 whereas in H5-51 position 33 is occupied by a bulky Trp, which stacks against H-Tyr52 and drives CDR H2 away from the center. The four LC CDRs L1 feature 3 different lengths (11, 12 and 17 residues) having a total of 4 different canonical structure assignments. Of these LCs, L1-39 and L3-11 have the same canonical structure, L1-11-1, and superimpose very well (Fig. 3A, B). For the remaining 2, L3-20 has 2 different assignments, L1-12-1 and L1-12-2, while L4-1 has a single assignment, L1-17-1. Figure 3.The superposition of CDR L1 backbones for all HC:LC pairs with light chains: (A) L1-39, (B) L3-11, (C) L3-20 and (D) L4-1. The superposition of CDR L1 backbones for all HC:LC pairs with light chains: (A) L1-39, (B) L3-11, (C) L3-20 and (D) L4-1. L4-1 has the longest CDR L1, composed of 17 amino acid residues (Fig. 3D). Despite this, the conformations are tightly clustered (rmsd is 0.20 Å). The backbone conformations of the stem regions superimpose well. Some changes in conformation occur between residues 30a and 30f (residues 8 and 13 of 17 in CDR L1). This is the tip of the loop region, which appears to have similar conformations that fan out the structures because of the slight differences in torsion angles in the backbone near Tyr30a and Lys30f. L3-20 is the most variable in CDR L1 among the 4 germlines as indicated by an rmsd of 0.54 Å (Fig. 3C). Two structures, H3-53:L3-20 and H5-51:L3-20 are assigned to canonical structure L1-12-1 with virtually identical backbone conformations. The third structure, H3-23:L3-20, has CDR L1 as L1-12-2, which deviates from L1-12-1 at residues 29-32, i.e., at the site of insertion with respect to the 11-residue CDR. The fourth member of the set, H1-69:L3-20, was crystallized with 2 Fabs in the asymmetric unit. The conformation of CDR L1 in these 2 Fabs is slightly different, and both conformations fall somewhere between L1-12-1 and L1-12-2. This reflects the lack of accuracy in the structure due to low resolution of the X-ray data (3.3 Å). All four LCs have CDR L2 of the same length and canonical structure, L2-8-1 (Table 2). The CDR L2 conformations for each of the LCs paired with the 4 HCs are clustered more tightly than any of the other CDRs (rmsd values are in the range 0.09-0.16 Å), and all 4 sets have virtually the same conformation despite the sequence diversity of the loop. No significant conformation outliers are observed (Fig. 4). Figure 4.The superposition of CDR L2 backbones for all HC:LC pairs with light chains: (A) L1-39, (B) L3-11, (C) L3-20 and (D) L4-1. The superposition of CDR L2 backbones for all HC:LC pairs with light chains: (A) L1-39, (B) L3-11, (C) L3-20 and (D) L4-1. As with CDR L2, all 4 LCs have CDR L3 of the same length and canonical structure, L3-9-cis7-1 (Table 2). The conformations of CDR L3 for L1-39, L3-11, and particularly for L320, are not as tightly clustered as those of L4-1 (Fig. 5). The slight conformational variability occurs in the region of amino acid residues 90-92, which is in contact with CDR H3. Figure 5.The superposition of CDR L3 backbones for all HC:LC pairs with light chains: (A) L1-39, (B) L3-11, (C) L3-20 and (D) L4-1. The superposition of CDR L3 backbones for all HC:LC pairs with light chains: (A) L1-39, (B) L3-11, (C) L3-20 and (D) L4-1. As mentioned earlier, all 16 Fabs have the same CDR H3, for which the amino acid sequence is derived from the anti-CCL2 antibody CNTO 888. The loop and the 2 β-strands of the CDR H3 in this ‘parent’ structure are stabilized by H-bonds between the carbonyl oxygen and peptide nitrogen atoms in the 2 strands. An interesting feature of these CDR H3 structures is the presence of a water molecule that interacts with the peptide nitrogens and carbonyl oxygens near the bridging loop connecting the 2 β-strands. This water is present in both the bound (4DN4) and unbound (4DN3) forms of CNTO 888. The stem region of CDR H3 in the parental Fab is in a ‘kinked’ conformation, in which the indole nitrogen of Trp103 forms a hydrogen bond with the carbonyl oxygen of Leu100b. The carboxyl group of Asp101 forms a salt bridge with Arg94. These interactions are illustrated in Fig. S2. Despite having the same amino acid sequence in all variants, CDR H3 has the highest degree of structural diversity and disorder of all of the CDRs in the experimental set. Three of the 21 Fab structures (including multiple copies in the asymmetric unit), H5-51:L3-11, H551:L3-20 and H3-23:L4-1 (one of the 2 Fabs), have missing (disordered) residues at the apex of the CDR loop. Another four of the Fabs, H3-23:L1-39, H3-53:L1-39, H3-53:L3-11 and H3-53:L4-1 have missing side-chain atoms. The variations in CDR H3 conformation are illustrated in Fig. 6 for the 18 Fab structures that have ordered backbone atoms. Figure 6.Ribbon representations of (A) the superposition of all CDR H3s of the structures with complete backbone traces. (B) The CDR H3s rotated 90° about the y axis of the page. The structure of each CDR H3 is represented with a different color. Ribbon representations of (A) the superposition of all CDR H3s of the structures with complete backbone traces. (B) The CDR H3s rotated 90° about the y axis of the page. The structure of each CDR H3 is represented with a different color. In 10 of the 18 Fab structures, H1-69:L1-39, H1-69:L3-11 (2 Fabs), H1-69:L4-1, H3-23:L3-11 (2 Fabs), H3-23:L3-20, H3-53:L3-11, H3-53:L3-20 and H5-51:L1-39, the CDRs have similar conformations to that found in 4DN3. The bases of these structures have the ‘kinked’ conformation with the H-bond between Trp103 and Leu100b. A representative CDR H3 structure for H1-69:L1-39 illustrating this is shown in Fig. 7A. The largest backbone conformational deviation for the set is at Tyr99, where the C=O is rotated by 90° relative to that observed in 4DN3. Also, it is worth noting that only one of these structures, H1-69:L4-1, has the conserved water molecule in CDR H3 observed in the 4DN3 and 4DN4 structures. In fact, it is the only Fab in the set that has a water molecule present at this site. The CDR H3 for this structure is shown in Fig. S3. Figure 7.A comparison of representatives of the “kinked” and “extended” structures. (A) The “kinked” CDR H3 of H1-69:L3-11 with purple carbon atoms and yellow dashed lines connecting the H-bond pairs for Leu100b O and Trp103 NE1, Arg94 NE and Asp101 OD1, and Arg94 NH2 and Asp101 OD2. (B) The “extended” CDR H3 of H1-69:L3-20 with green carbon atoms and yellow dashed lines connecting the H-bond pairs for Asp101 OD1 and OD2 and Trp103 NE1. A comparison of representatives of the “kinked” and “extended” structures. (A) The “kinked” CDR H3 of H1-69:L3-11 with purple carbon atoms and yellow dashed lines connecting the H-bond pairs for Leu100b O and Trp103 NE1, Arg94 NE and Asp101 OD1, and Arg94 NH2 and Asp101 OD2. (B) The “extended” CDR H3 of H1-69:L3-20 with green carbon atoms and yellow dashed lines connecting the H-bond pairs for Asp101 OD1 and OD2 and Trp103 NE1. The remaining 8 Fabs can be grouped into 5 different conformational classes. Three of the Fabs, H3-23:L1-39, H3-23:L4-1 and H3-53:L1-39, have distinctive conformations. The stem regions in these 3 cases are in the ‘kinked’ conformation consistent with that observed for 4DN3. The five remaining Fabs, H5-51:L4-1 (2 copies), H1-69:L3-20 (2 copies) and H3-53:L4-1, have 3 different CDR H3 conformations (Fig. S4). The stem regions of CDR H3 for the H5-51:L4-1 Fabs are in the ‘kinked’ conformation while, surprisingly, those of the H1-69:L3-20 pair and H3-53:L4-1 are in the ‘extended’ conformation (Fig. 7B). The VH and VL domains have a β-sandwich structure (also often referred as a Greek key motif) and each is composed of a 4-stranded and a 5-stranded antiparallel β-sheets. The two domains pack together such that the 5-stranded β-sheets, which have hydrophobic surfaces, interact with each other bringing the CDRs from both the VH and VL domains into close proximity. The domain packing of the variants was assessed by computing the domain interface interactions, the VH:VL tilt angles, the buried surface area and surface complementarity. The results of these analyses are shown in Tables 3, 4 and S2. The VH:VL interface is pseudosymmetric, and involves 2 stretches of the polypeptide chain from each domain, namely CDR3 and the framework region between CDRs 1 and 2. These stretches form antiparallel β-hairpins within the internal 5-stranded β-sheet. There are a few principal inter-domain interactions that are conserved not only in the experimental set of 16 Fabs, but in all human antibodies. They include: 1) a bidentate hydrogen bond between L-Gln38 and H-Gln39; 2) H-Leu45 in a hydrophobic pocket between L-Phe98, L-Tyr87 and L-Pro44; 3) L-Pro44 stacked against H-Trp103; and 4) L-Ala43 opposite the face of H-Tyr91 (Fig. 8). With the exception of L-Ala43, all other residues are conserved in human germlines. Position 43 may be alternatively occupied by Ser, Val or Pro (as in L4-1), but the hydrophobic interaction with H-Tyr91 is preserved. These core interactions provide enough stability to the VH:VL dimer so that additional VH-VL contacts can tolerate amino acid sequence variations in CDRs H3 and L3 that form part of the VH:VL interface. Figure 8.The conserved VH:VL interactions as viewed along the VH/VL axis. The VH residues are in blue, the VL residues are in orange. The conserved VH:VL interactions as viewed along the VH/VL axis. The VH residues are in blue, the VL residues are in orange. In total, about 20 residues are involved in the VH:VL interactions on each side (Fig. S5). Half of them are in the framework regions and those residues (except residue 61 in HC, which is actually in CDR2 in Kabat's definition) are conserved in the set of 16 Fabs. The side chain conformations of these conserved residues are also highly similar. One notable exception is H-Trp47, which exhibits 2 conformations of the indole ring. In most of the structures, it has the χ2 angle of ∼80°, while the ring is flipped over (χ2 = −100°) in H5-51:L3:11 and H5-51:L3-20. Interestingly, these are the only 2 structures with residues missing in CDR H3 because of disorder, although both structures are determined at high resolution and the rest of the structure is well defined. Apparently, residues flanking CDR H3 in the 2 VH:VL pairings are inconsistent with any stable conformation of CDR H3, which translates into a less restricted conformational space for some of them, including H-Trp47. The relative orientation of VH and VL has been measured in a number of different ways. Presented here are the results of 2 different approaches for determining the orientation of one domain relative to the other. The first approach uses ABangles, the results of which are shown in Table S2. The four LCs all are classified as Type A because they have a proline at position 44, and the results for each orientation parameter are within the range of values of this type reported by Dunbar and co-workers. In fact, the parameter values for the set of 16 Fabs are in the middle of the distribution observed for 351 non-redundant antibody structures determined at 3.0 Å resolution or better. The only exception is HC1, which is shifted toward smaller angles with the mean value of 70.8° as compared to the distribution centered at 72° for the entire PDB. This probably reflects the invariance of CDR H3 in the current set as opposed to the CDR H3 diversity in the PDB. The second approach used for comparing tilt angles involved computing the difference in the tilt angles between all pairs of structures. For structures with 2 copies of the Fab in the asymmetric unit, only one structure was used. The differences between independent Fabs in the same structure are 4.9° for H1-69:L3-20, 1.6° for H1-69:L3-11, 1.4° for H3-23:L4-1, 3.3° for H3-23:L3-11, and 2.5° for H5-51:L4-1. With the exception of H1-69:L3-20, the angles are within the range of 2-3° as are observed in the identical structures in the PDB. In H1-69:L3-20, one of the Fabs is substantially disordered so that part of CDR H2 (the outer β-strand, residues 55-60) is completely missing. This kind of disorder may compromise the integrity of the VH domain and its interaction with the VL. Indeed, this Fab has the largest twist angle HC2 within the experimental set that exceeds the mean value by 2.5 standard deviations (Table S2). The differences in the tilt angle are shown for all pairs of V regions in Table 3. They range from 0.6° to 11.0°. The smallest differences in the tilt angle are between the Fabs in isomorphous crystal forms. The largest deviations in the tilt angle, up to 11.0°, are found for 2 structures, H1-69:L3-20 and H3-23:L3-20, that stand out from the other Fabs. One of the 2 structures, H1-69:L3-20, has its CDR H3 in the ‘extended’ conformation; the other structure has it in the ‘kinked’ conformation. Two examples illustrating large (10.5°) and small (1.6°) differences in the tilt angles are shown in Fig. 9. Figure 9.An illustration of the difference in tilt angle for 2 pairs of variants by the superposition of the VH domains of (A) H1-69:L3-20 on that of H5-51:L1-39 (the VL domain is off by a rigid-body roatation of 10.5°) and (B) H1-69:L4-1 on that of H5-51:L1-39 (the VL domain is off by a rigid-body roatation of 1.6°). Table 3.Differences in VH:VL tilt