Skip to main content
NCBI home page
Acta Crystallographica Section F: Structural Biology Communications logoLink to Acta Crystallographica Section F: Structural Biology Communications
. 2020 Jan 1;76(Pt 1):1–7. doi: 10.1107/S2053230X19015395

The RING domain of mitochondrial E3 ubiquitin ligase 1 and its complex with Ube2D2: crystallization and X-ray diffraction

Sang-Ok Lee a,b, Chong-Kil Lee b, Kyoung-Seok Ryu c,d,*, Seung-Wook Chi a,e,*
PMCID: PMC6957114  PMID: 31929179

This study presents the crystallization and X-ray diffraction analysis of the RING domain of mitochondrial E3 ubiquitin ligase 1 (MUL1-RING) and its complex with Ube2D2.

Keywords: mitochondrial E3 ubiquitin ligase 1, MUL1, RING domain, MUL1-RING, Ube2D2, ubiquitylation, crystallization

Abstract

Mitochondrial E3 ubiquitin ligase 1 (MUL1) is located in the mitochondrial outer membrane and regulates various biological processes, including apoptosis, cell growth, mitophagy and mitochondrial dynamics. The C-terminal region of MUL1 faces the cytoplasm and contains the RING domain (MUL1-RING) where the Ub~E2 thioester binds. Unlike most RING-type E3 enzymes, MUL1-RING alone does not have an additional region that recruits a substrate protein, yet is still able to ubiquitylate the substrate, the p53 protein. Nevertheless, the exact mechanism of the ubiquitylation of p53 by MUL1-RING has not yet been elucidated. In order to understand this novel ubiquitylation mechanism, it is necessary to determine the three-dimensional structures of MUL1-RING and of its complex with the cognate E2 enzyme. Here, Ube2D2 was validated as a functional E2 enzyme for the ubiquitylation of the p53 transactivation domain (p53-TAD) by MUL1-RING, and purification and crystallization processes for MUL1-RING and the MUL1-RING–Ube2D2 complex are reported.

1. Introduction  

Mitochondrial E3 ubiquitin ligase 1 (MUL1) is a mitochondrial E3 ligase that consists of a small cytoplasmic region (residues 1–8), a transmembrane-1 domain (TM-1; residues 9–29) domain, a mitochondrial intermembrane region (residues 30–238), a transmembrane-2 domain (TM-2; residues 239–259) and a C-terminal cytoplasmic region (residues 260–352) that contains the MUL1-RING domain. The ubiquitylation activity of MUL1 is important for the regulation of mitochondrial dynamics (fusion and fission), including mitophagy (Braschi et al., 2009 ▸), and is also involved in various biological processes such as apoptosis and cell growth (Neuspiel et al., 2008 ▸; Zhang et al., 2008 ▸). MUL1 regulates the activation of NF-κB signaling, which causes stress-induced hyperfusion of mitochondria (Zemirli et al., 2014 ▸). The presence of MUL1-RING is critical for both the ubiquitylation (Li et al., 2008 ▸) and SUMOylation (Braschi et al., 2009 ▸) activities of MUL1. MUL1 promotes the degradation of ULK1 (unc-51-like autophagy activating kinase 1) and MFN-2 (mitofusin 2) through ubiquitylation (Yun et al., 2014 ▸). On the other hand, MUL1 induces the SUMOylation of DNM1L/Drp1 to increase mitochondrial fission (Nunnari & Suomalainen, 2012 ▸) and delays mitochondrial fusion by inhibiting ubiquitylation (Deng et al., 2008 ▸).

