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. Author manuscript; available in PMC: 2024 Jun 4.
Published in final edited form as: J Struct Biol. 2024 Apr 13;216(2):108093. doi: 10.1016/j.jsb.2024.108093

Cryo-EM structure of bacterial nitrilase reveals insight into oligomerization, substrate recognition, and catalysis

Sergio Aguirre-Sampieri a,1, Ana Casañal b, Paul Emsley c, Georgina Garza-Ramos a,*
PMCID: PMC7616060  EMSID: EMS196545  PMID: 38615726

Abstract

Many enzymes can self-assemble into higher-order structures with helical symmetry. A particularly noteworthy example is that of nitrilases, enzymes in which oligomerization of dimers into spiral homo-oligomers is a requirement for their enzymatic function. Nitrilases are widespread in nature where they catalyze the hydrolysis of nitriles into the corresponding carboxylic acid and ammonia. Here, we present the Cryo-EM structure, at 3 Å resolution, of a C-terminal truncate nitrilase from Rhodococcus sp. V51B that assembles in helical filaments. The model comprises a complete turn of the helical arrangement with a substrate-intermediate bound to the catalytic cysteine. The structure was solved having added the substrate to the protein. The length and stability of filaments was made more substantial in the presence of the aromatic substrate, benzonitrile, but not for aliphatic nitriles or dinitriles. The overall structure maintains the topology of the nitrilase family, and the filament is formed by the association of dimers in a chain-like mechanism that stabilizes the spiral. The active site is completely buried inside each monomer, while the substrate binding pocket was observed within the oligomerization interfaces. The present structure is in a closed configuration, judging by the position of the lid, suggesting that the intermediate is one of the covalent adducts. The proximity of the active site to the dimerization and oligomerization interfaces, allows the dimer to sense structural changes once the benzonitrile was bound, and translated to the rest of the filament, stabilizing the helical structure.

Keywords: Nitrilase, Filament forming enzymes, Enzyme catalysis, Covalent adduct, Benzonitrile, Filamentation

Graphical abstract.

Graphical abstract

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1. Introduction

Nitrilase enzymes (EC 3.5.5.1) catalyze the direct conversion of organic nitriles to the corresponding carboxylic acids and ammonia. They are widely distributed in nature, where they can be found in bacteria, fungi, and plants (O’Reilly and Turner, 2003). The first nitrilase was discovered in the early 1960s (Thimann and Sundararaman, 1964), and since then some of these cell factories have been utilized for the commercial production of carboxylic acids at an industrial scale. Instead of the conventional chemical synthesis, biocatalytic routes are developed to meet green chemistry requirements on cost-efficiency, waste reduction, energy consumption, and other synthesizing objectives (Gong et al., 2012), providing synthesis possibilities that are difficult or impossible to achieve by conventional catalytic methods. Although the physiological role of nitrilases is not yet clear, its participation is recognized in the synthesis of 3-indoleacetic acid and cyanolipids, or in the hydrolysis of β-cyanoalanine, an intermediate in the cyanide detoxification pathway in plants (Piotrowski, 2008). Significant differences among nitrilases are related to substrate specificity, quaternary structure and aggregation properties. According to their specificity, nitrilases are classified into three categories: those that hydrolyze aromatic or heterocyclic nitriles; those that preferentially degrade aliphatic nitriles, and those that act on arylacetonitriles (O’Reilly and Turner, 2003). In addition, some bacterial and fungal nitrilases can assemble into filamentous structures, property that has been observed in vivo for the nitrilases of Arabidopsis thaliana, in which filamentous structures were found to aggregate in proximity to the leaf cell nucleus, as response to mechanical wounding or herbicide treatment and prior to cell death (Cutler and Somerville, 2005; Doskocilova,et al., 2013). It is proposed that these filamentous structures could provide a platform for molecular interactions in the regulation of cellular processes, or act as a scaffold for other nitrilase-associated proteins (Thuku et al., 2007).

Nitrilases belong to the first branch of the nitrilase superfamily, which comprises 13 classes of hydrolases acting on a wide variety of nonpeptide carbon–nitrogen bonds (Pace and Brenner, 2001). Despite large variations in the sequence of the different enzyme classes, they all share a characteristic homodimeric building block with an αββα – αββα super sandwich fold, and a conserved catalytic triad formed by Glu, Lys, and Cys (Brenner, 2002). Nitrilases are homo-oligomers with a monomer size of approximately 30–45 kDa, which can assemble as helical filaments of variable length. These filament structures have been reported for the cyanide dihydratase from Bacillus pumilus, (Jandhyala et al., 2003) and nitrilases from Rhodococcus rhodochrous J1 (RrJ1Nit) (Thuku et al., 2007), Fusarium solani O1 (Vejvoda et al., 2009), Nit1, Nit3, and Nit4 from Arabidopsis thaliana, and Nit1 and Nit2 from Capsella rubella, (Woodward et al., 2018). It is known that several enzymes form large polymers with subunits assembled into filaments (Aughey and Liu, 2015). These kinds of structures occur frequently in biological systems and are functionally important for providing stability and improving enzyme activity; however, for many of these structures, we still lack an understanding of the regulation and polymerization mechanism (Barry et al., 2014; Park and Horton, 2020; Yeates and Padilla, 2002). In this regard, it has been shown that oligomerization in helical filaments of nitrilases is an essential requirement for its catalytic activity, so it’s necessary to know how the interactions in the different interfaces stabilize the quaternary structure, as well as the correlation among them and the enzymatic activation process, a common phenomenon in this class of enzymes. A particularly noteworthy example is that of RrJ1Nit, Nagasawa et al. (2000) showed that the wild-type enzyme is an inactive dimer that forms an active 410 kDa oligomer after incubation with ammonium sulfate or benzonitrile substrate. Later, Thuku et al. (2007) observed a third active structural form of RrJ1Nit that assembles as regular helical filaments of variable sizes. These filaments were generated after the cleavage of 39 amino acids at the C-terminal end of the enzyme, removing the steric hindrance imposed by the C-terminus and allowing extended helices to form (Thuku et al., 2007). Furthermore, the interactions that stabilize the nitrilases in their various oligomeric assemblies, their contact interfaces, and the position of the C-terminal end has been identified by negative stain TEM (Thuku et al., 2009; Woodward et al., 2018) and more recently by Cryo-EM (Mulelu et al., 2019).