angles. H1-69:L1-39H1-69:L3-11H1-69:L3-20H1-69:L4-1H3-23:L1-39H3-23:L3-11H3-23:L3-20H3-23:L4-1H3-53:L1-39H3-53:L3-11H3-53:L3-20H3-53:L4-1H5-51:L1-39H5-51:L3-11H5-51:L3-20H5-51:L4-1H1-69:L1-3902.18.91.14.23.09.51.53.33.63.11.61.82.92.45.2H1-69:L3-11 07.32.92.52.08.41.32.62.93.21.83.94.64.45.0H1-69:L3-20 09.25.08.77.47.68.98.69.47.910.510.111.09.7H1-69:L4-1 04.63.910.11.84.44.74.12.31.62.52.36.2H3-23:L1-39 04.08.02.84.84.85.43.66.06.26.66.6H3-23:L3-11 09.33.02.12.93.33.34.65.85.05.2H3-23:L3-20 08.97.97.07.67.910.59.710.76.2H3-23:L4-1 03.63.83.71.53.23.73.85.6H3-53:L1-39 01.01.62.94.65.34.83.1H3-53:L3-11 01.32.94.85.25.02.3H3-53:L3-20 02.53.84.23.92.2H3-53:L4-1 02.93.03.34.2H5-51:L1-39 01.90.65.8H5-51:L3-11 01.95.7H5-51:L3-20 05.8H5-51:L4-1 0 An illustration of the difference in tilt angle for 2 pairs of variants by the superposition of the VH domains of (A) H1-69:L3-20 on that of H5-51:L1-39 (the VL domain is off by a rigid-body roatation of 10.5°) and (B) H1-69:L4-1 on that of H5-51:L1-39 (the VL domain is off by a rigid-body roatation of 1.6°). Differences in VH:VL tilt angles. The results of the PISA contact surface calculation and surface complementarity calculation are shown in Table 4. The interface areas are calculated as the average of the VH and VL contact surfaces. Six of the 16 structures have CDR H3 side chains or complete residues missing, and therefore their interfaces are much smaller than in the other 10 structures with complete CDRs (the results are provided for all Fabs for completeness). Among the complete structures, the interface areas range from 684 to 836 Å. Interestingly, the 2 structures that have the largest tilt angle differences with the other variants, H3-23:L3-20 and H1-69:L3-20, have the smallest VH:VL interfaces, 684 and 725 Å, respectively. H3-23:L3-20 is also unique in that it has the lowest value (0.676) of surface complementarity. Table 4.VH:VL surface areas and surface complementarity.Chain PairsPDBContact surfaceVH (Å)Contact surfaceVL (Å)Interface(Å)Surface complementarityH1-69:L1-395I157277717490.743H1-69:L3-115I168028708360.762H1-69:L3-205I177137367250.723H1-69:L4-15I187297367330.734H3-23:L1-395I197958178060.722H3-23:L3-115I1A8228348280.725H3-23:L3-205I1C6706986840.676H3-23:L4-15I1D7437707570.708H3-53:L1-395I1E6987197090.712H3-53:L3-115I1G7477587530.690H3-53:L3-205I1H7437357390.687H3-53:L4-15I1I6896936910.711H5-51:L1-394KMT7618087850.728H5-51:L3-115I1J6487146810.717H5-51:L3-205I1K6226436330.740H5-51:L4-15I1L7907927910.7041Some side chain atoms in CDR H3 are missing.2Residues in CDR H3 are missing: YGE in H5-51:L3-11, GIY in H5-51:L3-20. VH:VL surface areas and surface complementarity. Some side chain atoms in CDR H3 are missing. Residues in CDR H3 are missing: YGE in H5-51:L3-11, GIY in H5-51:L3-20. Melting temperatures (Tm) were measured for all Fabs using differential scanning calorimetry (Table 5). It appears that for each given LC, the Fabs with germlines H1-69 and H3-23 are substantially more stable than those with germlines H3-53 and H5-51. In addition, L1-39 provides a much higher degree of stabilization than the other 3 LC germlines when combined with any of the HCs. As a result, the Tm for pairs H1-69:L1-39 and H3-23:L1-39 is 12-13° higher than for pairs H3-53:L3-20, H3-53:L4-1, H5-51:L3-20 and H5-51:L4-1. Table 5.Melting temperatures for the 16 Fabs. L3-20L4-1L3-11L1-39HC averageH1-6973.674.875.680.376.1H3-2374.875.24.881.576.6H3-5368.468.071.573.970.5H5-5168.468.471.977.071.4LC average71.371.673.578.2 1Colors: blue (Tm < 70°C), green (70°C < Tm < 73°C), yellow (73°C < Tm < 78°C), orange (Tm > 78°C). Melting temperatures for the 16 Fabs. Colors: blue (Tm < 70°C), green (70°C < Tm < 73°C), yellow (73°C < Tm < 78°C), orange (Tm > 78°C). These findings correlate well with the degree of conformational disorder observed in the crystal structures. Parts of CDR H3 main chain are completely disordered, and were not modeled in Fabs H5-51:L3-20 and H5-51:L3-11 that have the lowest Tms in the set. No electron density is observed for a number of side chains in CDRs H3 and L3 in all Fabs with germline H3-53, which indicates loose packing of the variable domains. All those molecules are relatively unstable, as is reflected in their low Tms. This is the first report of a systematic structural investigation of a phage germline library. The 16 Fab structures offer a unique look at all pairings of 4 different HCs (H1-69, H3-23, H3-53, and H5-51) and 4 different LCs (L1-39, L3-11, L3-20 and L4-1), all with the same CDR H3. The structural data set taken as a whole provides insight into how the backbone conformations of the CDRs of a specific heavy or light chain vary when it is paired with 4 different light or heavy chains, respectively. A large variability in the CDR conformations for the sets of HCs and LCs is observed. In some cases the CDR conformations for all members of a set are virtually identical, for others subtle changes occur in a few members of a set, and in some cases larger deviations are observed within a set. The five variants that crystallized with 2 copies of the Fab in the asymmetric unit serve somewhat as controls for the influence of crystal packing on the conformations of the CDRs. In four of the 5 structures the CDR conformations are consistent. In only one case, that of H1-69:L3-20 (the lowest resolution structure), do we see differences in the conformations of the 2 copies of CDRs H1 and L1. This variability is likely a result of 2 factors, crystal packing interactions and internal instability of the variable domain. For the CDRs with canonical structures, the largest changes in conformation occur for CDR H1 of H1-69 and H3-53. The other 2 HCs, H3-23 and H5-51, have canonical structures that are remarkably well conserved (Fig. 1). Of the 4 HCs, H1-69 has the greatest number of canonical structure assignments (Table 2). H1-69 is unique in having a pair of glycine residues at positions 26 and 27, which provide more conformational freedom in CDR H1. Besides IGHV1-69, only the germlines of the VH4 family possess double glycines in CDR H1, and it will be interesting to see if they are also conformationally unstable. Having all 16 VH:VL pairs with the same CDR H3 provides some insights into why molecular modeling efforts of CDR H3 have proven so difficult. As mentioned in the Results section, this data set is composed of 21 Fabs, since 5 of the 16 variants have 2 Fab copies in the asymmetric unit. For the 18 Fabs with complete backbone atoms for CDR H3, 10 have conformations similar to that of the parent, while the others have significantly different conformations (Fig. 6). Thus, it is likely that the CDR H3 conformation is dependent upon 2 dominating factors: 1) amino acid sequence; and 2) VH and VL context. More than half of the variants retain the conformation of the parent despite having differences in the VH:VL pairing. This subset includes 2 structures with 2 copies of the Fab in the asymmetric unit, all of which are nearly identical in conformation. This provides an internal control showing a consistency in the conformations. The remaining 8 structures exhibit “non-parental” conformations, indicating that the VH and VL context can also be a dominating factor influencing CDR H3. Importantly, there are 5 distinctive conformations in this subset. This subset also has 2 structures with 2 Fab copies in the asymmetric unit. Each pair has nearly identical conformations providing an internal check on the consistency of the conformations. Interestingly, as described earlier, these 2 pairs differ in the stem regions with the H1-69:L3-20 pair in the ‘extended’ conformation and H5-51:L4-1 pair in the ‘kinked’ conformation. The conformations are different from each other, as well as from the parent. The CDR H3 conformational analysis shows that, for each set of variants of one HC paired with the 4 different LCs, both “parental” and “non-parental” conformations are observed. The same variability is observed for the sets of variants composed of one LC paired with each of the 4 HCs. Thus, no patterns of conformational preference for a particular HC or LC emerge to shed any direct light on what drives the conformational differences. This finding supports the hypothesis of Weitzner et al. that the H3 conformation is controlled both by its sequence and its environment. In looking at a possible correlation between the tilt angle and the conformation of CDR H3, no clear trends are observed. Two variants, H1-69:L3-20 and H3-23:L3-20, have the largest differences in the tilt angles compared to other variants as seen in Table 3. The absolute VH:VL orientation parameters for the 2 Fabs (Table S2) show significant deviation in HL, LC1 and HC2 values (2-3 standard deviations from the mean). One of the variants, H3-23:L3-20, has the CDR H3 conformation similar to the parent, but the other, H1-69:L3-20, is different. As noted in the Results section, the 2 variants, H1-69:L3-20 and H3-23:L3-20, are outliers in terms of the tilt angle; at the same time, both have the smallest VH:VL interface. These smaller interfaces may perhaps translate to a significant deviation in how VH is oriented relative to VL than the other variants. These deviations from the other variants can also be seen to some extent in VH:VL orientation parameters in Table S2, as well as in the smaller number of residues involved in the VH:VL interfaces of these 2 variants (Fig. S5). These differences undoubtedly influence the conformation of the CDRs, in particular CDR H1 (Fig. 1A) and CDR L1 (Fig. 3C), especially with the tandem glycines and multiple serines present, respectively. Pairing of different germlines yields antibodies with various degrees of stability. As indicated by the melting temperatures, germlines H1-69 and H3-23 for HC and germline L1-39 for LC produce more stable Fabs compared to the other germlines in the experimental set. Structural determinants of the differential stability are not always easy to decipher. One possible explanation of the clear preference of LC germline L1-39 is that CDR L3 has smaller residues at positions 91 and 94, allowing for more room to accommodate CDR H3. Other germlines have bulky residues, Tyr, Arg and Trp, at these positions, whereas L1-39 has Ser and Thr. Various combinations of germline sequences for VL and VH impose certain constraints on CDR H3, which has to adapt to the environment. A more compact CDR L3 may be beneficial in this situation. At the other end of the stability range is LC germline L3-20, which yields antibodies with the lowest Tms. While pairings with H3-53 and H5-51 may be safely called a mismatch, those with H1-69 and H3-23 have Tms about 5-6° higher. Curiously, the 2 Fabs, H1-69:L3-20 and H3-23:L3-20, deviate markedly in their tilt angles from the rest of the panel. It is possible that by adopting extreme tilt angles the structure modulates CDR H3 and its environment, which apparently cannot be achieved solely by conformational rearrangement of the CDR. Note that most of the VH:VL interface residues are invariant; therefore, significant change of the tilt angle must come with a penalty in free energy. Yet, for the 2 antibodies, the total gain in stability merits the domain repacking. Overall, the stability of the Fab, as measured by Tm, is a result of the mutual adjustment of the HC and LC variable domains and adjustment of CDR H3 to the VH:VL cleft. The final conformation represents an energetic minimum; however, in most cases it is very shallow, so that a single mutation can cause a dramatic rearrangement of the structure. In summary, the analysis of this structural library of germline variants composed of all pairs of 4 HCs and 4LCs, all with the same CDR H3, offers some unique insights into antibody structure and how pairing and sequence may influence, or not, the canonical structures of the L1, L2, L3, H1 and H2 CDRs. Comparison of the CDR H3s reveals a large set of variants with conformations similar to the parent, while a second set has significant conformational variability, indicating that both the sequence and the structural context define the CDR H3 conformation. Quite unexpectedly, 2 of the variants, H1-69:L3-20 and H3-53:L4-1, have the ‘extended’ stem region differing from the other 14 that have a ‘kinked’ stem region. Why this is the case is unclear at present. These data reveal the difficulty of modeling CDR H3 accurately, as shown again in Antibody Modeling Assessment II. Furthermore, antibody CDRs, H3 in particular, may go through conformational changes upon binding their targets, making structural prediction for docking purposes an even more difficult task. Fortunately, for most applications of antibody modeling, such as engineering affinity and biophysical properties, an accurate CDR H3 structure is not always necessary. For those applications where accurate CDR structures are essential, such as docking, the results in this work demonstrate the importance of experimental structures. With the recent advances in expression and crystallization methods, Fab structures can be obtained rapidly. The set of 16 germline Fab structures offers a unique dataset to facilitate software development for antibody modeling. The results essentially support the underlying idea of canonical structures, indicating that most CDRs with germline sequences tend to adopt predefined conformations. From this point of view, a novel approach to design combinatorial antibody libraries would be to cover the range of CDR