Ubiquitylation is mediated by an enzymatic cascade reaction involving a ubiquitin-activating enzyme (E1), a ubiquitin-conjugating enzyme (E2) and a ubiquitin ligase enzyme (E3) (Pickart & Eddins, 2004 ▸). The repetitive process of ubiquitin transfer leads to the polyubiquitylation of substrate proteins and their degradation by the 26S proteasome; the attached ubiquitin is then recycled, contributing to free ubiquitin (Komander & Rape, 2012 ▸). E3 ligases can be categorized into two main classes according to their mechanism of ubiquitin transfer. HECT-E3 ligases receive activated ubiquitin that is attached to E2 via a thioester (E2~UB) to form an intermediate, HECT-E3~UB, and then transfer ubiquitin to the target protein. In contrast, RING-E3 ligases directly deliver ubiquitin from E2~UB to the target protein (Pruneda et al., 2012 ▸; Metzger et al., 2014 ▸; Deshaies & Joazeiro, 2009 ▸), in which a lysine ∊-amino group of the target protein attacks the carbonyl group of the E2~UB thioester and results in the formation of an isopeptide bond between the C-terminal Gly76 of ubiquitin and the lysine side chain of the target. The key feature of most RING-type E3 ligases is the presence of both a RING domain and another domain that recruit E2~UB and target protein, respectively (Plechanovová et al., 2012 ▸).

MUL1-RING has been shown to play an important role in apoptosis, cell growth and the regulation of mitochondrial function. Interestingly, MUL1-RING alone is able to bind to and ubiquitylate the substrate protein p53 (Jung et al., 2011 ▸; Peng et al., 2016 ▸). We recently reported the solution structure of MUL1-RING and analyzed the interaction between MUL1-RING and the p53 transactivation domain (p53-TAD) using NMR spectroscopy (Lee et al., 2019 ▸). However, the binding affinity between these two proteins is very weak, and the exact mechanism of MUL1-RING-mediated ubiquitylation of p53-TAD is still not clear. Therefore, it is important to determine the 3D structures of both free MUL1-RING and its complex with the cognate E2 enzyme in order to elucidate the unique ubiquitylation mechanism of MUL1-RING. Previous studies of the ubiquitylation activity of MUL1 with various E2 enzymes have shown that Ube2D1 and Ube2D3, but not Ube2D2, play a role in the ubiquitylation of p53 (Jung et al., 2011 ▸). Since these human E2 enzymes belong to the yeast Ubc4 family and their amino-acid sequences are almost identical (Dou et al., 2012 ▸; Ozkan et al., 2005 ▸), it is necessary to validate the enzymatic activity of these E2 enzymes during the ubiquitylation of p53-TAD by MUL1-RING alone. Here, we selected Ube2D2 for complex formation with MUL1-RING based on an in vitro ubiquitylation assay and report the purification and crystallization processes of MUL1-RING and the MUL1-RING–Ube2D2 complex in order to study the novel ubiquitylation mechanism of MUL1-RING alone.

2. Materials and methods  

2.1. Expression and purification  

To express the target proteins, the genes for human MUL1-RING (residues 298–352) and Ube2D2 (residue 1–147) were cloned into the pGEX4T-3 vector using the BamHI and XhoI restriction-enzyme sites (Table 1 ▸). The recombinant plasmid was transformed into Escherichia coli Rosetta 2 (DE3) cells. Expression of the glutathione S-transferase (GST)-tagged proteins was induced by incubation with 0.5 mM isopropyl β-d-1-thiogalactopyranoside for 6 h at 30°C. The GST-fusion proteins were purified by GST-affinity column chroma­tography with phosphate-buffered saline (PBS) buffer and the GST tag was removed by thrombin digestion. The MUL1-RING protein was further purified by size-exclusion chroma­tography (SEC) on a HiLoad 16/600 Superdex 75 column (GE Healthcare) with SEC buffer (50 mM MES, 50 mM NaCl, 5 µM ZnSO4, 10 mM DTT pH 6.5). The E2 protein was purified by SEC with a buffer consisting of 10 mM Tris–HCl, 100 mM NaCl, 1 mM DTT pH 7.5.

Table 1. Production information for MUL1-RING and Ube2D2.

  MUL1-RING Ube2D2
Source organism H. sapiens H. sapiens
DNA source H. sapiens genomic DNA H. sapiens genomic DNA
Forward primer† (5′–3′) GAGGATCCAGTCTGAAGAGCGCCTG GAGGATCCATGGCTCTGAAGAGAATC
Reverse primer‡ (5′–3′) GACTCGAGTTAGCTGTTGTACAGGGGTATC GACTCGAGTTACATCGCATACTTCTGAGTC
Cloning and expression vector pGEX4T-3 pGEX4T-3
Cloning host E. coli DH5α E. coli DH5α
Expression host E. coli Rosetta 2 (DE3) E. coli Rosetta 2 (DE3)
Amino-acid sequence§ GSLKSACVVCLSSFKSCVFLECGHVCSCTECYRALPEPKKCPICRQAITRVIPLYNS GSMALKRIHKELNDLARDPPAQCSAGPVGDDMFHWQATIMGPNDSPYQGGVFFLTIHFPTDYPFKPPKVAFTTRIYHPNINSNGSICLDILRSQWSPALTISKVLLSICSLLCDPNPDDPLVPEIARIYKTDREKYNRIAREWTQKYAM
Open in a new tab
†

The BamHI site is underlined.