To date the atomic structures of three nitrilase family members belonging to branch 1 have been reported, these are the thermoactive nitrilase from Pyrococcus abyssi (PaNit) (PDB 3IVZ) (Raczynska et al., 2011), the nitrilase from Syechocystis sp. strain PCC6803 (Nit6803) (PDB 3WUY) (Zhang et al., 2014), and the Nit4 from Arabidopsis thaliana (AtNit4) (PDB 6I00) (Mulelu et al., 2019). PaNit is a homodimer with a C-terminal end ~100 amino acids shorter than most bacterial and plant nitrilases and a shared identity of only 22–26 % with these. In this regard, it was recently shown that the PaNit enzyme has mostly amidase activity with barely detectable activity towards nitrile substrates and shows a 78 % sequence identity with the well-known amidase from P. yayanosii CH1 (AmiE) (Fu et al., 2014; Makumire et al., 2022). Therefore, it may be necessary to reclassify PaNit and the hypothetical protein PH0642 (PDB 1J31) (Sakai et al., 2004) from P. horikoshii as amidases in the nitrilase superfamily. On the other hand, the asymmetric unit of Nit6803 is a dimer that oligomerizes to form a C-shaped extended helix in the crystal structure, whereas in solution the enzyme forms a decamer. A 57-amino acid fragment of its C-terminus was trimmed during the crystallization process which allowed the protein to crystallize (Zhang et al., 2014). The Cryo-EM structure of the helical filament of AtNit4 shows a left-handed helical assembly with 4.9 dimers per helical turn. For this active helical nitrilase, the predominant interaction that stabilizes the filament occurs through the C-terminal ends of four adjacent monomers, all of which contribute with strands that form a row of four ß-sheets in the core of the filament (Mulelu et al., 2019). These interactions stabilize the helical structure keeping the enzyme in an active oligomeric form.

However, to date not a single atomic structure of a nitrilase with a substrate or intermediate molecule bound to its active site is available, limiting our knowledge of the structural determinants for substrate recognition and the catalytic mechanism. The Glu-Glu-Lys-Cys catalytic tetrad is well conserved among the nitrilase family, in which the catalytic cysteine residue is localized at the bottom of the active site pocket, at the top of a β-turn-α structural motif, known in the superfamily as the nucleophile elbow (Kumaran et al., 2003), that in an energized conformation, acts as the nucleophile (Andrade et al., 2007). It has been shown that the reaction mechanism leads to the formation of covalent adduct intermediaries (Stevenson et al., 1990). It’s proposed that the reaction mechanism is acid-base catalyzed, a catalytic glutamate would act as the general base, whereas the catalytic lysine/glutamate as the general acid (Nakai et al., 2000; Fernandes et al., 2006; Raczynska et al., 2011).

In this work, using a nitrilase from Rhodococcus sp V51B as a model system, we explored the effect of the filament quaternary structure formation on the nitrilase activity. We generated a C-terminal truncated mutant NITΔC39, which assembles into large helical filaments with enhanced catalytic activity. After incubation of NITΔC39 with its substrate benzonitrile, the length of the filaments increases up to a size of 500 nm. This property was used to determine the structure of the nitrilase filaments by Cryo-EM at 3 Å resolution. The structure comprises a complete turn of the helical arrangement with a substrate-intermediate bound to the catalytic cysteine, giving a glimpse of what the reaction mechanism may look like, and its relationship with the oligomerization process. The structural comparison of NITΔC39 with the other two nitrilase structures reveals a unique configuration of its active site, which is completely buried inside each monomer. Access to the active site is restricted by segments of two other monomers, right between the dimerization and high-order oligomerization interfaces.

Nitrilase filaments constitute an ideal prototype for the development of supramolecular structures and their implementation in biotechnological processes. As more structural information becomes available, structure-based design for improvement of catalysis and stability, and/or changes in substrate specificity and enantioselectivity will become possible.

2. Materials and methods

2.1. Cloning

The gene coding the bacterial nitrilase (NitA) (S.M.) was amplified by PCR from the total DNA of Rhodococcus sp. V51B (NCBI taxonomy ID. 1831) as a template, with oligonucleotides designed to introduce the NdeI and HindIII restriction sites flanking the NitA gene (Table S1) and cloned into pET24a+ (Novagen) vector.

2.2. Construction of C-terminal truncated mutants

Based on the sequence of the NitA gene, C-terminal deleted mutants NitΔC39 and NitΔC49 were generated by PCR using the nitA5´primer and different 3´ oligonucleotides to introduce the respective stop codons along the C-terminus. After the restriction of amplified DNA fragments, the mutant genes were ligated into the pGEM-T Easy (PROMEGA) vector and cloned in E. coli NEB5-α cells. Positive clones were selected, sequenced, and inserted into NdeI/HindIII cleaved pET24a (+). A C165A mutant was obtained from the pET24a (+) construct of NitΔC39 by site-directed mutagenesis using the respective mutagenic primers (Table S1).

2.3. Overexpression and protein purification

The recombinant plasmids carrying the nitrilase genes of the wild-type and C-terminal mutants were transformed into E. coli BL21(DE3) pLys and grown on LB-agar-kanamycin medium plates at 37 °C, 50 mL of LB liquid medium supplemented with kanamycin was inoculated with a fresh colony and grown at 37 °C overnight. The culture was used to inoculate 1L of LB medium supplemented with kanamycin and allowed to grow at 37 °C until an OD600 of 0.6 was achieved. Expression of the proteins was promoted with the addition of 0.4 mM isopropyl-β-D-thiogalactopyranoside (IPTG) for 8 h at 27 °C. Cells were harvested by centrifugation at 5,000 rpm for 20 min, resuspended in 40 mL of lysis buffer (100 mM Tris pH 7.8, 1 mM EDTA, 150 mM KCl and 0.2 mM PMSF), and disrupted by sonication. Cells debris was removed by centrifugation at 15,000 rpm, for 20 min at 4 °C. Precipitation of the soluble cell extract was carried out by fractionation with ammonium sulfate at 4 °C, obtaining the mutant proteins in the fraction between 0 and 20 % saturation, and between 20 and 45 % saturation for the WT enzyme. After centrifugation at 15,000 rpm for 20 min, the resulting precipitate was solubilized in 10 mM K-phosphate buffer pH 7.0, 1 mM EDTA, 200 mM NaCl and 14.25 mM β-mercaptoethanol (buffer-A). The solution was loaded on a Sephacryl S-300 column previously equilibrated with buffer-A, and fractions were analyzed by SDS-PAGE.