conformations that may not necessarily coincide with the germline usage in the human repertoire. This would insure more structural diversity, leading to a more diverse panel of antibodies that would bind to a broad spectrum of targets. The production, purification and crystallization of the Fabs reported in this article were described previously. Briefly, the 16 Fabs were produced by combining 4 different HC and 4 different LC germline constructs. The human HC germlines were IGHV1-69 (H1-69), IGHV3-23 (H3-23), IGHV3-53 (H3-53) and IGHV5-51 (H5-51) in the IMGT nomenclature. The human LC germlines were IGKV1-39 (L1-39), IGKV3-11 (L3-11), IGKV3-20 (L3-20) and IGKV4-1 (L4-1) corresponding to O12, L6, A27 and B3 in the V-BASE nomenclature. CDR H3 of the anti-CCL2 antibody CNTO 888 with the amino acid sequence ARYDGIYGELDF was used in all Fab constructs. The J region genes were IGHJ1 for the HC and IGKJ1 for the LC for all Fabs. Human IgG1 and κ constant regions were used in all Fab constructs. A 6xHis tag was added to the C-terminus of the HC to facilitate purification. The Fabs were expressed in HEK 293E cells and purified by affinity and size-exclusion chromatography. For crystallization, the Fabs were dialyzed into 20 mM Tris buffer, pH 7.4, with 50 mM NaCl and concentrated to 12-18 mg/mL. Automated crystallization screening was carried out using the vapor diffusion method at 20°C with an Oryx4 (Douglas Instruments) or a Mosquito (TTP Labtech) crystallization robot in a sitting drop format using Corning 3550 plates. Initial screening was carried out with an in-house 192-well screen optimized for Fab crystallization and the Hampton 96-well Crystal Screen HT (Hampton Research). For the majority of the Fabs, the crystallization protocol employed microseed matrix screening using self-seeding or cross-seeding approaches. A summary of the final crystallization conditions for each of the Fabs is presented in Table 1. For 13 of the Fab crystals, X-ray data collection was carried out at Janssen Research and Development, LLC using a Rigaku MicroMax™-007HF microfocus X-ray generator equipped with a Saturn 944 CCD detector and an X-stream™ 2000 cryocooling system (Rigaku), and for the remaining 3, X-ray data collection was carried out at the Advanced Photon Source (APS) synchrotron at Argonne National Laboratory using the IMCA 17-ID beamline with a Pilatus 6M detector. For X-ray data collection, the Fab crystals were soaked for a few seconds in a cryo-protectant solution containing the corresponding mother liquor supplemented with 17-25% glycerol (Table S1). The crystals for which data were collected in-house were flash cooled in the stream of nitrogen at 100 K. Crystals sent to the APS were flash cooled in liquid nitrogen prior to shipping them to the synchrotron. Diffraction data for all variants were processed with the program XDS. X-ray data statistics are given in Table 1. A summary of the methods used in the structure solution and refinement of the 16 Fabs is presented in Table S1. Twelve of the structures were solved by molecular replacement with Phaser using different combinations of search models for the VH, VL and constant domains. Four of the structures, H3-53:L1-39, H3-53:L3-11, H5-51:L1-39 and H5-51:L3-11, were solved by direct replacement followed by rigid body refinement with REFMAC. All structures were refined using REFMAC. Model adjustments were carried out using the program Coot. The refinement statistics are given in Table 1. Other crystallographic calculations were performed with the CCP4 suite of programs. The structural figures were prepared using the PyMOL Molecular Graphics System, Version 1.0 (Schrödinger, LLC). The canonical structure assignments (Table 2) were made using PyIgClassify, an online canonical structure classification tool (http://dunbrack2.fccc.edu/pyigclassify/) that uses the rules set forth by Dunbrack and coworkers. The conformational variability within the CDRs was assessed by calculating the root-mean-square deviation (rmsd) from the average structure that was generated after superposition of all structures of the set using the main-chain atoms of the CDR in question. The rmsd was calculated for all main-chain atoms (N, CA, C, O) of the CDR. The contact surface areas of the VH and VL domains at the VH:VL inteface were computed with the CCP4 program PISA. The surface complementarity of the VH and VL domains was computed using the CCP4 program SC. The orientation of the VH domain with respect to the VL domain was assessed using 2 different approaches. The first approach calculates the 6 VH:VL orientation parameters that describe the VH:VL relationship according to Dunbar and co-workers using a script downloaded from the website (http://opig.stats.ox.ac.uk/webapps/abangle). The six parameters include 5 angles, HL, H1, H2, L1 and L2, and a distance, dc. These parameters are derived by first defining 2 planes, one for each domain, based on core residues in the domains. The distance between the planes, dc, is determined along a vector between the planes that is used to establish a consistent coordinate system. The torsion angle between the domains, HL, is much like the VH:VL packing angle defined by Abhinandan and Martin. The tilt of one domain relative to the other is defined by the HC1 and LC1 angles, and the twist of one domain relative to the other is defined by the HC2 and LC2 angles. The second approach calculates the difference in the tilt angle between pairs of Fvs, which reflects the relative orientation between the VH and VL domains. The difference with respect to the reference structure is calculated by sequential root-mean-square superposition of the VL and VH domains using β-sheet core Cα positions (Chothia numbering scheme): 3–13, 18–25, 33–38, 43–49, 61–67, 70–76, 85–90, 97–103 for VL; 3–7, 18–24, 34–40, 44–51, 56–59, 67–72, 77–82a, 87–94, 102–110 for VH. The κ angle in the spherical polar angular system (ω, ϕ, κ) of the latter transformation is the difference in the tilt angle. DSC experiments were performed on a VP-capillary DSC system (MicroCal Inc., Northampton, MA) in which temperature differences between the reference and sample cell are continuously measured and calibrated to power units. Samples were heated from 10°C to 95°C at a heating rate of 60°C/hour. The pre-scan time was 15 minutes and the filtering period was 10 seconds. The concentration used in the DSC experiments was about 0.4 mg/mL in phosphate-buffered saline. Analysis of the resulting thermograms was performed using MicroCal Origin 7 software. Melting temperature of proteins was determined by deconvolution of the DSC scans using non-2 state model in the MicroCal Origin 7 software. Scans were deconvoluted using a non-2 state model with either 1-step transition or 2-step transition depending on the number of resolved peaks observed in a scan. Atomic coordinates and structure factors have been deposited in the Protein Data Bank with accession numbers 4KMT, 5I15, 5I16, 5I17, 5I18, 5I19, 5I1A, 5I1C, 5I1D, 5I1E, 5I1G, 5I1H, 5I1I, 5I1J, 5I1K and 5I1L.
PMC4854314	RNA protects a nucleoprotein complex against radiation damage	Systematic analysis of radiation damage within a protein–RNA complex over a large dose range (1.3–25 MGy) reveals significant differential susceptibility of RNA and protein. A new method of difference electron-density quantification is presented.With the wide use of high-flux third-generation synchrotron sources, radiation damage (RD) has once again become a dominant reason for the failure of structure determination using macromolecular crystallography (MX) in experiments conducted both at room temperature and under cryocooled conditions (100 K). Significant progress has been made in recent years in understanding the inevitable manifestations of X-ray-induced RD within protein crystals, and there is now a body of literature on possible strategies to mitigate the effects of RD (e.g. Zeldin, Brockhauser et al., 2013 ▸; Bourenkov & Popov, 2010 ▸). However, there is still no general consensus within the field on how to minimize RD during MX data collection, and debates on the dependence of RD progression on incident X-ray energy (Shimizu et al., 2007 ▸; Liebschner et al., 2015 ▸) and the efficacy of radical scavengers (Allan et al., 2013 ▸) have yet to be resolved. RD manifests in two forms. Global radiation damage is observed within reciprocal space as the overall decay of the summed intensity of reflections detected within the diffraction pattern as dose increases (Garman, 2010 ▸; Murray & Garman, 2002 ▸). Dose is defined as the absorbed energy per unit mass of crystal in grays (Gy; 1 Gy = 1 J kg), and is the metric against which damage progression should be monitored during MX data collection, as opposed to time. At 100 K, an experimental dose limit of 30 MGy has been recommended as an upper limit beyond which the biological information derived from any macromolecular crystal may be compromised (Owen et al., 2006 ▸). Specific radiation damage (SRD) is observed in the real-space electron density, and has been detected at much lower doses than any observable decay in the intensity of reflections. Indeed, the C—Se bond in selenomethionine, the stability of which is key for the success of experimental phasing methods, can be cleaved at a dose as low as 2 MGy for a crystal maintained at 100 K (Holton, 2007 ▸). SRD has been well characterized in a large range of proteins, and is seen to follow a reproducible order: metallo-centre reduction, disulfide-bond cleavage, acidic residue decarboxylation and methionine methylthio cleavage (Ravelli & McSweeney, 2000 ▸; Burmeister, 2000 ▸; Weik et al., 2000 ▸; Yano et al., 2005 ▸). Furthermore, damage susceptibility within each residue type follows a preferential ordering influenced by a combination of local environment factors (solvent accessibility, conformational strain, proximity to active sites/high X-ray cross-section atoms; Holton, 2009 ▸). Deconvoluting the individual roles of these parameters has been surprisingly challenging, with factors such as solvent accessibility currently under active investigation (Weik et al., 2000 ▸; Fioravanti et al., 2007 ▸; Gerstel et al., 2015 ▸). There are a number of cases where SRD manifestations have compromised the biological information extracted from MX-determined structures at much lower doses than the recommended 30 MGy limit, leading to false structural interpretations of protein mechanisms. Active-site residues appear to be particularly susceptible, particularly for photosensitive proteins and in instances where chemical strain is an intrinsic feature of the reaction mechanism. For instance, structure determination of the purple membrane protein bacteriorhodopsin required careful corrections for radiation-induced structural changes before the correct photosensitive intermediate states could be isolated (Matsui et al., 2002 ▸). The significant chemical strain required for catalysis within the active site of phosphoserine aminotransferase has been observed to diminish during X-ray exposure (Dubnovitsky et al., 2005 ▸). Since the majority of SRD studies to date have focused on proteins, much less is known about the effects of X-ray irradiation on the wider class of crystalline nucleoprotein complexes or how to correct for such radiation-induced structural changes. Understanding RD to such complexes is crucial, since DNA is rarely naked within a cell, instead dynamically interacting with proteins, facilitating replication, transcription, modification and DNA repair. As of early 2016, >5400 nucleoprotein complex structures have been deposited within the PDB, with 91% solved by MX. It is essential to understand how these increasingly complex macromolecular structures are affected by the radiation used to solve them. Nucleoproteins also represent one of the main targets of radiotherapy, and an insight into the damage mechanisms induced by X-ray irradiation could inform innovative treatments. When a typical macromolecular crystal is irradiated with ionizing X-rays, each photoelectron produced via interactions with both the macromolecule (direct damage) and solvent (indirect damage) can induce cascades of up to 500 secondary low-energy electrons (LEEs) that are capable of inducing further ionizations. Investigations on sub-ionization-level LEEs (0–15 eV) interacting with both dried and aqueous oligonucleotides (Alizadeh & Sanche, 2014 ▸; Simons, 2006 ▸) concluded that resonant electron attachment to DNA bases and the sugar-phosphate backbone could lead to the preferential cleavage of strong (∼4 eV, 385 kJ mol) sugar-phosphate C—O covalent bonds within the DNA backbone and then base-sugar N1—C bonds, eventually leading to single-strand breakages (SSBs; Ptasińska & Sanche, 2007 ▸). Electrons have been shown to be mobile at 77 K by electron spin resonance spectroscopy studies (Symons, 1997 ▸; Jones et al., 1987 ▸), with rapid electron quantum tunnelling and positive hole migration along the protein backbone and through stacked DNA bases indicated as a dominant mechanism by which oxidative and reductive damage localizes at distances from initial ionization sites at 100 K (O’Neill et al., 2002 ▸). The investigation of naturally forming nucleoprotein complexes circumvents the inherent challenges in making controlled comparisons of damage mechanisms between protein and nucleic acids