‡

The XhoI site is underlined.

§

After thrombin digestion.

The formation of a complex between MUL1-RING and Ube2D2 was first confirmed by SEC high-performance liquid chromatography (HPLC) using a Biosep-SEC-S3000 column (Phenomenex) with SEC buffer. To obtain the MUL1-RING–Ube2D2 complex, a mixture of MUL1-RING and Ube2D2 in a 1:1 molar ratio was prepared using their UV extinction coefficients (Gill & von Hippel, 1989 ▸) and the mixture was further purified using the same SEC procedure as described for MUL1-RING alone. All purified proteins were concentrated using Amicon Ultra Centrifugal filters (Millipore) for crystallization setup.

2.2. In vitro ubiquitylation assay  

The in vitro ubiquitylation of p53-TAD by MUL1-RING alone was assessed with four different E2 enzymes, Ube2D1, Ube2D2, Ube2D3 and Ube2L3 (UbcH7), following a previously reported method (Choi et al., 2015 ▸). Briefly, the in vitro ubiquitylation was performed with 20 µM GST-fused MUL1-RING, 200 µM ubiquitin, 1 µM E1, 20 µM E2 and 20 µM p53-TAD in a buffer consisting of 50 mM Tris–HCl, 10 µM ZnSO4, 1 mM MgCl2, 2 mM ATP pH 8.0 for 2.5 h at 25°C. The reaction samples were analyzed by sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS–PAGE) and the protein bands were visualized by Coomassie Blue staining. Ube2L3 was used as a negative control, since it does not have E3-independent reactivity with the lysine residue and has a preference for HECT-E3 ligases (Wenzel et al., 2011 ▸).

2.3. Crystallization of MUL1-RING and the MUL1-RING–Ube2D2 complex  

The MUL1-RING protein (15 mg ml−1) was prepared in SEC buffer for crystallization setup. Initial sparse-matrix screening of the crystallization conditions was performed using commercial crystallization solution kits (Index, Crystal Screen, PEG/Ion, SaltRx and MembFac from Hampton Research and Wizard Classic from Molecular Dimensions) at 20°C. The crystals of MUL1-RING alone grew in five days using the sitting-drop vapor-diffusion method with 100 µl reservoir buffer consisting of 0.1 M bis-Tris pH 6.5, 0.2–0.4 M lithium sulfate monohydrate, 25%(w/v) polyethylene glycol (PEG) 3350, in which 1 µl protein solution and 1 µl reservoir solution were mixed together (Table 2 ▸). The cryoprotectant solution for the crystal of MUL1-RING alone was prepared by adding 25% glycerol to the reservoir solution.

Table 2. Crystallization conditions.

  MUL1-RING MUL1-RING–Ube2D2
Method Sitting-drop vapor diffusion Hanging-drop vapor diffusion
Plate type 96-well 2-drop MRC Crystallization Plates (Molecular Dimensions) 24-well VDX plate (Hampton Research)
Temperature (K) 293 293
Protein concentration (mg ml−1) 15 10
Buffer composition of protein solution 50 mM MES, 50 mM NaCl, 5 µM ZnSO4, 10 mM DTT pH 6.5 50 mM MES, 50 mM NaCl, 5 µM ZnSO4, 10 mM DTT pH 6.5
Composition of reservoir solution 0.4 M lithium sulfate monohydrate, 0.1 M bis-Tris pH 6.5, 25%(w/v) PEG 3350 4%(v/v) Tacsimate pH 6.0, 12%(w/v) PEG 3350
Volume and ratio of drop 2 µl (1:1 ratio) 2 µl (1:1 ratio)
Volume of reservoir (µl) 100 1000
Open in a new tab