2.4. Dynamic light scattering (DLS)

Purified protein samples were dialyzed in 100 mM K-phosphate buffer pH 7.8, 100 mM KCl, and filtered through a 0.22 μm pore size membrane. Then, 400 μl of each sample at a concentration of 0.2 mg/ml were transferred to a quartz cell thermostated at 25 °C. Size distribution data was obtained with the use of a Zetasizer μV instrument equipped with a photodiode laser (830 nm) (Malvern Instruments Ltd.). Zetasizer software 7.13 was used to analyze the data. DLS values were averages of 10 distinct scans for 10 acquisitions over 10 s on a given sample.

2.5. Enzyme assays

Standard nitrilase activity assays were carried out in 1 mL of 100 mM K-phosphate buffer pH 7.8 (activity buffer), containing 5 μg/mL enzyme and substrates at the concentrations indicated. After 10 min incubation at 30 °C, the reaction was stopped with the addition of 0.1 mL of 2 N HCl. The reaction mixture was centrifuged at 14,000 rpm for 10 min at 4 °C. The supernatant was taken, and the ammonia content was determined by using a commercially available ammonia test system based on the phenol-hypochlorite method (Spectroquant® Merck-Milipore).

For the determination of kinetic parameters, the concentration of benzonitrile, adiponitrile, and valeronitrile was varied over a range of 0.01 to 1 mM, reaction aliquots were taken at intervals between 1 to 15 min. The ammonium concentration formed in the reaction mixture was determined by generating a curve with standard ammonium solution from 0.02 to 1.0 mM concentration.

2.6. Negative-stain electron microscopy

Purified protein at a final concentration of 0.5 mg/ml was incubated for 15 min in a carbon-coated copper grid before being dried by contact with filter paper. The sample was stained by adding a drop of 2 % phosphotungstic acid and air-dried for 2 min. The grids were observed in a JEOL JEM 12000 EII equipped with an 11-megapixel CCD Camera (GATAN, Orius SC1000), and imaged at a sampling of 4.32 Å pixel − 1 at 80 kV. The helical arrangement of the observed particles was confirmed with the use of Fiji (Image J) by an inverse fast Fourier transform of a single filament particle.

2.7. Cryo-electron microscopy

Purified protein at 1 mg/ml was centrifuged at 13,000 rpm for 15 min, and 100 μl supernatant was incubated for 15 min with 10 μl of concentrated benzonitrile (9.8 M) (Thermo Scientific). The holey grids (Quantifoil, Electron Microscopy Sciences) were glow-discharged for 60 sec before deposition of 3 μl sample (~3.8 mg ml − 1), blotted for 4 s, and vitrified by plunging into liquid ethane with the use of a Vitrobot IV (FEI) at 10 °C with 100 % humidity, and stored in a cryobox in liquid nitrogen. The grids were loaded on a Titan Krios microscope equipped with a Falcon 3 Direct Electron Detector (FEI) at 300 kV (MRC-LMB, Cambridge, UK). A total of 2,358 images consisting of 75 frames, with a 0.49 e/px/frame dose rate and with a 1.04 Å pixel size were collected in counting mode. (Fig. S1A).

2.8. Image processing

For image processing and map reconstruction, RELION3.0 (He and Scheres, 2017) was used along with MotionCor2 (Zheng et al., 2017), CTFFIND4 (Rohou and Grigorieff, 2015), and Gctf (Zhang, 2016). A total of 140,381 particles were used for 2D classification, whereby a featureless cylinder was generated. A second 2D classification was performed to choose the best 45 2D classes comprising 56,369 particles and used to generate a 3D classification that showed 6 3D classes, from these, two classes with the best particles were selected and used to calculate rise (18.4 Å) and twist (−288°). After various cycles of refinement, 3D classification and postprocessing, a 3 Å resolution map was generated (Fig. S1B–E).

2.9. Model building, refinement, and validation

For model building, the main chain of the pyrimidine degrading enzyme from Drosophila melanogaster (PDB 2VHH) was used as a template, mutations, errors in connectivity, unbuilt residues, and incorrect rotamers were corrected against the EM map in Coot (Emsley et al., 2010), and refined in a first-round using Refmac5 (Murshudov et al., 2011). Clashes were later corrected with the use of Isolde (Croll, 2018) in Chimera X (Goddard et al., 2018) and refined again with Phenix (Liebschner et al., 2019). A second model with benzaldehyde adducts bound to the catalytic cysteines was constructed through AceDRG (Long et al., 2017) in CCP4i2 (Potterton et al., 2018), adjusted with Coot, and validated (Fig. S1F). Coot was also used for modeling and geometry optimization of the different nitrile intermediates. Map and coordinates were deposited on the EMDB and wwPDB under the accession codes EMD-42779 and 8UXU respectively.

2.10. Cavity search and docking

CASTp3.0 (Computed Atlas of Surface Topography of proteins) webserver (Tian et al., 2018) was used for locating, delineating, and measuring geometric and topological properties of the active site cavity, with a default probe radius of 1.4 Å. Molecular Docking was performed with AutoDock Vina (Eberhardt et al., 2021), using as a search template the active site area previously identified by CASTp3.0.

2.11. Data visualization

Molecular visualization, structural analysis, and image preparation were performed using Fiji image J (Schindelin et al., 2012), UCSF Chimera (Pettersen et al., 2004), Chimera X, ChemDraw, PyMol v1.8.2.1 (Schrödinger and DeLano, 2020) and Coot.