crystallized separately. Recently, for a well characterized bacterial protein–DNA complex (C.Esp1396I; PDB entry 3clc; resolution 2.8 Å; McGeehan et al., 2008 ▸) it was concluded that over a wide dose range (2.1–44.6 MGy) the protein was far more susceptible to SRD than the DNA within the crystal (Bury et al., 2015 ▸). Only at doses above 20 MGy were precursors of phosphodiester-bond cleavage observed within AT-rich regions of the 35-mer DNA. For crystalline complexes such as C.Esp1396I, whether the protein is intrinsically more susceptible to X-ray-induced damage or whether the protein scavenges electrons to protect the DNA remains unclear in the absence of a non-nucleic acid-bound protein control obtained under exactly the same crystallization and data-collection conditions. To monitor the effects of nucleic acid binding on protein damage susceptibility, a crystal containing two protein molecules per asymmetric unit, only one of which was bound to RNA, is reported here (Fig. 1 ▸). Using newly developed methodology, we present a controlled SRD investigation at 1.98 Å resolution using a large (∼91 kDa) crystalline protein–RNA complex: trp RNA-binding attenuation protein (TRAP) bound to a 53 bp RNA sequence (GAGUU)10GAG (PDB entry 1gtf; Hopcroft et al., 2002 ▸). TRAP consists of 11 identical subunits assembled into a ring with 11-fold rotational symmetry. It binds with high affinity (K d ≃ 1.0 nM) to RNA segments containing 11 GAG/UAG triplets separated by two or three spacer nucleotides (Elliott et al., 2001 ▸) to regulate the transcription of tryptophan biosynthetic genes in Bacillus subtilis (Antson et al., 1999 ▸). In this structure, the bases of the G1-A2-G3 nucleotides form direct hydrogen bonds to the protein, unlike the U4-U5 nucleotides, which appear to be more flexible. Ten successive 1.98 Å resolution MX data sets were collected from the same TRAP–RNA crystal to analyse X-ray-induced structural changes over a large dose range (d 1 = 1.3 MGy to d 10 = 25.0 MGy). To avoid the previous necessity for visual inspection of electron-density maps to detect SRD sites, a computational approach was designed to quantify the electron-density change for each refined atom with increasing dose, thus providing a rapid systematic method for SRD study on such large multimeric complexes. By employing the high 11-fold structural symmetry within each TRAP macromolecule, this approach permitted a thorough statistical quantification of the RD effects of RNA binding to TRAP. As previously described (Hopcroft et al., 2002 ▸), the 53-base RNA (GAGUU)10GAG was synthesized by in vitro transcription with T7 RNA polymerase and gel-purified. TRAP from B. stearothermophilus was overexpressed in Escherichia coli and purified. TRAP–RNA crystals were prepared using a previously established hanging-drop crystallization protocol (Antson et al., 1999 ▸). By using a 2:1 molar ratio of TRAP to RNA, crystals successfully formed from the protein–RNA complex (∼15 mg ml) in a solution containing 70 mM potassium phosphate pH 7.8 and 10 mM l-tryptophan. The reservoir consisted of 0.2 M potassium glutamate, 50 mM triethanolamine pH 8.0, 10 mM MgCl2, 8–11% monomethyl ether PEG 2000. In order to accelerate crystallization, a further gradient was induced by adding 0.4 M KCl to the reservoir after 1.5 µl protein solution had been mixed with an equal volume of the reservoir solution. Wedge-shaped crystals of approximate length 70 µm (longest dimension) grew within 3 d and were vitrified and stored in liquid nitrogen immediately after growth. The cryosolution consisted of 12% monomethyl ether PEG 2000, 30 mM triethanolamine pH 8.0, 6 mM l-tryptophan, 0.1 M potassium glutamate, 35 mM potassium phosphate pH 7.8, 5 mM MgCl2 with 25% 2-methyl-2,4-pentanediol (MPD) included as a cryoprotectant. Data were collected at 100 K from a wedge-shaped TRAP–RNA crystal of approximate dimensions 70 × 20 × 40 µm (see Supplementary Fig. S2) on beamline ID14-4 at the ESRF using an incident wavelength of 0.940 Å (13.2 keV) and an ADSC Q315R mosaic CCD detector at 304.5 mm from the crystal throughout the data collection. The beam size was slitted to 0.100 mm (vertical) × 0.160 mm (horizontal), with a uniformly distributed profile, such that the crystal was completely bathed within the beam throughout data collection. Ten successive (1.98 Å resolution) 180° data sets (with Δφ = 1°) were collected over the same angular range from a TRAP–RNA crystal at 28.9% beam transmission. The TRAP–RNA macromolecule crystallized in space group C2, with unit-cell parameters a = 140.9, b = 110.9, c = 137.8 Å, α = γ = 90, β = 137.8° (the values quoted are for the first data set; see Supplementary Table S1 for subsequent values). For the first nine data sets the attenuated flux was recorded to be ∼5 × 10 photons s. A beam refill took place immediately before data set 10, requiring a flux-scale factor increase of 1.42 to be applied, based on the ratio of observed relative intensity I D/I 1 at data set 10 to that extrapolated from data set 9. RADDOSE-3D (Zeldin, Gerstel et al., 2013 ▸) was used to calculate the absorbed dose distribution during each data set (see input file; Supplementary Figs. S1 and S2). The crystal composition was calculated from the deposited TRAP–RNA structure (PDB entry 1gtf; Hopcroft et al., 2002 ▸). Crystal absorption coefficients were calculated in RADDOSE-3D using the concentration (mmol l) of solvent heavy elements from the crystallization conditions. The beam-intensity profile was modelled as a uniform (‘top-hat’) distribution. The diffraction-weighted dose (DWD) values (Zeldin, Brockhauser et al., 2013 ▸) are given in Supplementary Table S1. Each data set was integrated using iMosflm (Leslie & Powell, 2007 ▸) and was scaled using AIMLESS (Evans & Murshudov, 2013 ▸; Winn et al., 2011 ▸) using the same 5% R free set of test reflections for each data set. To phase the structure obtained from the first data set, molecular replacement was carried out with Phaser (McCoy et al., 2007 ▸), using an identical TRAP–RNA structure (PDB entry 1gtf; resolution 1.75 Å; Hopcroft et al., 2002 ▸) as a search model. The resulting TRAP–RNA structure (TR1) was refined using REFMAC5 (Murshudov et al., 2011 ▸), initially using rigid-body refinement, followed by repeated cycles of restrained, TLS and isotropic B-factor refinement, coupled with visual inspection in Coot (Emsley et al., 2010 ▸). TR1 was refined to 1.98 Å resolution, with a dimeric assembly of non-RNA-bound and RNA-bound TRAP rings within the asymmetric unit. Consistent with previous structures of the TRAP–RNA complex, the RNA sequence termini were not observed within the 2F o − F c map; the first spacer (U4) was then modelled at all 11 repeats around the TRAP ring and the second spacer (U5) was omitted from the final refined structure. For the later data sets, the observed structure-factor amplitudes from each separately scaled data set (output from AIMLESS) were combined with the phases of TR1 and the resulting higher-dose model was refined with phenix.refine (Adams et al., 2010 ▸) using only rigid-body and isotropic B-factor refinement. During this refinement, the TRAP–RNA complex and nonbound TRAP ring were treated as two separate rigid bodies within the asymmetric unit. Supplementary Table S1 shows the relevant summary statistics. The CCP4 program CAD was used to create a series of nine merged .mtz files combining observed structure-factor amplitudes for the first data set F obs(d 1) with each later data set F obs(d n) (for n = 2, …, 10). All later data sets were scaled against the initial low-dose data set in SCALEIT. For each data set an atom-tagged .map file was generated using the ATMMAP mode in SFALL (Winn et al., 2011 ▸). A full set of nine Fourier difference maps F obs(d n) − F obs(d 1) were calculated using FFT (Ten Eyck, 1973 ▸) over the full TRAP–RNA unit-cell dimensions, with the same grid-sampling dimensions as the atom-tagged .map file. All maps were cropped to the TRAP asymmetric unit in MAPMASK. Comparing the atom-tagged .map file and F obs(d n) − F obs(d 1) difference map at each dose, each refined atom was assigned a set of density-change values X. The maximum density-loss metric, D loss (units of e Å), was calculated to quantify the per-atom electron-density decay at each dose, assigned as the absolute magnitude of the most negative Fourier difference map voxel value in a local volume around each atom as defined by the set X. Model calculations were run for the simple amino acids glutamate and aspartate. In order to avoid decarboxylation at the C-terminus instead of the side chain on the C atom, the C-terminus of each amino acid was methylated. While the structures of the closed shell acids are well known, the same is not true of those in the oxidized state. The quantum-chemical calculations employed were chosen to provide a satisfactory description of the structure of such radical species and also provide a reliable estimation of the relative C—C(O2) bond strengths, which are otherwise not available. Structures of methyl-terminated (at the N- and C-termini) carboxylates were determined using analytic energy gradients with density functional theory (B3LYP functional; Becke, 1993 ▸) and a flexible basis set of polarized valence triple-zeta size with diffuse functions on the non-H atoms [6-311+G(d,p)] in the Gaussian 09 computational chemistry package (Frisch et al., 2009 ▸). The stationary points obtained were characterized as at least local minima by examination of the associated analytic Hessian. Effects of the medium were modelled using a dielectric cavity approach (Tomasi et al., 1999 ▸) parameterized for water. To quantify the exact effects of nucleic acid binding to a protein on SRD susceptibility, a high-throughput and automated pipeline was created to systematically calculate the electron-density change for every refined atom within the TRAP–RNA structure as a function of dose. This provides an atom-specific quantification of density–dose dynamics, which was previously lacking within the field. Previous studies have characterized SRD sites by reporting magnitudes of F obs(d n) − F obs(d 1) Fourier difference map peaks in terms of the sigma (σ) contour level (the number of standard deviations from the mean map electron-density value) at which peaks become visible. However, these σ levels depend on the standard deviation values of the map, which can deviate between data sets, and are thus unsuitable for quantitative comparison of density between different dose data sets. Instead, we use here a maximum density-loss metric (D loss), which is the per-atom equivalent of the magnitude of these negative Fourier difference map peaks in units of e Å. Large positive D loss values indicate radiation-induced atomic disordering reproducibly throughout the unit cells with respect to the initial low-dose data set. For each TRAP–RNA data set, the D loss metric successfully identified the recognized forms of protein SRD (Fig. 2 ▸ a), with clear Glu and Asp side-chain decarboxylation even in the first difference map calculated (3.9 MGy; Fig. 3 ▸ a). The main sequence of TRAP does not contain any Trp and Cys residues (and thus contains no disulfide bonds). The substrate Trp amino-acid ligands also exhibited disordering of the free terminal carboxyl groups at higher doses (Fig. 2 ▸ a); however, no clear Fourier difference peaks could be observed visually. Even for radiation-insensitive residues (e.g. Gly) the average D loss increases with dose: this is the effect of global radiation damage, since as dose increases the electron density associated with each refined atom becomes weaker as the atomic occupancy decreases (Fig. 2 ▸ b). Only Glu and Asp residues exhibit a rate of D loss increase that consistently exceeds the average decay (Fig. 2 ▸ b, dashed line) at each dose. Additionally, the density surrounding ordered solvent molecules was determined to significantly diminish with increasing dose (Fig. 2 ▸ b). The rate of D loss (attributed to side-chain decarboxylation) was consistently larger for Glu compared with Asp residues over the large dose range (Fig. 2 ▸ b and Supplementary Fig. S3); this observation is consistent with our calculations on model systems (see above) that suggest that, without considering differential hydrogen-bonding environments, CO2 loss is more exothermic by around 8 kJ mol from oxidized Glu residues than from their Asp counterparts. Visual inspection of Fourier difference maps illustrated the clear lack of RNA electron-density degradation with increasing dose compared with the obvious protein damage manifestations (Figs. 3 ▸ b and 3 ▸ c). Only at the highest doses investigated (>20 MGy) was density loss observed at the RNA phosphate and C—O bonds of the phosphodiester backbone. However, the median D loss was lower by a factor of >2 for RNA P atoms than for Glu and Asp side-chain groups at 25.0 MGy (Supplementary Fig. S4), and furthermore could not be numerically distinguished from Gly C atoms within TRAP, which are not radiation-sensitive at the doses tested here (Supplementary Fig. S3). For the large number of acidic residues per TRAP ring (four Asp and six Glu residues per protein monomer), a strong dependence of decarboxylation susceptibility on local environment was observed (Fig. 4 ▸). For each Glu C or Asp C atom, D loss provided a direct measure