The MUL1-RING–Ube2D2 complex (10 mg ml−1) was prepared in SEC buffer and sparse-matrix screening was then performed using a Mosquito Crystal nanolitre protein crystallization robot (TTP Labtech) with similar crystallization solution kits (Index, Crystal Screen and PEG/Ion from Hampton Research and ProPlex from Molecular Dimensions) at 20°C. The MUL1-RING (10 mg ml−1) and Ube2D2 (10 mg ml−1) proteins were also used separately to distinguish the complex crystals from crystals consisting of a single protein component during sparse-matrix screening. After various optimizations of the crystallization conditions, crystals of the MUL1-RING–Ube2D2 complex grew in three days using the hanging-drop vapor-diffusion method with 1 ml reservoir buffer consisting of 4%(v/v) Tacsimate pH 6.0, 12%(w/v) PEG 3350 (Table 2 ▸). The cryoprotectant solution for the crystal of the MUL1-RING–Ube2D2 complex was prepared by increasing the concentration of PEG 3350 in the reservoir solution to 40%(w/v).

2.4. Data collection and processing for MUL1-RING and the MUL1-RING–Ube2D2 complex  

All X-ray diffraction data were recorded on beamline 7A at Pohang Accelerator Laboratory (PAL), Republic of Korea. Single crystals of MUL1-RING and the MUL1-RING–Ube2D2 complex were transferred into the cryoprotectant solution using a cryoloop and were then flash-cooled with cold nitrogen gas (100 K). The X-ray diffraction data were indexed, merged and scaled using the HKL-2000 software (Otwinowski & Minor, 1997 ▸). The data-collection statistics are summarized in Table 3 ▸.

Table 3. X-ray diffraction and data-collection statistics for MUL1-RING and the MUL1-RING–Ube2D2 complex.

Values in parentheses are for the highest resolution shell.

  MUL1-RING MUL1-RING–Ube2D2
Diffraction source Beamline 7A, PAL Beamline 7A, PAL
Wavelength (Å) 1.2837 0.9793
Temperature (K) 100 100
Detector ADSC Q270r ADSC Q270r
Crystal-to-detector distance (mm) 150 350
Rotation range per image (°) 1.0 1.0
Total rotation range (°) 360 360
Exposure time per image (s) 0.2 1.0
Space group P212121 P21
a, b, c (Å) 59.226, 66.593, 68.271 47.207, 141.241, 68.047
α, β, γ (°) 90.00, 90.00, 90.00 90.00, 104.81, 90.00
Mosaicity range (°) 0.707–1.103 0.850
Resolution range (Å) 50.00–1.80 50.00–2.70
Total No. of reflections 1034568 453307
No. of unique reflections 25532 21200
Completeness (%) 99.0 (96.8) 90.1 (74.8)
Multiplicity 8.4 (4.4) 5.1 (3.0)
R meas 0.084 (0.407) 0.093 (0.337)
〈I/σ(I)〉 24.5 (2.3) 20.2 (3.3)
Wilson B factor (Å2) 16.9 51.6
Open in a new tab

3. Results and discussion  

We first examined whether Ube2D1, Ube2D2 and Ube2D3, the sequences of which are almost identical (Fig. 1 ▸ a), were able to support the ubiquitylation of p53-TAD by MUL1-RING alone, since (i) a previous report showed that Ube2D2 was not effective in the ubiquitylation of p53 by MUL1 (Jung et al., 2011 ▸) and (ii) the backbone chemical shifts of Ube2D2 were already available, which may facilitate further NMR studies based on the crystal structure of the MUL1-RING–Ube2D2 complex. The in vitro ubiquitylation of p53-TAD by MUL1-RING alone was assessed for Ube2D1, Ube2D2, Ube2D3 and Ube2L3, in which Ube2L3 was used as a negative control. As predicted, all three E2 enzymes, but not Ube2L3, were able to support polyubiquitylation by MUL1-RING alone (Fig. 1 ▸ b). Next, we also confirmed that MUL1-RING formed a stable complex with Ube2D2: SEC–HPLC and SDS–PAGE analyses showed that MUL1-RING co-eluted with Ube2D2 (Fig. 2 ▸).

Figure 1.