3. Results

3.1. Specificity and kinetic properties

Starting from the recombinant Rhodococcus sp V51B WT nitrilase (RV51BNit), a C-terminal truncated mutant, in which the last 39 amino acids residues at the carboxyl terminus were removed, was generated (NitΔC39). This mutant was designed considering the proteolysis site reported for R. rhodochrous J1 nitrilase, since they share 97 % identity. Similar to what was reported by Thuku et al. (2007), the NitΔC39 mutant assembles into regular, active, and stable helical filaments of variable length between 50–200 nm (Fig. 1A), with an enhanced catalytic activity 5 times higher than the WT enzyme (Fig. S2).

Fig. 1. Filament formation is prompted after the incubation with benzonitrile.

Fig. 1

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A) Negative stain TEM images of NitΔC39 before and after incubation with valeronitrile, adiponitrile, and benzonitrile respectively. B) Representative Cryo-EM images before and after incubation with benzonitrile. C) 2D class averages of NitΔC39 with benzonitrile from Cryo-EM images. D) Observed quality of model fitting into map.

The NitΔC39 mutant, as the WT nitrilase, has preference for aromatic nitriles, showing its highest activity towards benzonitrile, 4-Cl benzonitrile, and cyanopyridine substrates. The enzyme exhibited low preference through the 2-substituted derivates of these substrates, such as 2-Cl benzonitrile, 4-Cl 2-cyanopyridine, or 2-cyanopyridine. For the dinitriles and aliphatic nitriles tested, the activity was 5 to 25 % with respect to benzonitrile (Fig. 2). We determined the kinetic constants of NitΔC39 for some selected substrates (Table S2). The kcat and the kcat/KM values for benzonitrile were higher than those for adiponitrile and valeronitrile, with a specificity constant of 200-fold and 40-fold higher, respectively.

Fig. 2. Substrate selectivity of NitΔC39.

Fig. 2

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Mixture reactions contained 100 mM K-phosphate buffer pH 7.8, 5 μg/mL enzyme and 10 mM substrate were incubated at 30 °C for 5 min. The reaction was arrested by mixing 1 mL of the reaction mixture with 0.1 mL of 2 N HCl and the amount of ammonia released was spectrophotometrically determined. The specific activity for benzonitrile (16.7 μmol min−1 mg−1) was taken as 100 %.

3.2. Substrate-induced oligomerization

After screening different buffer conditions, protein concentration, and ligands, we noticed that certain substrates could induce heavy aggregation of NitΔC39. It is known that Rhodococcus rhodochrous J1 nitrilase assembles into higher oligomers and acquires activity in response to benzonitrile (Nagasawa et al., 2000). We observed the effect of 3 different nitriles (one aromatic, benzonitrile; one dinitrile, adiponitrile; and one aliphatic, valeronitrile) on the oligomeric state of the protein, through DLS, negative-stain electron microscopy, and by Cryo-EM (Figs. 1A and S3). We noticed that while benzonitrile dramatically induces enzyme oligomerization from filaments around 100–200 nm to 500 nm lengths, adiponitrile seems to decrease filament formation, while valeronitrile doesn’t seem to have a significant effect on the oligomeric state of the protein (Figs. 1A and S3). Interestingly an inactive mutant NitΔC39 – C165A also assembles as filaments regardless of the presence of benzonitrile, while RV51BNit wild type mostly forms irregular aggregates. Indeed, it is possible to induce filament formation in an even shorter mutant, NitΔC49 after incubation with benzonitrile (Fig. S2).

3.3. The overall structure of NitΔC39

To further investigate the effect of the substrate on the oligomerization and activation of the enzyme, we reconstructed NitΔC39 at 3 Å resolution by Cryo-EM (Fig. S1). In general, filament quality (length and number of usable particles) was improved after incubation with benzonitrile. A clearly helical arrangement was visible in 2D classes. The structure comprises two turns of the helical arrangement, and although the quality of the map decreases from the center, 7 dimers could be modeled with certainty into density (Fig. 3A). The filament is formed by the association of dimers, which are the minimum building blocks, as has been observed in other nitrilase homologs. The helical structure is a left-handed spiral, in which each turn is composed of 5 dimers, with a helical twist of −72° per dimer, an outer diameter of ~13.5 nm, and a ~4.2 nm “five-point star” shape hollow core (Fig. 3B).

Fig. 3. Electronic potential map of NitΔC39 quaternary structure.

Fig. 3

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A) External side view. Seven dimeric units are presented, dimers are numbered [1–7] from top to bottom. Monomers in each dimer are presented in different colors. The filament is comprised of a left-handed helix with a pitch of 9.25 nm, a helical rise of 18.4 Å and a helical twist of −288°. B) Top view. The cylinder is composed of 5 dimers per turn, with a helical twist of −72°, an outer diameter of ~13.5 nm, and a “star shape” inner tunnel with a diameter of ~4.2 nm. C-terminal ends contribute to channel formation between dimers in a longitudinal plane, that can be seen next to the edges of the star. C) Side view of the filament core. Three dimers [3 to 5] are presented: one in state blue and cyan [5], another in gray and orange [4], and the last in forest and pale green [3]. Dashed lines indicate external diameter length. D) In the interior of the helix, the C-terminal ends of the previous [5] and contiguous [3] monomers of adjacent dimers interact with the C-terminal ends in the middle dimer [4] through the oligomerization interfaces. The map is presented with 70 % transparency over a black background with a stick representation of the mentioned C-terminal tails.

The model comprises 318 residues (2–319) from a total of 328. The map lacks clear density for the last nine C-terminal amino acid residues, which were not modeled. Details on the Cryo-EM statistics are summarized in Table 1. The C-terminal end of four adjacent monomers interact in the inner core of the spiral, at the center of each dimer, in an attaching mechanism that stabilizes the spiral (Fig. 3C and D), as previously reported by Mulelu et al. (2019). In the reconstructed map it is possible to observe a continuous density bound to the catalytic cysteine (C165), indicating the presence of a covalently bound molecule, due to the addition and saturation with benzonitrile substrate, possibly an intermediary of the reaction mechanism.

Table 1. Cryo-EM data collection, refinement, and validation statistics PDB 8UXU.