of the rate of side-chain carboxyl-group disordering and subsequent decarboxylation. For acidic residues with no differing interactions between nonbound and bound TRAP (Fig. 4 ▸ a), similar damage was apparent between the two rings within the asymmetric unit, as expected. However, TRAP residues directly on the RNA-binding interfaces exhibited greater damage accumulation in nonbound TRAP (Fig. 4 ▸ b), and for residues at the ring–ring interfaces (where crystal contacts were detected) bound TRAP exhibited enhanced SRD accumulation (Fig. 4 ▸ c). Three acidic residues (Glu36, Asp39 and Glu42) are involved in RNA interactions within each of the 11 TRAP ring subunits, and Fig. 5 ▸ shows their density changes with increasing dose. Hotelling’s T-squared test (the multivariate counterpart of Student’s t-test) was used to reject the null hypothesis that the means of the D loss metric were equal for the bound and nonbound groups in Fig. 5 ▸. A significant reduction in D loss is seen for Glu36 in RNA-bound compared with nonbound TRAP, indicative of a lower rate of side-chain decarboxylation (Fig. 5 ▸ a; p = 6.06 × 10). For each TRAP ring subunit, the Glu36 side-chain carboxyl group accepts a pair of hydrogen bonds from the two N atoms of the G3 RNA base. In our analysis, Asp39 in the TRAP–(GAGUU)10GAG structure appears to exhibit two distinct hydrogen bonds to the G1 base within each of the 11 TRAP–RNA interfaces, as does Glu36 to G3; however, the reduction in density disordering upon RNA binding is far less significant for Asp39 than for Glu36 (Fig. 5 ▸ b, p = 0.0925). One oxygen (O) of Glu42 appears to form a hydrogen bond to a nearby water within each TRAP RNA-binding pocket, with the other (O) being involved in a salt-bridge interaction with Arg58 (Hopcroft et al., 2002 ▸; Antson et al., 1999 ▸). Salt-bridge interactions have previously been suggested to reduce the glutamate decarboxylation rate within the large (∼62.4 kDa) myrosinase protein structure (Burmeister, 2000 ▸). A significant difference was observed between the D loss dynamics for the nonbound/bound Glu42 O atoms (Fig. 5 ▸ c; p = 0.007) but not for the Glu42 O atoms (Fig. 5 ▸ d; p = 0.239), indicating that the stabilizing strength of this salt-bridge interaction was conserved upon RNA binding and that the water-mediated hydrogen bond had a greater relative susceptibility to atomic disordering in the absence of RNA. The density-change dynamics were statistically indistinguishable between bound and nonbound TRAP for each Glu42 carboxyl group C atom (p = 0.435), indicating that upon RNA binding the conserved salt-bridge interaction ultimately dictated the overall Glu42 decarboxylation rate. The RNA-stabilizing effect was not restricted to radiation-sensitive acidic residues. The side chain of Phe32 stacks against the G3 base within the 11 TRAP RNA-binding interfaces (Antson et al., 1999 ▸). With increasing dose, the D loss associated with the Phe32 side chain was significantly reduced upon RNA binding (Fig. 5 ▸ e; Phe32 C; p = 0.0014), an indication that radiation-induced conformation disordering of Phe32 had been reduced. The extended aliphatic Lys37 side chain stacks against the nearby G1 base, making a series of nonpolar contacts within each RNA-binding interface. The D loss for Lys37 side-chain atoms was also reduced when stacked against the G1 base (Fig. 5 ▸ f; p = 0.0243 for Lys37 C atoms). Representative Phe32 and Lys37 atoms were selected to illustrate these trends. Here, MX radiation-induced specific structural changes within the large TRAP–RNA assembly over a large dose range (1.3–25.0 MGy) have been analysed using a high-throughput quantitative approach, providing a measure of the electron-density distribution for each refined atom with increasing dose, D loss. Compared with previous studies, the results provide a further step in the detailed characterization of SRD effects in MX. Our methodology, which eliminated tedious and error-prone visual inspection, permitted the determination on a per-atom basis of the most damaged sites, as characterized by F obs(d n) − F obs(d 1) Fourier difference map peaks between successive data sets collected from the same crystal. Here, it provided the precision required to quantify the role of RNA in the damage susceptibilities of equivalent atoms between RNA-bound and nonbound TRAP, but it is applicable to any MX SRD study. The RNA was found to be substantially more radiation-resistant than the protein, even at the highest doses investigated (∼25.0 MGy), which is in strong concurrence with our previous SRD investigation of the C.Esp1396I protein–DNA complex (Bury et al., 2015 ▸). Consistent with that study, at high doses of above ∼20 MGy, F obs(d n) − F obs(d 1) map density was detected around P, O3′ and O5′ atoms of the RNA backbone, with no significant difference density localized to RNA ribose and basic subunits. RNA backbone disordering thus appears to be the main radiation-induced effect in RNA, with the protein–base interactions maintained even at high doses (>20 MGy). The U4 phosphate exhibited marginally larger D loss values above 20 MGy than G1, A2 and G3 (Supplementary Fig. S4). Since U4 is the only refined nucleotide not to exhibit significant base–protein interactions around TRAP (with a water-mediated hydrogen bond detected in only three of the 11 subunits and a single Arg58 hydrogen bond suggested in a further four subunits), this increased U4 D loss can be explained owing to its greater flexibility. At 25.0 MGy, the magnitude of the RNA backbone D loss was of the same order as for the radiation-insensitive Gly C atoms and on average less than half that of the acidic residues of the protein (Supplementary Fig. S3). Consequently, no clear single-strand breaks could be located, and since RNA-binding within the current TRAP–(GAGUU)10GAG complex is mediated predominantly through base–protein interactions, the biological integrity of the RNA complex was dictated by the rate at which protein decarboxylation occurred. RNA interacting with TRAP was shown to offer significant protection against radiation-induced structural changes. Both Glu36 and Asp39 bind directly to RNA, each through two hydrogen bonds to guanine bases (G3 and G1, respectively). However, compared with Asp39, Glu36 is strikingly less decarboxylated when bound to RNA (Fig. 4 ▸). This is in good agreement with previous mutagenesis and nucleoside analogue studies (Elliott et al., 2001 ▸), which indicated that the G1 nucleotide does not bind to TRAP as strongly as do A2 and G3, and plays little role in the high RNA-binding affinity of TRAP (K d ≃ 1.1 ± 0.4 nM). For Glu36 and Asp39, no direct quantitative correlation could be established between hydrogen-bond length and D loss (linear R of <0.23 for all doses; Supplementary Fig. S5). Thus, another factor must be responsible for this clear reduction in Glu36 CO2 decarboxylation in RNA-bound TRAP. The Glu36 carboxyl side chain also potentially forms hydrogen bonds to His34 and Lys56, but since these interactions are conserved irrespective of G3 nucleotide binding, this cannot directly account for the stabilization effect on Glu36 in RNA-bound TRAP. Radiation-induced decarboxylation has been proposed to be mediated by preferential positive-hole migration to the side-chain carboxyl group, with rapid proton transfer trapping the hole at the carboxyl group (Burmeister, 2000 ▸; Symons, 1997 ▸):where the forward rate is K 1 and the backward rate is K −1, where the forward rate is K 2. When bound to RNA, the average solvent-accessible area of the Glu36 side-chain O atoms is reduced from ∼15 to 0 Å. We propose that with no solvent accessibility Glu36 decarboxylation is inhibited, since the CO2-formation rate K 2 is greatly reduced, and suggest that steric hindrance prevents each radicalized Glu36 CO2 group from achieving the planar conformation required for complete dissociation from TRAP. The electron-recombination rate K −1 remains high, however, owing to rapid electron migration through the protein–RNA complex to refill the Glu36 positive hole (the precursor for Glu decarboxylation). Upon RNA binding, the Asp39 side-chain carboxyl group solvent-accessible area changes from ∼75 to 35 Å, still allowing a high CO2-formation rate K 2. Previous studies have reported inconsistent results concerning the dependence of the acidic residue decarboxylation rate on solvent accessibility (Weik et al., 2000 ▸; Fioravanti et al., 2007 ▸; Gerstel et al., 2015 ▸). The prevalence of radical attack from solvent channels surrounding the protein in the crystal is a questionable cause, considering previous observations indicating that the strongly oxidizing hydroxyl radical is immobile at 100 K (Allan et al., 2013 ▸; Owen et al., 2012 ▸). Furthermore, the suggested electron hole-trapping mechanism which induces decarboxylation within proteins at 100 K has no clear mechanistic dependence on the solvent-accessible area of each carboxyl group. By comparing equivalent acidic residues with and without RNA, we have now deconvoluted the role of solvent accessibility from other local protein environment factors, and thus propose a suitable mechanism by which exceptionally low solvent accessibility can reduce the rate of decarboxylation. Overall, no direct correlation between solvent accessibility and decarboxylation susceptibility was observed, but it is very clear that inaccessible residues are protected. Apart from these RNA-binding interfaces, RNA binding was seen to enhance decarboxylation for residues Glu50, Glu71 and Glu73, all of which are involved in crystal contacts between TRAP rings (Fig. 4 ▸ c). However, for each of these residues the exact crystal contacts are not preserved between bound and nonbound TRAP or even between monomers within one TRAP ring. For example, in bound TRAP, Glu73 hydrogen-bonds to a nearby lysine on each of the 11 subunits, whereas in nonbound TRAP no such interaction exists and Glu73 interacts with a variable number of refined waters in each subunit. Thus, the dependence of decarboxylation rates on these interactions could not be established. Radiation-induced side-chain conformational changes have been poorly characterized in previous SRD investigations owing to their strong dependence on packing density and geometric strain. Such structural changes are known to have significant roles within enzymatic pathways, and experimenters must be aware of these possible confounding factors when assigning true functional mechanisms using MX. Our results show that RNA binding to TRAP physically stabilizes non-acidic residues within the TRAP macromolecule, most notably Lys37 and Phe32, which stack against the G1 and G3 bases, respectively. It has been suggested (Burmeister, 2000 ▸) that Tyr residues can lose their aromatic –OH group owing to radiation-induced effects; however, no energetically favourable pathway for –OH cleavage exists and this has not been detected in aqueous radiation-chemistry studies. In TRAP, D loss increased at a similar rate for both the Tyr O atoms and aromatic ring atoms, suggesting that full ring conformational disordering is more likely. Indeed, no convincing reproducible Fourier difference peaks above the background map noise were observed around any Tyr terminal –OH groups. The RNA-stabilization effects on protein are observed at short ranges and are restricted to within the RNA-binding interfaces around the TRAP ring. For example, Asp17 is located ∼6.8 Å from the G1 base, outside the RNA-binding interfaces, and has indistinguishable C atom D loss dose-dynamics between RNA-bound and nonbound TRAP (p > 0.9). An increase in the dose at which functionally important residues remain intact has biological ramifications for understanding the mechanisms at which ionizing radiation damage is mitigated within naturally forming DNA–protein and RNA–protein complexes. Observations of lower protein radiation-sensitivity in DNA-bound forms have been recorded in solution at RT at much lower doses (∼1 kGy) than those used for typical MX experiments [e.g. an oestrogen response element–receptor complex (Stísová et al., 2006 ▸) and a DNA glycosylase and its abasic DNA target site (Gillard et al., 2004 ▸)]. In these studies, the main damaging species is predicted to be the oxidizing hydroxyl radical produced through solvent irradiation, which is known to add to double covalent bonds within both DNA and RNA bases to induce strand breaks and base modification (Spotheim-Maurizot & Davídková, 2011 ▸; Chance et al., 1997 ▸). It was suggested that physical screening of DNA by protein shielded the DNA–protein interaction sites from radical damage, yielding an extended life-dose for the nucleoprotein complex compared with separate protein and DNA constituents at RT. However, in the current MX study at 100 K, the main damaging species are believed to be migrating LEEs and holes produced directly within the protein–RNA components or in closely associated solvent. The results presented here suggest that biologically relevant nucleoprotein complexes also exhibit prolonged life-doses under the effect of LEE-induced structural changes, involving direct physical protection of key RNA-binding residues. Such reduced radiation-sensitivity in this case ensures that the interacting protein remains bound long enough to the RNA to complete its function, even whilst exposed to ionizing radiation. Within the nonbound TRAP macromolecule, the acidic residues within the unoccupied RNA-binding interfaces (Asp39, Glu36, Glu42) are notably amongst the most susceptible residues within the asymmetric unit (Fig. 4 ▸). When exposed to X-rays, these residues will be preferentially damaged by X-rays and subsequently reduce the affinity with which TRAP binds to RNA. Within the cellular environment, this mechanism could reduce the risk that radiation-damaged proteins might bind to RNA, thus avoiding the detrimental introduction of incorrect DNA-repair, transcriptional and base-modification pathways. The Python scripts written to calculate the per atom D loss metric are available from the authors on request. The following references are cited in the Supporting Information for this article: Chen et al. (2010 ▸).