Figure 1

Open in a new tab

An in vitro ubiquitylation assay by MUL1-RING alone. (a) Multiple sequence alignment of Ube2D1, Ube2D2, Ube2D3 and Ube2L3. Sequence alignment of Ube2D1, Ube2D2, Ube2D3 and Ube2L3 was carried out using Clustal Omega and ESPript 3.0 (Sievers & Higgins, 2018 ▸; Robert & Gouet, 2014 ▸). (b) The in vitro ubiquitylation of p53-TAD mediated by MUL1-RING was assessed for Ube2D1, Ube2D2 and Ube2D3. Ube2L3 (UbcH7) was used as a negative control. Molecular-weight markers are labeled in kDa.

Figure 2.

Figure 2

Open in a new tab

SEC elution profile of the MUL1-RING–Ube2D2 mixture. MUL1-RING, Ube2D2 and their mixture were analyzed by (a) SEC–HPLC and (b) SDS–PAGE. In (b), lane M contains molecular-weight markers (labeled in kDa), lane 1 contains Ube2D2, lane 2 contains MUL1-RING and lane 3 contains MUL1-RING–Ube2D2.

MUL1-RING crystals were obtained using the sitting-drop method (Fig. 3 ▸ a). X-ray diffraction data from a MUL1-RING crystal were collected to 1.80 Å resolution (Fig. 4 ▸ a). The crystal belonged to the orthorhombic space group P212121 (unit-cell parameters a = 59.22, b = 66.59, c = 68.27 Å). The data set, with a resolution range of 50–1.80 Å, had 99% completeness and an R merge of 8.4% (Table 3 ▸). Assuming the presence of four MUL1-RING molecules in the asymmetric unit, the corresponding Matthews coefficient (Matthews, 1968 ▸) and solvent content are 2.67 Å3 Da−1 and 53.98%, respectively. To solve the crystal structure of MUL1-RING, we performed molecular replacement (MR) using MOLREP (Vagin & Teplyakov, 2010 ▸) and Phaser (McCoy et al., 2007 ▸). In addition to the recent NMR ensemble structures of MUL1-RING (PDB entry 6k2k; Lee et al., 2019 ▸), a homology model of MUL1-RING was calculated using the Phyre2 web portal (Kelley et al., 2015 ▸), in which the structure of baculoviral IAP repeat-containing protein 2 (PDB entry 3t6p; 41% identity; Dueber et al., 2011 ▸) was used as a reference model. Unfortunately, neither model was able to derive a correct MR solution. The failure of MR with NMR ensemble structures of MUL1-RING is interesting, since the structures were calculated with NH residual dipolar coupling data in addition to NOE distance and Talos angle restraints. The crystal structure of MUL1-RING might differ from the solution structure, or possible structural flexibility of MUL1-RING in solution may cause the failure of MR. To study the unique ubiquitylation mechanism of MUL1-RING, the structure of the complex of MUL1-RING with its cognate E2 enzyme is critical. Therefore, the crystal structure of MUL1-RING alone can simply be solved after determination of the complex structure.

Figure 3.

Figure 3

Open in a new tab

(a, b) Crystals of human MUL1-RING (a) and of MUL1-RING complexed with Ube2D2 (b). (c) SDS–PAGE of a dissolved crystal of the MUL1-RING–Ube2D2 complex. Molecular-weight markers are labeled in kDa.

Figure 4.

Figure 4

Open in a new tab

X-ray diffraction patterns of crystals of MUL1-RING (a) and of MUL1-RING complexed with Ube2D2 (b). The diffraction images were collected on beamline 7A at PAL.