Data collection
Microscope Titan Krios
Voltage (kV) 300
Detector Falcon III
Magnification 75 K
Pixel Size (Å) 1.04
Defocus range (nm) 500–5000
Exposure time (s) 2.04
Dose rate (e- pixel−1 s−1) 0.45 e/px/frame
Number of images 2358
Number of frames per image 75
Map reconstruction
Initial particle number 140,381
Final particle number 56,369
Map size (Å) 360, 360, 360
Helical twist (° ) –288°
Helical rise (Å) 18.4 Å
Resolution determination method FSC 0.143 cut-off
Resolution (Å) 3.019
Model composition
Non-hydrogen atoms 33,559
Protein residues (all/built) 328/318
Ligands 14
R.m.s. deviations
Bond lengths (Å) 0.32
Bond angles (° ) 0.5
MolProbity score 1.53
Validation
Clashscore 6
CaBLAM outliers (%) 1.16
Ramachandran plot
Favored (%) 97
Allowed (%) 3
Outliers 0.00
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3.4. Oligomerization mechanism

In the assembly mechanism of NitΔC39, one of the monomers is rotated 180° relative to the other along an axis perpendicular to the Z axis and will interact to form a dimeric unit through dimerization interface ‘A’, as previously described for other nitrilases (Lundgren et al., 2008; Sewell et al., 2003; Raczynska et al., 2011). When two monomers associate, a “super-sandwich” (αßßα-αßßα) structure is formed. The main contributions to interface ‘A’ may be seen at the C-terminal end of each monomer, which associate themselves through two antiparallel β-strands (Fig. 4A). Dimeric symmetrical interactions between S305-N307, N309-V303, K174-E217, H297-K133 and R139 can be observed in the C-terminal segment of two monomers in the same dimer, but also another interaction among Y298-T135 or Y298-W166 in distinct monomers within the same dimer is observed (Fig. S4). These interactions across residues near the C-terminal end of the protein, which plays an essential role in protein dimerization and higher oligomerization, and residues surrounding the active site pocket, explains at least partially, why oligomerization has an essential effect on the enzymatic activity of the enzyme. Curiously, amino acids forming part of the ‘A’ surface, are positioned from and downriver the active site, from position 132 onwards. The C-terminal end of contiguous monomers in adjacent dimers extends to interact with the dimerization interface ‘A’, in which the C-terminal ends of each monomer interact to form a row of 4-stranded β-sheets in the inner core of the filament, as seen in AtNit4 (Mulelu et al., 2019). The ‘C’ interface arises from C-terminal ends interactions across adjacent monomers in distinct dimers, but also between contacts in adjacent dimers; a monomer from one dimer will interact with the one positioned next to it in the contiguous dimer through an extensive surface, generating symmetrical interactions between residues K133-E144, N307-N317 and R139-R139. These interactions will repeat on each side of the dimers in a chain-like mechanism that attaches and stabilize the spiral. Interactions involving the C-terminal end are shared across ‘A’ and ‘C’ interfaces (Fig. 4B). Amino acid residues contributing to the formation of ‘A’ and ‘C’ interfaces are outlined in Fig. S5. Also ‘A’ and ‘C’ interfaces contribute to the formation of the active site entrance, which is positioned facing the exterior of the filament, within the dimerization and higher-oligomerization interfaces, in the surface where 3 monomers interact, two in the same dimeric subunit and another from the contiguous dimer (Fig. 4C).

Fig. 4. Nitrilase helical arrangement.

Fig. 4

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Left to right: A) Dimer representation. Monomers (gray and orange) interact through C-terminal ends giving rise to the ‘A’ interface (green) to form a dimer with a super sandwich αββα–αββα characteristic fold. B) Tetramer representation. Dimers interact across the ‘C’ interface (magenta), giving rise to a supra-molecular assembly. Residues contributing to the active site entrance in the contiguous dimer are shown in blue. Dashed lines highlight association interfaces. C) External side and top views of an octamer surface representation. Both ‘A’ and ‘C’ interfaces contribute to the formation of the active site entrance (cyan arrows), which lies between the interface formed by 3 monomers. Bottom: Schematic representation of the different oligomeric states. The square size for the octamer top view is presented from furthest (smaller) to closest (bigger) dimer according to the “point of view”.

3.5. Substrate-binding pocket

According to CASTp3.0, the NitΔC39 pocket volume molecular surface is 218.4 Å and has a molecular surface area of 245.4 Å2. A loop arising from α9 in the adjacent monomer in the same dimer extends over the entrance of the active site pocket like a lid (A291), but also a methionine residue (M67) from the contiguous dimer contributes to the formation of this lid area (Fig. 5A). The cavity is formed by a few aromatic and hydrophobic residues, such as Y54, Y56 in a loop between ß2-α2, W59 in ß2, T135 and E138 in ß7-η1, C165 and W166 in ß9-η2, P190, G191, M192, S193, and L194 between ß10- η3 and F202 in η3. This type of aromatic-rich pocket has also been described for other nitrilases (Liu et al., 2013; Zhang et al., 2014). The catalytic tetrad, E48, K131, E138, and C165 is positioned at the bottom of the active site, away from the lid. The pocket is accessible from the surrounding media only through a funnel-shaped entrance formed by T135, H136, W166, and F202. Curiously, a single rotameric change in H136 is enough to close this funnel entrance. The catalytic cysteine C165 is positioned in the catalytic elbow, which for NitΔC39 is η2 (Fig. 5B). This residue is in an energized conformation, as judged by its localization in a disallowed region of the Ramachandran plot.

Fig. 5. The bacterial nitrilase active site.

Fig. 5

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A) Location of the active site in the NitΔC39 dimer. Dimers are numbered according to Fig. 2. Monomers in the same dimer [4] are shown in gray and orange. Segments from monomers in adjacent dimers that contribute to the active site entrance are shown in state blue [5] and forest green [3]. The catalytic residues and residues contributing to active site entrance in different monomers are shown as sticks. The position of the active site in each monomer is indicated by a dashed square. B) A semi-transparent solvent-accessible surface representation of the catalytic cavity (yellow), residues contributing to the binding pocket are shown as gray sticks. The entrance to the active site pocket is a funnel-shaped tunnel located in the contact interface between the two monomers in the same dimer [4] (gray and orange) and a monomer from the contiguous dimer [5] (blue). The pocket entrance residues in each monomer are shown as sticks in different colors, T135, H136, W166, and F202 in gray, A291 in orange, and M67 in blue. C) Presence of a reaction intermediary within the active site. The electronic potential map corresponding to 165 S-benzoyl cysteine (C165**) is shown as a mesh, the map is contoured at 0.035 level. The benzaldehyde adduct is shown in magenta. The catalytic residues E48, K131, E138, and C165 are labeled with *. D) Mode of substrate binding. A benzonitrile molecule modeled in the same position and orientation as the adduct is shown in magenta. Distances between catalytic residues and benzonitrile are shown in blue, the distance between E138 and K131 is shown in yellow, and residues with a VDW overlap ≥ 0.5 Å are shown. A dashed circle indicates the position of the cyanocarbon atom in the nitrile.