PMC4792962	A unified mechanism for proteolysis and autocatalytic activation in the 20S proteasome	Biogenesis of the 20S proteasome is tightly regulated. The N-terminal propeptides protecting the active-site threonines are autocatalytically released only on completion of assembly. However, the trigger for the self-activation and the reason for the strict conservation of threonine as the active site nucleophile remain enigmatic. Here we use mutagenesis, X-ray crystallography and biochemical assays to suggest that Lys33 initiates nucleophilic attack of the propeptide by deprotonating the Thr1 hydroxyl group and that both residues together with Asp17 are part of a catalytic triad. Substitution of Thr1 by Cys disrupts the interaction with Lys33 and inactivates the proteasome. Although a Thr1Ser mutant is active, it is less efficient compared with wild type because of the unfavourable orientation of Ser1 towards incoming substrates. This work provides insights into the basic mechanism of proteolysis and propeptide autolysis, as well as the evolutionary pressures that drove the proteasome to become a threonine protease.The 20S proteasome core particle (CP) is the key non-lysosomal protease of eukaryotic cells. Its seven different α and seven different β subunits assemble into four heptameric rings that are stacked on each other to form a hollow cylinder. While the inactive α subunits build the two outer rings, the β subunits form the inner rings. Only three out of the seven different β subunits, namely β1, β2 and β5, bear N-terminal proteolytic active centres, and before CP maturation these are protected by propeptides123. In the last stage of CP biogenesis, the prosegments are autocatalytically removed through nucleophilic attack by the active site residue Thr1 on the preceding peptide bond involving Gly(-1)45. Release of the propeptides creates a functionally active CP that cleaves proteins into short peptides. Although the chemical nature of the substrate-binding channel and hence substrate preferences are unique to each of the distinct active β subunits67, all active sites employ an identical reaction mechanism to hydrolyse peptide bonds2. Nucleophilic attack of Thr1O on the carbonyl carbon atom of the scissile peptide bond creates a first cleavage product and a covalent acyl-enzyme intermediate. Hydrolysis of this complex by the addition of a nucleophilic water molecule regenerates the enzyme and releases the second peptide fragment89. The proteasome belongs to the family of N-terminal nucleophilic (Ntn) hydrolases10, and the free N-terminal amine group of Thr1 was proposed to deprotonate the Thr1 hydroxyl group to generate a nucleophilic Thr1O for peptide-bond cleavage2911. This mechanism, however, cannot explain autocatalytic precursor processing because in the immature active sites, Thr1N is part of the peptide bond with Gly(-1), the bond that needs to be hydrolysed. An alternative candidate for deprotonating the Thr1 hydroxyl group is the side chain of Lys33 as it is within hydrogen-bonding distance to Thr1OH (2.7 Å). In principle it could function as the general base during both autocatalytic removal of the propeptide and protein substrate cleavage. Here we provide experimental evidences for this distinct view of the proteasome active-site mechanism. Data from biochemical and structural analyses of proteasome variants with mutations in the β5 propeptide and the active site strongly support the model and deliver novel insights into the structural constraints required for the autocatalytic activation of the proteasome. Furthermore, we determine the advantages of Thr over Cys or Ser as the active-site nucleophile using X-ray crystallography together with activity and inhibition assays. Proteasome-mediated degradation of cell-cycle regulators and potentially toxic misfolded proteins is required for the viability of eukaryotic cells8. Inactivation of the active site Thr1 by mutation to Ala has been used to study substrate specificity and the hierarchy of the proteasome active sites1412131415. Yeast strains carrying the single mutations β1-T1A or β2-T1A, or both, are viable, even though one or two of the three distinct catalytic β subunits are disabled and carry remnants of their N-terminal propeptides4 (Table 1). These results indicate that the β1 and β2 proteolytic activities are not essential for cell survival. By contrast, the T1A mutation in subunit β5 has been reported to be lethal or nearly so113. Viability is restored if the β5-T1A subunit has its propeptide (pp) deleted but expressed separately in trans (β5-T1A pp trans), although substantial phenotypic impairment remains11516 (Table 1). Our present crystallographic analysis of the β5-T1A pp trans mutant demonstrates that the mutation per se does not structurally alter the catalytic active site and that the trans-expressed β5 propeptide is not bound in the β5 substrate-binding channel (Supplementary Fig. 1a). The extremely weak growth of the β5-T1A mutant pp cis described by Chen and Hochstrasser1 compared with the inviability reported by Heinemeyer et al.13 prompted us to analyse this discrepancy. Sequencing of the plasmids, testing them in both published yeast strain backgrounds and site-directed mutagenesis revealed that the β5-T1A mutant pp cis is viable, but suffers from a marked growth defect that requires extended incubation of 4–5 days for initial colony formation (Table 1 and Supplementary Methods). We also identified an additional point mutation K81R in subunit β5 that was present in the allele used in ref. 1. This single amino-acid exchange is located at the interface of the subunits α4, β4 and β5 (Supplementary Fig. 1b) and might weakly promote CP assembly by enhancing inter-subunit contacts. The slightly better growth of the β5-T1A-K81R mutant allowed us to solve the crystal structure of a yeast proteasome (yCP) with the β5-T1A mutation, which is discussed in the following section (for details see Supplementary Note 1). In the final steps of proteasome biogenesis, the propeptides are autocatalytically cleaved from the mature β-subunit domains1. For subunit β1, this process was previously inferred to require that the propeptide residue at position (-2) of the subunit precursor occupies the S1 specificity pocket of the substrate-binding channel formed by amino acid 45 (for details see Supplementary Note 2)5. Furthermore, it was observed that the prosegment forms an antiparallel β-sheet in the active site, and that Gly(-1) adopts a γ-turn conformation, which by definition is characterized by a hydrogen bond between Leu(-2)O and Thr1NH (ref. 5). Here we again analysed the β1-T1A mutant crystallographically but in addition determined the structures of the β2-T1A single and β1-T1A-β2-T1A double mutants (Protein Data Bank (PDB) entry codes are provided in Supplementary Table 1). In subunit β1, we found that Gly(-1) indeed forms a sharp turn, which relaxes on prosegment cleavage (Fig. 1a and Supplementary Fig. 2a). However, the γ-turn conformation and the associated hydrogen bond initially proposed is for geometric and chemical reasons inappropriate and would not perfectly position the carbonyl carbon atom of Gly(-1) for nucleophilic attack by Thr1. Regarding the β2 propeptide, Thr(-2) occupies the S1 pocket but is less deeply anchored compared with Leu(-2) in β1, which might be due to the rather large β2-S1 pocket created by Gly45. Thr(-2) positions Gly(-1)O via hydrogen bonding (∼2.8 Å) in a perfect trajectory for the nucleophilic attack by Thr1O (Fig. 1b and Supplementary Fig. 2b). Next, we examined the position of the β5 propeptide in the β5-T1A-K81R mutant. Surprisingly, Gly(-1) is completely extended and forces the histidine side chain at position (-2) to occupy the S2 instead of the S1 pocket, thereby disrupting the antiparallel β-sheet. Nonetheless, the carbonyl carbon of Gly(-1) would be ideally placed for nucleophilic attack by Thr1O (Fig. 1c and Supplementary Fig. 2c,d). As the K81R mutation is located far from the active site (Thr1C–Arg81C: 24 Å), any influence on propeptide conformation can be excluded. Instead, the plasticity of the β5 S1 pocket caused by the rotational flexibility of Met45 might prevent stable accommodation of His(-2) in the S1 site and thus also promote its immediate release after autolysis. Processing of β-subunit precursors requires deprotonation of Thr1OH; however, the general base initiating autolysis is unknown. Remarkably, eukaryotic proteasomal β5 subunits bear a His residue in position (-2) of the propeptide (Supplementary Fig. 3a). As histidine commonly functions as a proton shuttle in the catalytic triads of serine proteases17, we investigated the role of His(-2) in processing of the β5 propeptide by exchanging it for Asn, Lys, Phe and Ala. All yeast mutants were viable at 30 °C, but suffered from growth defects at 37 °C with the H(-2)N and H(-2)F mutants being most affected (Supplementary Fig. 3b and Table 1). In agreement, the chymotrypsin-like (ChT-L) activity of H(-2)N and H(-2)F mutant yCPs was impaired in situ and in vitro (Supplementary Fig. 3c). Structural analyses revealed that the propeptides of all mutant yCPs shared residual 2FO–FC electron densities. Gly(-1) and Phe/Lys(-2) were visualized at low occupancy, while Ala/Asn(-2) could not be assigned. This observation indicates a mixture of processed and unprocessed β5 subunits and partially impaired autolysis18, thereby excluding any essential role of residue (-2) as the general base. Next, we examined the effect of residue (-2) on the orientation of the propeptide by creating mutants that combine the T1A (K81R) mutation(s) with H(-2)L, H(-2)T or H(-2)A substitutions. Leu(-2) is encoded in the yeast β1 subunit precursor (Supplementary Fig. 3a); Thr(-2) is generally part of β2-propeptides (Supplementary Fig. 3a); and Ala(-2) was expected to fit the β5-S1 pocket without inducing conformational changes of Met45, allowing it to accommodate ‘β1-like' propeptide positioning. As expected from β5-T1A mutants, the yeasts show severe growth phenotypes, with minor variations (Supplementary Fig. 4a and Table 1). We determined crystal structures of the β5-H(-2)L-T1A, β5-H(-2)T-T1A and the β5-H(-2)A-T1A-K81R mutants (Supplementary Table 1). For the β5-H(-2)A-T1A-K81R variant, only the residues Gly(-1) and Ala(-2) could be visualized, indicating that Ala(-2) leads to insufficient stabilization of the propeptide in the substrate-binding channel (Supplementary Fig. 4d). By contrast, the prosegments of the β5-H(-2)L-T1A and the β5-H(-2)T-T1A mutants were significantly better resolved in the 2FO–FC electron-density maps yet not at full occupancy (Supplementary Fig. 4b,c and Supplementary Table 1), suggesting that the natural propeptide bearing His(-2) is most favourable. Nevertheless, both Leu(-2) and Thr(-2) were found to occupy the S1 specificity pocket formed by Met45 (Fig. 2a,b and Supplementary Fig. 4f–h). This result proves that the naturally occurring His(-2) of the β5 propeptide does not stably fit into the S1 site. Since Gly(-1) adopts the same position in both wild-type (WT) and mutant β5 propeptides, and since in all cases its carbonyl carbon is perfectly placed for nucleophilic attack by Thr1O (Fig. 2b), we propose that neither binding of residue (-2) to the S1 pocket nor formation of the antiparallel β-sheet is essential for autolysis of the propeptide. Next, we determined the crystal structure of a chimeric yCP having the yeast β1-propeptide replaced by its β5 counterpart18. Although we observed fragments of 2FO–FC electron density in the β1 active site, the data were not interpretable. Bearing in mind that in contrast to Thr(-2) in β2, Leu(-2) in subunit β1 is not conserved among species (Supplementary Fig. 3a), we created a β2-T(-2)V proteasome mutant. As proven by the β2-T1A crystal structures, Thr(-2) hydrogen bonds to Gly(-1)O. Although this interaction was not observed for the β5-H(-2)T-T1A mutant (Fig. 2c and Supplementary Fig. 4c,i), exchange of Thr(-2) by Val in β2, a conservative mutation regarding size but drastic with respect to polarity, was found to inhibit maturation of this subunit (Fig. 2d and Supplementary Fig. 4e,j). Notably, the 2FO–FC electron-density map displays a different orientation for the β2 propeptide than has been observed for the β2-T1A proteasome. In particular, Val(-2) is displaced from the S1 site and Gly(-1) is severely shifted (movement of the carbonyl oxygen atom of 3.8 Å), thereby preventing nucleophilic attack of Thr1 (Fig. 2d and Supplementary Fig. 4j,k). These