The crystallization conditions for the MUL1-RING–Ube2D2 protein complex were screened using the sitting-drop method, and the final crystal form was obtained using the hanging-drop method after various crystal optimizations (Fig. 3 ▸ b). In addition, we confirmed that the crystal actually contained MUL1-RING and Ube2D2 by SDS–PAGE of a dissolved crystal of the complex (Fig. 3 ▸ c). X-ray diffraction data were collected from a crystal of the complex to 2.7 Å resolution (Fig. 4 ▸ b). The crystal symmetry was determined to be monoclinic (space group P21; unit-cell parameters a = 47.21, b = 141.24, c = 68.05 Å, α = γ = 90.00, β = 104.81°) and the statistics of the diffraction data are summarized in Table 3 ▸. The calculated Matthews coefficient (V M) was 2.38 Å3 Da−1 with a solvent content of 48.26%, and thus four complex molecules appeared to be present in the asymmetric unit. We attempted to find an MR solution for the X-ray diffraction data from the MUL1-RING–Ube2D2 complex using the same NMR structures and the homologous model of MUL1-RING, but these trials were not successful. However, a Phaser MR analysis using a previous crystal structure of Ube2D2 (PDB entry 2esk; Ozkan et al., 2005 ▸) clearly found four solutions with TFZ values of 13.0, 24.9, 26.0 and 30.0.

It would be interesting to observe whether there are any structural differences in MUL1-RING among the NMR ensemble structures of MUL1-RING, the crystal structure of MUL1-RING alone and the crystal structure of the MUL1-RING–Ube2D2 complex that may explain the reason why MR using the NMR ensemble structure of MUL1-RING failed and may reveal unique structural features of the MUL1-RING–Ube2D2 complex compared with other RING-E2 proteins. The 3D structures of MUL1-RING and the MUL1-RING–Ube2D2 complex are likely to provide a structural basis for understanding the unique ubiquitylation mechanism of MUL1-RING alone that distinguishes it from other RING-E3 proteins.

Acknowledgments

The authors are grateful to the staff of beamline 7A at Pohang Accelerator Laboratory, Republic of Korea and to Dr Jin-Sik Kim for helpful discussions.

Funding Statement

This work was funded by National Research Foundation of Korea grants NRF-2017R1E1A1A01074403, NRF-2019M3E5D4069903, and NRF-2019M3A9C4076156. Korea Basic Science Institute grant T39632 to Kyoung-Seok Ryu. Korea Research Institute of Bioscience and Biotechnology grant .