Inside the catalytic pocket, the electronic potential map showed a continuous density that extends over the length of the catalytic cysteine C165 (Figs. 5C and S1). We modeled into this density the different reaction intermediaries in the hydrolysis of benzonitrile to benzoic acid (Fig. S7). Best fitting was obtained for an intermediary with just one double-bond ramification of the cyanocarbon atom of the nitrile. Taking into the assumption that the first steps of the reaction occur rapidly, as suggested by Stevenson et al. (1990), we first modeled a benzaldehyde molecule within the density, and later a covalent benzaldehyde adduct (Figs. 5C and S6).

3.6. Comparison with other nitrilases

According to CATH classification (Orengo et al., 1997), the NitΔC39 monomer has a 4-layer αßßα sandwich architecture and is classified within the homologous superfamily of carbon–nitrogen hydrolases, also known just as nitrilases. Nitrilase enzymes identified with DALI-server (Holm et al., 2023) are Syechocystis sp. PCC6803 nitrilase (Nit6803) (PDB 3WUY, Z-score 41.4, r.m.s.d. 1.5 Å, %id 39), and Arabidopsis Nit4 (AtNit4) (PDB 6I00, Z-score 33.8, r.m.s.d. 1.6 Å, %id 37). Among them, NitΔC39 is closest related in substrate affinity to Nit6803, although the oligomeric structure and C-terminal interactions resemble the ones from AtNit4. The main differences between both filament structures are exhibited in NitΔC39 α2- α3 (I58-F70) with a small helix and a loop, and in AtNit4 α3 (G85-D101) with a longer loop, NitΔC39 has a loop insertion (η3) from G191 to G203. Also, the AtNit4 model lacks atoms from R251-V276, which in NitΔC39 (T230-R249) form two small α helixes (α7 & α8) connected by a loop (Figs. 6A and S5). Strikingly, major differences can be seen in the active site configuration of the three enzymes (Fig. 6B). The NitΔC39 active site cavity is the only one among the three that is buried inside each monomer, its active site lid is contained within the same dimeric unit, which appears to be restricting access from the surrounding media. Due to the presence of a covalent adduct most probably the lid is observed in a closed configuration. The AtNit4 active site is partially exposed, its lid is projected from a monomer in the contiguous dimer, which completes the active site in a closed configuration. In Nit6803, the active site cavity is completely exposed, and like NitΔC39, the lid comes from the monomer in the same dimeric unit. The deletion of 57 residues at the C-terminal end of Nit6803 brought an inactive enzyme with an irregular helical twist, which is thought to lead the active site to an open configuration (Mulelu et al., 2019; Zhang et al., 2014).

Fig. 6. Comparison of nitrilases with known structures.

Fig. 6

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A) Structural comparison between NitΔC39, AtNit4 (PDB 6I00) and Nit6803 (PDB 3WUY). An overall monomeric fold of the four enzymes, the backbone is colored from blue (0.5 Å) to red (4 Å) depending on the backbone root-mean-square deviation (r.m.s.d.) between the three enzymes, above a 4 Å deviation, NitΔC39 is presented in green, Nit6803 in pale cyan, and AtNit4 in grey. Unbuild main chain atom (unb.) in AtNit4 are connected by a yellow cylinder. The active site location is indicated by a yellow star in each monomer. B) Active site pocket of each enzyme, the different cavities are presented in a similar orientation according to the catalytic tetrad (*). Residues contributing to the active site cavity in the same monomer are presented in gray, and residues contributing to the lid are presented in orange. The surfaces are colored by hydrophobicity, from blue → hydrophilic, to yellow → hydrophobic. Residues in each enzyme that contribute to the active site formation are labeled by one letter code and number. Lid: C = Closed conformation. O = Open conformation.

3.7. Mode of substrate binding

As suggested by Liu et al. (2013) the closing of the lid precedes the formation of the thioester intermediate. Indeed, Andrade et al. (2007) proposed that for the substrate to enter and leave the active site cavity, residues around the entrance of the cavity and the lid must move to allow the entrance of the substrate and the latter release of products. The position of the lid, restricting access from the media to the funnel-shape entrance, and the size and volume of this tunnel, which is not big enough for the aromatic ring of the benzonitrile to diffuse through, suggests some mobility in the residues around the active site entrance should occur, hence the present structure is in a “closed” state and supports the hypothesis that the bound molecule in the active site is a covalent reaction intermediary. The intermediary is held in place by aromatic and hydrophobic residues, Y54, W59, W166, L192, M194, and F202, all of which seem to orient and hold the molecule. The aromatic ring of the intermediary is sandwiched between Y54 and W166, and between W59 and M192, while W166 and F202 interact with this ring through anion-π interactions (Fig. 5C), in turn W166 also interacts with F169 positioned at the ‘A’ interface through a π-π stacking interaction. Molecular docking analysis of a NitΔC39 monomer with benzonitrile diffused in its active site, localizes the benzonitrile molecule within the binding pocket and positions the cyano group of benzonitrile facing the sulfur atom of the catalytic C165, which is necessary for the nucleophilic attack to occur, and imposes constraints on the possible position and orientation of the bound substrate (Data not shown).

3.8. Implications for the reaction mechanism

Modeling of a substrate molecule in the same position and orientation as the adduct (Figs. 5D and S6), places the nitrogen atom in the nitrile at a ~3.4 Å distance from the catalytic residues E48 and E138, and at ~3.6 Å from K131, and positions the cyanocarbon atom in the nitrile, parallel to the catalytic cysteine, at a ~1.6 Å distance from the sulfur atom. For the nucleophilic attack to start, the nitrile molecule needs to be placed next to the catalytic cysteine in a parallel position at the bottom of the active site (Fig. 5D). Given the distances between the substrate and the aromatic residue F202 at the upper part of the catalytic pocket, is feasible to imagine this residue pushing the substrate closest to the catalytic tetrad after the closing of the lid, pressing the cyano carbon atom in the nitrile towards the sulfur atom in the catalytic cysteine and initiating the enzymatic reaction.