results further confirm that correct positioning of the active-site residues and Gly(-1) is decisive for the maturation of the proteasome. Proton shuttling from the proteasomal active site Thr1OH to Thr1NH2 via a nucleophilic water molecule was suggested to initiate peptide-bond hydrolysis2910. However, in the immature particle Thr1NH2 is blocked by the propeptide and cannot activate Thr1O. Instead, Lys33NH2, which is in hydrogen-bonding distance to Thr1O (2.7 Å) in all catalytically active β subunits (Fig. 3a,b)29, was proposed to serve as the proton acceptor19. Owing to its likely protonation at neutral pH, however, it was assumed not to act as the general base259. A proposed catalytic tetrad model involving Thr1OH, Thr1NH2, Lys33NH2 and Asp17O, as well as a nucleophilic water molecule as the proton shuttle appeared to accommodate all possible views of the proteasomal active site8920. Twenty years later, with a plethora of yCP X-ray structures in hand, we decided to re-analyse the active site of the proteasome and to resolve the uncertainty regarding the nature of the general base. Mutation of β5-Lys33 to Ala causes a strongly deleterious phenotype, and previous structural and biochemical analyses confirmed that this is caused by failure of propeptide cleavage, and consequently, lack of ChT-L activity1413 (Fig. 4a, Supplementary Fig. 3b and Table 1; for details see Supplementary Note 1). The phenotype of the β5-K33A mutant was however less pronounced than for the β5-T1A-K81R yeast (Fig. 4a). This discrepancy in growth was traced to an additional point mutation L(-49)S in the β5-propeptide of the β5-K33A mutant (see also Supplementary Note 1). Structural comparison of the β5-L(-49)S-K33A and β5-T1A-K81R active sites revealed that mutation of Lys33 to Ala creates a cavity that is filled with Thr1 and the remnant propeptide. This structural alteration destroys active-site integrity and abolishes catalytic activity of the β5 active site4 (Supplementary Fig. 5a). Additional proof for the key function of Lys33 was obtained from the β5-K33A mutant, with the propeptide expressed separately from the main subunit (pp trans)15. The Thr1 N terminus of this mutant is not blocked by the propeptide, yet its catalytic activity is reduced by ∼83% (Supplementary Fig. 6b). Consistent with this, the crystal structure of the β5-K33A pp trans mutant in complex with carfilzomib only showed partial occupancy of the ligand at the β5 active sites (Supplementary Fig. 5b and Supplementary Table 1). Since no acetylation of the Thr1 N terminus was observed for the β5-K33A pp trans apo crystal structure416, the reduced reactivity towards substrates and inhibitors indicates that Lys33NH2, rather than Thr1NH2, deprotonates and activates Thr1OH. Furthermore, the crystal structure of the β5-K33A pp trans mutant without inhibitor revealed that Thr1O strongly coordinates a well-defined water molecule (∼2 Å; Fig. 3c and Supplementary Fig. 5c,d). This water hydrogen bonds also to Arg19O (∼3.0 Å) and Asp17O (∼3.0 Å), and thereby presumably enables residual activity of the mutant. Remarkably, the solvent molecule occupies the position normally taken by Lys33NH2 in the WT proteasome structure (Fig. 3c), further corroborating the essential role of Lys33 as the general base for autolysis and proteolysis. Conservative substitution of Lys33 by Arg delays autolysis of the β5 precursor and impairs yeast growth (for details see Supplementary Note 1). While Thr1 occupies the same position as in WT yCPs, Arg33 is unable to hydrogen bond to Asp17, thereby inactivating the β5 active site24 (Supplementary Fig. 5e). The conservative mutation of Asp17 to Asn in subunit β5 of the yCP also provokes a severe growth defect (Supplementary Note 1, Supplementary Fig. 6a and Table 1). Notably, only with the additional point mutation L(-49)S present in the β5 propeptide could we purify a small amount of the β5-D17N mutant yCP. As determined by crystallographic analysis, this mutant β5 subunit was partially processed (Table 1) but displayed impaired reactivity towards the proteasome inhibitor carfilzomib compared with the subunits β1 and β2, and with WT β5 (Supplementary Fig. 7a). In contrast to the cis-construct, expression of the β5 propeptide in trans allowed straightforward isolation and crystallization of the D17N mutant proteasome. The ChT-L activity of the β5-D17N pp in trans CP towards the canonical β5 model substrates N-succinyl-Leu-Leu-Val-Tyr-7-amino-4-methylcoumarin (Suc-LLVY-AMC) and carboxybenzyl-Gly-Gly-Leu-para-nitroanilide (Z-GGL-pNA) was severely reduced (Supplementary Fig. 6b), confirming that Asp17 is of fundamental importance for the catalytic activity of the mature proteasome. Even though the β5-D17N pp trans yCP crystal structure appeared identical to the WT yCP (Supplementary Fig. 7b), the co-crystal structure with the α′, β′ epoxyketone inhibitor carfilzomib visualized only partial occupancy of the ligand in the β5 active site (Supplementary Fig. 7a). This observation is consistent with a strongly reduced reactivity of β5-Thr1 and the crystal structure of the β5-D17N pp cis mutant in complex with carfilzomib. Autolysis and residual catalytic activity of the β5-D17N mutants may originate from the carbonyl group of Asn17, which albeit to a lower degree still can polarize Lys33 for the activation of Thr1. In agreement, an E17A mutant in the proteasomal β-subunit of the archaeon Thermoplasma acidophilum prevents autolysis and catalysis21. Strikingly, although the X-ray data on the β5-D17N mutant with the propeptide expressed in cis and in trans looked similar, there was a pronounced difference in their growth phenotypes observed (Supplementary Fig. 6a and Supplementary Fig. 7b). On the basis of these results, we propose that CPs from all domains of life use a catalytic triad consisting of Thr1, Lys33 and Asp/Glu17 for both autocatalytic precursor processing and proteolysis (Fig. 3d). This model is also consistent with the fact that no defined water molecule is observed in the mature WT proteasomal active site that could shuttle the proton from Thr1O to Thr1NH2. To explore this active-site model further, we exchanged the conserved Asp166 residue for Asn in the yeast β5 subunit. Asp166O is hydrogen-bonded to Thr1NH2 via Ser129OH and Ser169OH, and therefore was proposed to be involved in catalysis2. The β5-D166N pp cis yeast mutant is significantly impaired in growth and its ChT-L activity is drastically reduced (Supplementary Fig. 6a,b and Table 1). X-ray data on the β5-D166N mutant indicate that the β5 propeptide is hydrolysed, but due to reorientation of Ser129OH, the interaction with Asn166O is disrupted (Supplementary Fig. 8a). Instead, a water molecule is bound to Ser129OH and Thr1NH2 (Supplementary Fig. 8b), which may enable precursor processing. The hydrogen bonds involving Ser169OH are intact and may account for residual substrate turnover. Soaking the β5-D166N crystals with carfilzomib and MG132 resulted in covalent modification of Thr1 at high occupancy (Supplementary Fig. 8c). In the carfilzomib complex structure, Thr1O and Thr1N incorporate into a morpholine ring structure and Ser129 adopts its WT-like orientation. In the MG132-bound state, Thr1N is unmodified, and we again observe that Ser129 is hydrogen-bonded to a water molecule instead of Asn166. Whereas Asn can to some degree replace Asp166 due to its carbonyl group in the side chain, Ala at this position was found to prevent both autolysis and catalysis21. These results suggest that Asp166 and Ser129 function as a proton shuttle and affect the protonation state of Thr1N during autolysis and catalysis. Mutation of Thr1 to Cys inactivates the 20S proteasome from the archaeon T. acidophilum21. In yeast, this mutation causes a strong growth defect (Fig. 4a and Table 1), although the propeptide is hydrolysed, as shown here by its X-ray structure. In one of the two β5 subunits, however, we found the cleaved propeptide still bound in the substrate-binding channel (Fig. 4c). His(-2) occupies the S2 pocket like observed for the β5-T1A-K81R mutant, but in contrast to the latter, the propeptide in the T1C mutant adopts an antiparallel β-sheet conformation as known from inhibitors like MG132 (Fig. 4c–e and Supplementary Fig. 9b). On the basis of the phenotype of the T1C mutant and the propeptide remnant identified in its active site, we suppose that autolysis is retarded and may not have been completed before crystallization. Owing to the unequal positions of the two β5 subunits within the CP in the crystal lattice, maturation and propeptide displacement may occur at different timescales in the two subunits. Despite propeptide hydrolysis, the β5-T1C active site is catalytically inactive (Fig. 4b and Supplementary Fig. 9a). In agreement, soaking crystals with the CP inhibitors bortezomib or carfilzomib modifies only the β1 and β2 active sites, while leaving the β5-T1C proteolytic centres unmodified even though they are only partially occupied by the cleaved propeptide remnant. Moreover, the structural data reveal that the thiol group of Cys1 is rotated by 74° with respect to the hydroxyl side chain of Thr1 (Fig. 4f and Supplementary Fig. 9b). This presumably results from the larger radius of the sulfur atom compared with oxygen. Consequently, the hydrogen bond bridging the active-site nucleophile and Lys33 in WT CPs is broken with Cys1. Notably, the 2FO–FC electron-density map of the T1C mutant also indicates that Lys33NH2 is disordered. Together, these observations suggest that efficient peptide-bond hydrolysis requires that Lys33NH2 hydrogen bonds to the active site nucleophile. All proteasomes strictly employ threonine as the active-site residue instead of serine. To investigate the reason for this singularity, we analysed a β5-T1S mutant, which is viable but suffers from growth defects (Fig. 4a and Table 1). Activity assays with the β5-specific substrate Suc-LLVY-AMC demonstrated that the ChT-L activity of the T1S mutant is reduced by 40–45% compared with WT proteasomes depending on the incubation temperature (Fig. 4b and Supplementary Fig. 9c). By contrast, turnover of the substrate Z-GGL-pNA, used to monitor ChT-L activity in situ but in a less quantitative fashion, is not detectably impaired (Supplementary Fig. 9a). Crystal structure analysis of the β5-T1S mutant confirmed precursor processing (Fig. 4g), and ligand-complex structures with bortezomib and carfilzomib unambiguously corroborated the reactivity of Ser1 (Fig. 5). However, the apo crystal structure revealed that Ser1O is turned away from the substrate-binding channel (Fig. 4g). Compared with Thr1O in WT CP structures, Ser1O is rotated by 60°. This renders it unavailable for direct nucleophilic attack onto incoming substrates and first requires its reorientation, which is expected to delay substrate turnover. Because both conformations of Ser1O are hydrogen-bonded to Lys33NH2 (Fig. 4h), the relay system is capable of hydrolysing peptide substrates, albeit at lower rates compared with Thr1. The active-site residue Thr1 is fixed in its position, as its methyl group is engaged in hydrophobic interactions with Thr3 and Ala46 (Fig. 4h). Consequently, the hydroxyl group of Thr1 requires no reorientation before substrate cleavage and is thus more catalytically efficient than Ser1. In agreement, at an elevated growing temperature of 37 °C the T1S mutant is unable to grow (Fig. 4a). In vitro, the mutant proteasome is less susceptible to proteasome inhibition by bortezomib (3.7-fold) and carfilzomib (1.8-fold; Fig. 5). Nevertheless, inhibitor complex structures indicate identical binding modes compared with the WT yCP structures, with the same inhibitors2223. Notably, the affinity of the tetrapeptide carfilzomib is less impaired, as it is better stabilized in the substrate-binding channel than the dipeptide bortezomib, which lacks a defined P3 site and has only a few interactions with the surrounding protein. Hence, the mean residence time of carfilzomib at the active site is prolonged and the probability to covalently react with Ser1 is increased. Considered together, these results provide a plausible explanation for the invariance of threonine as the active-site nucleophile in proteasomes in all three domains of life, as well as in proteasome-like proteases such as HslV (ref. 24). The 20S proteasome CP is the major non-lysosomal protease in eukaryotic cells, and its assembly is highly organized. The β-subunit propeptides, particularly that of β5, are key factors that help drive proper assembly of the CP complex1. In addition, they prevent irreversible inactivation of the Thr1 N terminus by N-acetylation41516. By contrast, the prosegments of β subunits are dispensable for archaeal proteasome assembly, at least when heterologously expressed in Escherichia coli25. In eukaryotes, deletion of or failure to cleave the β1 and β2 propeptides is well tolerated513141516. However, removal of the β5 prosegment or any interference with its cleavage causes severe phenotypic defects113. These observations highlight the unique function and importance of the β5 propeptide as well as the β5 active site for maturation and function of the eukaryotic CP. Here we have described the atomic structures of various β5-T1A mutants, which allowed for the first time visualization of the residual β5 propeptide. Depending on the (-2) residue we observed various propeptide conformations, but Gly(-1) is in all structures perfectly located for the nucleophilic attack by Thr1O, although it does not adopt the tight turn observed for the prosegment of subunit β1. From these data we conclude that only the positioning of Gly(-1) and Thr1 as well as the integrity of the proteasomal active site are required for autolysis. In this regard, inappropriate N-acetylation of the Thr1 N terminus cannot be removed by Thr1O due to the rotational freedom and flexibility of the acetyl group. The propeptide needs some anchoring in the substrate-binding channel to properly position Gly(-1), but this seems to be independent of the orientation of residue (-2). Autolytic activation of the CP constitutes one of the final steps of proteasome biogenesis26, but the trigger for propeptide cleavage had remained enigmatic. On the basis of the numerous CP:ligand complexes solved during the past 18 years and in the current study, we provide a revised interpretation of proteasome active-site architecture. We propose a catalytic triad for the active site of the CP consisting of residues Thr1, Lys33 and Asp/Glu17, which are conserved among all proteolytically active eukaryotic, bacterial and archaeal proteasome subunits. Lys33NH2 is expected to act as the proton acceptor during autocatalytic removal of the propeptides19, as well as during substrate proteolysis, while Asp17O orients Lys33NH2 and makes it more prone to protonation by raising its pKa (hydrogen bond distance: Lys33NH3–Asp17O: 2.9 Å). Analogously to the proteasome, a Thr–Lys–Asp triad is also found in L-asparaginase27. Thus, specific protein surroundings can significantly alter the chemical properties of amino acids such as Lys to function as an acid–base catalyst28. In this new view of the proteasomal active site, the positively charged Thr1NH3-terminus hydrogen bonds to the amide nitrogen of incoming peptide substrates and stabilizes as well as activates them for the endoproteolytic cleavage by Thr1O (Fig. 3d). Consistent with this model, the positively charged Thr1 N terminus is engaged in hydrogen bonds with inhibitory compounds like fellutamide B (ref. 29), α-ketoamides30, homobelactosin C (ref. 31) and salinosporamide A (ref. 32). Furthermore, opening of the β-lactone compound omuralide2 by Thr1 creates a C3-hydroxyl group, whose proton originates from Thr1NH3. The resulting uncharged Thr1NH2 is hydrogen-bridged to the C3-OH group. In agreement, acetylation of the Thr1 N terminus irreversibly blocks hydrolytic activity1516, and binding of substrates is prevented for steric reasons. By acting as a proton donor during catalysis, the Thr1 N terminus may also favour cleavage of substrate peptide bonds (Fig. 3d). In all proteases, collapse of the tetrahedral transition state results in selective breakage of the substrate amide bond, while the covalent interaction between the substrate and the enzyme persists. Cleavage of the scissile peptide bond requires protonation of the emerging free amine, and in the proteasome, the Thr1 amine group is likely to assume this function. Analogously, Thr1NH3 might promote the bivalent reaction mode of epoxyketone inhibitors by protonating the epoxide moiety to create a positively charged trivalent oxygen atom that is subsequently nucleophilically attacked by Thr1NH2. During autolysis the Thr1 N terminus is engaged in a hydroxyoxazolidine ring intermediate (Fig. 3d), which is unstable and short-lived. Breakdown of this tetrahedral transition state releases the Thr1 N terminus that is protonated by aspartic acid 166 via Ser129OH to yield Thr1NH3. The residues Ser129 and Asp166 are expected to increase the pKa value of Thr1N, thereby favouring its charged state. Consistent with playing an essential role in proton shuttling, the mutation D166A prevents autolysis of the archaeal CP21 and the exchange D166N impairs catalytic activity of the yeast CP about 60%. The mutation D166N lowers the pKa of Thr1N, which is thus more likely to exist in the uncharged deprotonated state (Thr1NH2). This renders the N terminus less suitable to stabilize substrates and to protonate the first cleavage product during catalysis, although it favours its ability to act as a nucleophile. This interpretation agrees with the strongly reduced catalytic activity of the β5-D166N mutant on the one hand, and the ability to react readily with carfilzomib on the other. Hence, the proteasome can be viewed as having a second triad that is essential for efficient proteolysis. While Lys33NH2 and Asp17O are required to deprotonate the Thr1 hydroxyl side chain, Ser129OH and Asp166OH serve to protonate the N-terminal amine group of Thr1. In accord with the proposed Thr1–Lys33–Asp17 catalytic triad, crystallographic data on the proteolytically inactive β5-T1C mutant demonstrate that the interaction of Lys33NH2 and Cys1 is broken. Consequently, efficient substrate turnover or covalent modification by ligands is prevented. However, owing to Cys being a strong nucleophile, the propeptide can still be cleaved off over time. While only one single turnover is necessary for autolysis, continuous enzymatic activity is required for significant and detectable substrate hydrolysis. Notably, in the Ntn hydrolase penicillin acylase, substitution of the catalytic N-terminal Ser residue by Cys also inactivates the enzyme but still enables precursor processing33. To investigate why the CP specifically employs threonine as its active-site residue, we used a β5-T1S mutant of the yCP and characterized it biochemically and structurally. Activity assays with the β5-T1S mutant revealed reduced turnover of Suc-LLVY-AMC. We also observed slightly lower affinity of the β5-T1S mutant yCP for the Food and Drug Administration-approved proteasome inhibitors bortezomib and carfilzomib. Structural analyses support these findings with the T1S mutant and provide an explanation for the strict use of Thr residues in proteasomes. Thr1 is well anchored in the active site by hydrophobic interactions of its C methyl group with Ala46 (C), Lys33 (carbon side chain) and Thr3 (C). Notably, proteolytically active proteasome subunits from archaea, yeast and mammals, including constitutive, immuno- and thymoproteasome subunits, either encode Thr or Ile at position 3, indicating the importance of the C for fixing the position of the nucleophilic Thr1. In contrast to Thr1, the hydroxyl group of Ser1 occupies the position of the Thr1 methyl side chain in the WT enzyme, which requires its reorientation relative to the substrate to allow cleavage (Fig. 4g,h). Notably, in the threonine aspartase Taspase1, mutation of the active-site Thr234 to Ser also places the side chain in the position of the methyl group of Thr234 in the WT, thereby reducing catalytic activity34. Similarly, although the serine mutant is active, threonine is more efficient in the context of the proteasome active site. The greater suitability of threonine for the proteasome active site, which has been noted in biochemical as well as in kinetic studies35, constitutes a likely reason for the conservation of the Thr1 residue in all proteasomes from bacteria to eukaryotes. Site-directed mutagenesis was performed by standard techniques using oligonucleotides listed in Supplementary Table 2. The pre2/doa3 (β5) mutant alleles in the centromeric, TRP1- or LEU2-marked shuttle vectors YCplac22 and pRS315, respectively, were verified by sequencing and subsequently introduced into the yeast strains MHY784 (ref. 1) or YWH20 (ref. 13), which express WT PRE2 from a URA3-marked plasmid. Counter-selection against the URA3 marker with 5-fluoroorotic acid yielded strains expressing only the mutant forms of β5. The strain producing a processed β5-T1A variant and the β5 propeptide in trans is a derivative of YWH212 (ref. 15). It carries an additional deletion of the NAT1 gene to avoid N-acetylation of Ala1; this strain exhibits extremely slow growth rates and served for crystallographic analysis only. All strains used in this study are listed in Supplementary Table 3. Yeast strains were grown in 18-l cultures at 30 °C in YPD into early stationary phase, and the yCPs were purified according to published procedures36. In brief, 120 g yeast cells were solubilized in 150 ml of 50 mM KH2PO4/K2HPO4 buffer (pH 7.5) and disrupted with a French press. Cell debris were removed by centrifugation for 30 min at 21,000 r.p.m. (4 °C). The resulting supernatant was filtered and ammonium sulfate (saturated solution) was added to a final concentration of 30% (v/v). This solution was loaded on a Phenyl Sepharose 6 Fast Flow column (GE Healthcare) pre-equilibrated with 1 M ammonium sulfate in 20 mM KH2PO4/K2HPO4 (pH 7.5). CPs were eluted by applying a linear gradient from 1 to 0 M ammonium sulfate. Proteasome-containing fractions were pooled and loaded onto a hydroxyapatite column (Bio-Rad) equilibrated with 20 mM KH2PO4/K2HPO4 (pH 7.5). Elution of the CPs was achieved by increasing the phosphate buffer concentration from 20 to 500 mM. Anion-exchange chromatogaphy (Resource Q column (GE Healthcare), elution gradient from 0 to 500 mM sodium chloride in 20 mM Tris-HCl (pH 7.5)) and subsequent size-exclusion chromatography (Superose 6 10/300 GL (GE Healthcare), 20 mM Tris-HCl (pH 7.5) and 150 mM NaCl) resulted in pure CPs for crystallization and activity assays. ChT-L (β5) activity of CPs was monitored by fluorescence spectroscopy using the model substrate Suc-LLVY-AMC. Purified yCPs (66 nM in 100 mM Tris-HCl, pH 7.5) were incubated with 300 μM substrate for 1 h at room temperature or 37 °C. The reactions were stopped by diluting samples 1:10 in 20 mM Tris-HCl, pH 7.5. AMC fluorophores released by proteasomal activity were measured in triplicate with a Varian Cary Eclipse Fluorescence Spectrophotometer (Agilent Technologies) at λexc=360 nm and λem=460 nm. Purified yCPs were mixed with dimethylsulfoxide as a control or serial dilutions of inhibitor and incubated for 45 min at room temperature. A final concentration of yCP of 66 nM was used. After addition of the peptide substrate Suc-LLVY-AMC to a final concentration of 200 μM and incubation for 1 h at room temperature, the reaction was stopped by diluting the samples 1:10 in 20 mM Tris-HCl, pH 7.5. AMC fluorophores released by residual proteasomal activity were measured in triplicate at λexc=360 nm and λem=460 nm. Relative fluorescence units were normalized to the dimethylsulfoxide-treated control. The calculated residual activities were plotted against the logarithm of the applied inhibitor concentration and fitted with GraphPad Prism 5. The IC50 value, the ligand concentration that leads to 50% inhibition of the enzymatic activity, was deduced from the fitted data. Mutant yCPs were crystallized as previously described for WT 20S proteasomes3637. Crystals were grown at 20 °C using the hanging drop vapour diffusion method. Drops contained a 1:1 mixture of protein (40 mg ml) and reservoir solution (25 mM magnesium acetate, 100 mM 2-(N-morpholino)ethanesulfonic acid (MES; pH 6.8) and 9–13% (v/v) 2-methyl-2,4-pentanediol (MPD)). Crystals were cryoprotected by addition of 5 μl cryobuffer (20 mM magnesium acetate, 100 mM MES, pH 6.8, and 30% (v/v) MPD). Inhibitor complex structures were obtained by incubating crystals in 5 μl cryobuffer supplemented with bortezomib or carfilzomib at a final concentration of 1.5 mM for at least 8 h. Diffraction data were collected at the beamline X06SA at the Paul Scherrer Institute, SLS, Villigen, Switzerland (λ=1.0 Å). Evaluation of reflection intensities and data reduction were performed with the programme package XDS38. Molecular replacement was carried out with the coordinates of the yeast 20S proteasome (PDB entry code: 5CZ4) by rigid body refinements (REFMAC5; ref. 39). MAIN40 and COOT41 were used to build models. TLS (Translation/Libration/Screw) refinements finally yielded excellent Rwork and Rfree, as well as root mean squared deviation bond and angle values. The coordinates, proven to have good stereochemistry from the Ramachandran plots, were deposited in the RCSB Protein Data Bank (Supplementary Table 1). The coordinates for the yeast 20S proteasome deposited under the entry code 1RYP do not represent the WT yCP but the double-mutant β5-K33R β1-T1A. At the time of deposition (in 1997), these data were the best available on the yCP. As yCP structure determination has become routine today, and structure refinement procedures have significantly improved, we here provide coordinates for the WT yCP at 2.3 Å resolution (PDB entry code: 5CZ4). Furthermore, the structures of most mutant yCPs described in this work were determined in their apo and ligand-bound states. For mutants with proteolytically inactive β5 subunits, the best crystallographic data obtained are given. For ligands or propeptide segments that were only partially defined in the 2FO–FC electron-density map the occupancy was reduced (for details see Supplementary Table 1). Accession codes: Coordinates and structure factors have been deposited in the Protein Data Bank, www.pdb.org (for PDB entry codes see Supplementary Table 1). How to cite this article: Huber, E. M. et al. A unified mechanism for proteolysis and autocatalytic activation in the 20S proteasome. Nat. Commun. 7:10900 doi: 10.1038/ncomms10900 (2016).
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