References

  1. Braschi, E., Zunino, R. & McBride, H. M. (2009). EMBO Rep. 10, 748–754. [DOI] [PMC free article] [PubMed]
  2. Choi, Y.-S., Lee, Y.-J., Lee, S.-Y., Shi, L., Ha, J.-H., Cheong, H.-K., Cheong, C., Cohen, R. E. & Ryu, K.-S. (2015). J. Biol. Chem. 290, 2251–2263. [DOI] [PMC free article] [PubMed]
  3. Deng, H., Dodson, M. W., Huang, H. & Guo, M. (2008). Proc. Natl Acad. Sci. USA, 105, 14503–14508. [DOI] [PMC free article] [PubMed]
  4. Deshaies, R. J. & Joazeiro, C. A. (2009). Annu. Rev. Biochem. 78, 399–434. [DOI] [PubMed]
  5. Dou, H., Buetow, L., Sibbet, G. J., Cameron, K. & Huang, D. T. (2012). Nat. Struct. Mol. Biol. 19, 876–883. [DOI] [PMC free article] [PubMed]
  6. Dueber, E. C., Schoeffler, A. J., Lingel, A., Elliott, J. M., Fedorova, A. V., Giannetti, A. M., Zobel, K., Maurer, B., Varfolomeev, E., Wu, P., Wallweber, H. J., Hymowitz, S. G., Deshayes, K., Vucic, D. & Fairbrother, W. J. (2011). Science, 334, 376–380. [DOI] [PubMed]
  7. Gill, S. C. & von Hippel, P. H. (1989). Anal. Biochem. 182, 319–326. [DOI] [PubMed]
  8. Jung, J. H., Bae, S., Lee, J. Y., Woo, S. R., Cha, H. J., Yoon, Y., Suh, K.-S., Lee, S.-J., Park, I.-C., Jin, Y.-W., Lee, K.-H., An, S. & Lee, J. H. (2011). Cell Death Differ. 18, 1865–1875. [DOI] [PMC free article] [PubMed]
  9. Kelley, L. A., Mezulis, S., Yates, C. M., Wass, M. N. & Sternberg, M. J. E. (2015). Nat. Protoc. 10, 845–858. [DOI] [PMC free article] [PubMed]
  10. Komander, D. & Rape, M. (2012). Annu. Rev. Biochem. 81, 203–229. [DOI] [PubMed]
  11. Lee, M.-S., Lee, S.-O., Lee, M.-K., Yi, G.-S., Lee, C.-K., Ryu, K.-S. & Chi, S.-W. (2019). Biochem. Biophys. Res. Commun. 516, 533–539. [DOI] [PubMed]
  12. Li, W., Bengtson, M. H., Ulbrich, A., Matsuda, A., Reddy, V. A., Orth, A., Chanda, S. K., Batalov, S. & Joazeiro, C. A. (2008). PLoS One, 3, e1487. [DOI] [PMC free article] [PubMed]
  13. Matthews, B. W. (1968). J. Mol. Biol. 33, 491–497. [DOI] [PubMed]
  14. McCoy, A. J., Grosse-Kunstleve, R. W., Adams, P. D., Winn, M. D., Storoni, L. C. & Read, R. J. (2007). J. Appl. Cryst. 40, 658–674. [DOI] [PMC free article] [PubMed]
  15. Metzger, M. B., Pruneda, J. N., Klevit, R. E. & Weissman, A. M. (2014). Biochim. Biophys. Acta, 1843, 47–60. [DOI] [PMC free article] [PubMed]
  16. Neuspiel, M., Schauss, A. C., Braschi, E., Zunino, R., Rippstein, P., Rachubinski, R. A., Andrade-Navarro, M. A. & McBride, H. M. (2008). Curr. Biol. 18, 102–108. [DOI] [PubMed]
  17. Nunnari, J. & Suomalainen, A. (2012). Cell, 148, 1145–1159. [DOI] [PMC free article] [PubMed]
  18. Otwinowski, Z. & Minor, W. (1997). Methods Enzymol. 276, 307–326. [DOI] [PubMed]
  19. Ozkan, E., Yu, H. & Deisenhofer, J. (2005). Proc. Natl Acad. Sci. USA, 102, 18890–18895. [DOI] [PMC free article] [PubMed]
  20. Peng, J., Ren, K.-D., Yang, J. & Luo, X.-J. (2016). Mitochondrion, 28, 49–53. [DOI] [PubMed]
  21. Pickart, C. M. & Eddins, M. J. (2004). Biochim. Biophys. Acta, 1695, 55–72. [DOI] [PubMed]
  22. Plechanovová, A., Jaffray, E. G., Tatham, M. H., Naismith, J. H. & Hay, R. T. (2012). Nature (London), 489, 115–120. [DOI] [PMC free article] [PubMed]
  23. Pruneda, J. N., Littlefield, P. J., Soss, S. E., Nordquist, K. A., Chazin, W. J., Brzovic, P. S. & Klevit, R. E. (2012). Mol. Cell, 47, 933–942. [DOI] [PMC free article] [PubMed]
  24. Robert, X. & Gouet, P. (2014). Nucleic Acids Res. 42, W320–W324. [DOI] [PMC free article] [PubMed]
  25. Sievers, F. & Higgins, D. G. (2018). Protein Sci. 27, 135–145. [DOI] [PMC free article] [PubMed]
  26. Vagin, A. & Teplyakov, A. (2010). Acta Cryst. D66, 22–25. [DOI] [PubMed]
  27. Wenzel, D. M., Lissounov, A., Brzovic, P. S. & Klevit, R. E. (2011). Nature (London), 474, 105–108. [DOI] [PMC free article] [PubMed]
  28. Yun, J., Puri, R., Yang, H., Lizzio, M. A., Wu, C., Sheng, Z.-H. & Guo, M. (2014). Elife, 3, e01958. [DOI] [PMC free article] [PubMed]
  29. Zemirli, N., Pourcelot, M., Ambroise, G., Hatchi, E., Vazquez, A. & Arnoult, D. (2014). FEBS J. 281, 3095–3112. [DOI] [PubMed]
  30. Zhang, B., Huang, J., Li, H.-L., Liu, T., Wang, Y.-Y., Waterman, P., Mao, A.-P., Xu, L.-G., Zhai, Z., Liu, D., Marrack, P. & Shu, H.-B. (2008). Cell Res. 18, 900–910. [DOI] [PMC free article] [PubMed]

Articles from Acta Crystallographica. Section F, Structural Biology Communications are provided here courtesy of International Union of Crystallography

RESOURCES