The finding of a ligand in the NitΔC39 active site, solved as a benzaldehyde-adduct, which is a predicted reaction intermediary, gives us a snapshot into the nitrilase reaction mechanism: Once the benzonitrile has been oriented and positioned, the activated cysteine will perform the nucleophilic attack over the cyanocarbon atom in the nitrile, leading to the formation of a phenylmethanimine adduct, which is stabilized by one or the two glutamates (Fig. S7). After the incorporation of a water molecule an amino benzyl-alcohol tetrahedral intermediate is formed, followed by ammonia release and formation of a benzaldehyde adduct, incorporation of a second water molecule then breaks this intermediary and leads to the release of the benzoic acid product (Fig. 7).

Fig. 7. Proposed mechanism of the hydrolysis of nitrile-enzyme intermediates.

Fig. 7

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The path from benzonitrile to ammonia and benzoic acid: The carbon atom of the cyano group is attacked by the nucleophilic catalyst C165. An acyl intermediate is formed, and an ammonia molecule is released with the collaboration of a catalytic glutamate. The electron movements are shown as curved arrows and a dashed arrow indicate the release of amonia. A bridging water molecule was added according with the proposed nitrilase mechanism (Stevenson et al., 1990; Fernandes et al., 2006; Tang et al., 2023).

4. Discussion

Here we report a 3 Å structure of a filament-forming bacterial nitrilase from Rhodoccochus sp V51B. The truncated mutant (NitΔC39) shows the topology of the nitrilase family, and as in the case of AtNit4, a strongly packed dimer assembles into high-order oligomers forming a left-handed helical filament. In NitΔC39 the substrate binding pocket is located within the interface where three monomers interact, facing the exterior of the filament. We found that the helical filament structure is promoted and stabilized after the binding of the substrate, suggesting that due to the proximity of the active site to the dimerization and oligomerization interfaces, the dimer will sense structural changes in the active site that are translated to the rest of the filament, stabilizing the helical structure. Indeed, this relationship may explain why in AtNit4 pitch and rise are correlated with substrate specificity, given that changes in the oligomerization interfaces will lead to changes in the size and shape of the active site and its entrance. It also confirms that interfaces ‘A’ and ‘C’ are solely responsible for stabilizing the helical structure, allowing spiral elongation, as reported for AtNit4 (PDB 6I00). And, contrary to what has been reported previously (Thuku et al., 2007), no ‘D’ interface was observed, indicating that interactions between and across dimers are the main forces for the spiral formation. Furthermore, the influence of the quaternary structure on the architecture of the substrate-binding pocket has been recently noticed, in several helix-forming plant nitrilases a correlation between the helical twist of the filaments and the preference for different substrate sizes was observed, concluding that enzymes whose helical fibers have shorter twists show a preference for larger substrates and vice versa. The quaternary structure of nitrilase Nit6803 shows a short helical twist (Δφ = −60°), while AtNit4 has a longest helical twist (Δφ = −73°). Compared to these structures, the NitΔC39 filaments have a helical twist (Δφ = −72°) closest to AtNit4. However, like Nit6803, NitΔC39 shows broad substrate specificity, with a preference for small aromatic nitriles (Table S1, Fig. 2). Taking this into account, we see discrepancies when correlating these properties in the three enzymes.

In the NitΔC39 structure as in AtNIT4, the C-terminal end is located within the interior of the filament, forming part of interfaces ‘A’ and ‘C’, so its importance in the oligomerization mechanism and enzymatic activation has been highlighted. In AtNIT4 the last 10 C-terminal residues participate in completing an extended layer of four-stranded anti-parallel β-sheets in the inner core of the filament. Although in our model the last 10 residues remain unresolved, this segment could be completing the formation of this layer. In fact, the deletion of these 10 amino acid residues in the truncated mutant NitΔC49, abolishes its ability to assemble as helical filaments, until the binding of benzonitrile substrate induce its polymerization into active long filaments (Fig. S2). These structural changes that lead to filament elongation after the binding of the substrate, are shared in both truncated mutants regardless of their C-terminal length.

The NitΔC39 active site entrance is located near the dimerization and tetramerization interfaces, ‘A’ and ‘C’ respectively, and the catalytic residues E48, K131, E138, and C165 are positioned at the bottom of the substrate binding pocket, like other nitrilase superfamily members (Liu et al., 2013; Lundgren et al., 2008; Maurer et al., 2018; Mulelu, et al., 2019). This pocket is shown to be rich in aromatic residues and it has been suggested that for Nit6803 this aromaticity is necessary for the correct arrangement of the different aromatic substrates (Zhang et al., 2014). In fact, both enzymes display higher specificity towards aromatic substrates. The NitΔC39 catalytic pocket is only accessible through a funnel-shaped tunnel, like that of the amidases from Pseudomonas aeruginosa and P. abyssi, in which cases it has been suggested that some mobility in the neighboring residues allow the access of substrates and the release of reaction products. In yeast Nit2, due to the proximity of the active site entrance to the dimer interface, is proposed that the dimer is poised to sense conformational changes at the active site, which in turn influences the structure of the hairpin lid, triggering the enzymatic reaction (Liu et al., 2013).

The hydrolysis of benzonitrile is expected to follow the reaction mechanism proposed for the nitrilase superfamily. We hypothesize that the NitΔC39 nitrilase reaction mechanism proceeds through 3 steps; in the first step, a glutamic acid will activate the catalytic cysteine (C165) through a bridging water molecule, once activated the thiol group will perform a nucleophilic attack on the cyanocarbon atom in the nitrile (benzonitrile), concurrently to the proton transfer from the substrate to the lysine (K131), leading to the formation of a thioimidate intermediary (phenylmethanimine), the water molecule then attacks the cyanocarbon atom in the nitrile leading to the formation of a tetrahedral intermediate (amino-benzyl alcohol) which is stabilized by one or both catalytic glutamates (E48 and E138). In the second step, after the incorporation of a second water molecule, the intermediate breaks leading to the formation of ammonia and an acyl-enzyme (benzaldehyde). After the release of ammonia, this thioester intermediate undergoes a general base-catalyzed nucleophilic attack by the water molecule, forming a second stabilized tetrahedral intermediate. In the third step, the intermediate will collapse and spontaneously release the acid product (benzoic acid) restoring the enzyme (Figs. 7 and S7). The role of the invariant cysteine C165 as the catalytic nucleophile was elucidated by mutating Cys165 to Ala, which renders a completely inactive enzyme.

To our knowledge, a few atomic structures of nitrilase superfamily members with adducts in their active sites are available, these are the yNit2 in complex with alpha-KG and with OA (PDBs 4HG3 and 4HG5 respectively) (Liu et al., 2013), XC1258 from Xanthomonas campestris with an arsenic adduct in the active site (PDB 2E11) (Chin et al., 2007), the structure of the aliphatic amidase from P. aeruginosa showing a trapped acyl transfer reaction intermediate (PDB 2UXY) (Andrade et al., 2007), and a couple mutants of the amidase from Nesterenkonia sp. AN1 in complex with butyramide and acrylamide at the active site cysteine (PDBs 4IZS and 4IZU respectively) (Kimani et al., 2014). Superposition of these structures with that of NitΔC39, reveals a common architectural arrangement among the nitrilase superfamily (Fig. 8), suggesting that substrate selectivity and its posterior catalysis are mainly determined by the size and shape of the catalytic pocket, rather than by a specific configuration of the catalytic tetrad, which appears to be shared across these structures. Coordination, positioning, and stabilization of the nitrogen atom by the catalytic glutamate, and/or stabilization of the intermediary adducts by the catalytic lysine is observed in these structures. Although in NitΔC39 structure the carbonyl oxygen of the benzaldehyde adduct is not placed in the oxyanion hole, the visual analysis of the adduct modeled in the 14 chains, shows that in only one of them the position of this group is in close proximity to the amide backbone of W166 and the catalytic lysine K133 (Fig. S1F–h).

Fig. 8. Structural comparison of catalytic tetrads between NitΔC39 (gray) /S-benzoylcysteine (green) structure, and nitrilase superfamily members structures solved with covalently bound ligands or adducts in their active site.

Fig. 8

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PDB accession codes: 2E11) XC1258 from Xanthomonas campestris with an arsenic adduct, 2UXY) aliphatic amidase from Pseudomonas aeruginosa showing an acyl transfer reaction intermediate, 4HG3) yeast Nit2 in complex with alpha-KG, and 4IZU) amidase from Nesterenkonia sp AN1 in complex with acrylamide.

The detection of a reaction intermediary adducts by ion-spray mass spectroscopy, indicated that in nitrilases the rate of breakdown of the covalent intermediary determines the rate of hydrolysis (Stevenson et al., 1990). The present report confirms the existence of such intermediates and maps them covalently bound specifically to the invariant Cys165. At a final resolution of 3 Å we observe a map which is consistent with the position and orientation of the intermediary, the rotameric configuration of the catalytic residues, the architecture of the active site, and its relationship with the oligomerization interfaces (Figs. 5C, S1, S5). Higher resolution structures will clear our view on the positioning of the water molecules; also, distinct substrate-bound structures will decipher how the enzyme performs its hydrolytic activity towards diverse classes of substrates.

5. Conclusions

Recent developments have broadened the scope of the potential application of nitrilases. Their versatile biocatalytic nature and green applications by their capability to eliminate highly toxic nitriles play an important role in environmental protection and synthesis. The advances in the biotechnological processes, and a better understanding of the structure and the reaction mechanism would lead to improved properties of these enzymes, such as higher enzyme activity, higher stereo-specificity, and wide applicability over a range of pH and temperature. The structure reported here opens the way for further studies of the nitrilase specificity determinants and reaction mechanism. More biochemical and biophysical studies are necessary to decipher the oligomerization biological function and will facilitate the development of the biotechnological use of this class of enzymes. In summary, our work provides the structural framework explaining multiple previous observations based solely on chemical analysis and activity profiling of the nitrilase reaction mechanism and its relationship with the oligomerization process.

Supplementary Material

Supplementary data to this article can be found online at https://doi.org/10.1016/j.jsb.2024.108093.

Supplementary Data

Acknowledgements

S.A.S. thanks to Consejo Nacional de Humanidades, Ciencias y Tecnologías for the Ph.D. scholarship (CVU 486976) and Posgrado en Ciencias Bioquímicas-UNAM, for financial support. G.G.R. acknowledge support from DGAPA-PAPIIT (Dirección General de Asuntos del Personal Académico-Programa de Apoyo a Proyectos de Investigación e Innovación Tecnológica) grant IN218318, and from Consejo Nacional de Humanidades, Ciencias y Tecnologías-Ciencia Básica grant A1-S-7853. Rhodococcus sp V51B strain was kindly donated by Dr. Ignacio Regla. We thank, M.Sc. Eugenia Flores Robles for excellent technical assistance, Dra. Gloria Hernández Alcántara for help in obtaining the C169A mutant and Dr. Marco Igor Valencia for valuable suggestions for model building.

Footnotes

CRediT authorship contribution statement

Sergio Aguirre-Sampieri: Writing – original draft, Visualization, Validation, Methodology, Investigation, Formal analysis, Data curation, Conceptualization. Ana Casañal: Writing – original draft, Supervision, Software, Methodology, Formal analysis, Data curation, Conceptualization. Paul Emsley: Writing – review & editing, Supervision, Software, Methodology, Formal analysis, Data curation, Conceptualization. Georgina Garza-Ramos: Writing – original draft, Supervision, Resources, Project administration, Methodology, Investigation, Funding acquisition, Formal analysis, Conceptualization.

Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Data availability

Map and coordinates were deposited on the EMDB and wwPDB under the accession codes EMDB-42779 and 8UXU respectively.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supplementary Data
EMS196545-supplement-Supplementary_Data.docx (4MB, docx)

Data Availability Statement

Map and coordinates were deposited on the EMDB and wwPDB under the accession codes EMDB-42779 and 8UXU respectively.

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