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Snapshots of the RNA editing machine in trypanosomes captured at different assembly stages in vivo

2009; Springer Nature; Volume: 28; Issue: 6 Linguagem: Inglês

10.1038/emboj.2009.19

1460-2075

Monika M. Golas, Cordula Böhm, Bjoern Sander, Kerstin A. Effenberger, Michael Brecht, Holger Stark, H. Ulrich Göringer,

CRISPR and Genetic Engineering

Article5 February 2009free access Snapshots of the RNA editing machine in trypanosomes captured at different assembly stages in vivo Monika M Golas Monika M Golas Research Group of 3D Electron Cryomicroscopy, Max Planck Institute for Biophysical Chemistry, Göttingen, Germany Institute of Anatomy, University of Aarhus, Århus, Denmark Search for more papers by this author Cordula Böhm Cordula Böhm Department of Genetics, Darmstadt University of Technology, Darmstadt, Germany Search for more papers by this author Bjoern Sander Bjoern Sander Research Group of 3D Electron Cryomicroscopy, Max Planck Institute for Biophysical Chemistry, Göttingen, Germany Stereology and EM Research Laboratory, University of Aarhus, Århus, Denmark Search for more papers by this author Kerstin Effenberger Kerstin Effenberger Department of Genetics, Darmstadt University of Technology, Darmstadt, Germany Search for more papers by this author Michael Brecht Michael Brecht Department of Genetics, Darmstadt University of Technology, Darmstadt, Germany Search for more papers by this author Holger Stark Corresponding Author Holger Stark Research Group of 3D Electron Cryomicroscopy, Max Planck Institute for Biophysical Chemistry, Göttingen, Germany Göttingen Centre for Molecular Biology, University of Göttingen, Göttingen, Germany Search for more papers by this author H Ulrich Göringer Corresponding Author H Ulrich Göringer Department of Genetics, Darmstadt University of Technology, Darmstadt, Germany Search for more papers by this author Monika M Golas Monika M Golas Research Group of 3D Electron Cryomicroscopy, Max Planck Institute for Biophysical Chemistry, Göttingen, Germany Institute of Anatomy, University of Aarhus, Århus, Denmark Search for more papers by this author Cordula Böhm Cordula Böhm Department of Genetics, Darmstadt University of Technology, Darmstadt, Germany Search for more papers by this author Bjoern Sander Bjoern Sander Research Group of 3D Electron Cryomicroscopy, Max Planck Institute for Biophysical Chemistry, Göttingen, Germany Stereology and EM Research Laboratory, University of Aarhus, Århus, Denmark Search for more papers by this author Kerstin Effenberger Kerstin Effenberger Department of Genetics, Darmstadt University of Technology, Darmstadt, Germany Search for more papers by this author Michael Brecht Michael Brecht Department of Genetics, Darmstadt University of Technology, Darmstadt, Germany Search for more papers by this author Holger Stark Corresponding Author Holger Stark Research Group of 3D Electron Cryomicroscopy, Max Planck Institute for Biophysical Chemistry, Göttingen, Germany Göttingen Centre for Molecular Biology, University of Göttingen, Göttingen, Germany Search for more papers by this author H Ulrich Göringer Corresponding Author H Ulrich Göringer Department of Genetics, Darmstadt University of Technology, Darmstadt, Germany Search for more papers by this author Author Information Monika M Golas1,2, Cordula Böhm3, Bjoern Sander1,4, Kerstin Effenberger3, Michael Brecht3, Holger Stark 1,5 and H Ulrich Göringer 3 1Research Group of 3D Electron Cryomicroscopy, Max Planck Institute for Biophysical Chemistry, Göttingen, Germany 2Institute of Anatomy, University of Aarhus, Århus, Denmark 3Department of Genetics, Darmstadt University of Technology, Darmstadt, Germany 4Stereology and EM Research Laboratory, University of Aarhus, Århus, Denmark 5Göttingen Centre for Molecular Biology, University of Göttingen, Göttingen, Germany *Corresponding authors: 3D Electron Cryomicroscopy Group, Max Planck Institute for Biophysical Chemistry, Am Faßberg 11, 37077 Göttingen, Germany. Tel.: +49 551 201 1350; Fax: +49 551 201 1197; E-mail: [email protected] of Genetics, Darmstadt University of Technology, Schnittspahnstraße 10, 64287 Darmstadt, Germany. Tel.: +49 6151 16 28 55; Fax: +49 6151 16 56 40; E-mail: [email protected] The EMBO Journal (2009)28:766-778https://doi.org/10.1038/emboj.2009.19 PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info Mitochondrial pre-messenger RNAs in kinetoplastid protozoa are substrates of uridylate-specific RNA editing. RNA editing converts non-functional pre-mRNAs into translatable molecules and can generate protein diversity by alternative editing. Although several editing complexes have been described, their structure and relationship is unknown. Here, we report the isolation of functionally active RNA editing complexes by a multistep purification procedure. We show that the endogenous isolates contain two subpopulations of ∼20S and ∼35–40S and present the three-dimensional structures of both complexes by electron microscopy. The ∼35–40S complexes consist of a platform density packed against a semispherical element. The ∼20S complexes are composed of two subdomains connected by an interface. The two particles are structurally related, and we show that RNA binding is a main determinant for the interconversion of the two complexes. The ∼20S editosomes contain an RNA-binding site, which binds gRNA, pre-mRNA and gRNA/pre-mRNA hybrid molecules with nanomolar affinity. Variability analysis indicates that subsets of complexes lack or possess additional domains, suggesting binding sites for components. Together, a picture of the RNA editing machinery is provided. Introduction The RNA editing reaction in kinetoplastid protozoa such as Trypanosoma and Leishmania is characterised by a reaction cycle that inserts and deletes uridylates (U) into otherwise incomplete mitochondrial transcripts. Biochemically, the reaction can be divided into three steps, which are catalysed by proteins only. Proteins involved in RNA editing are assembled in large macromolecular machines known as editosomes (Madison-Antenucci et al, 2002; Stuart et al, 2005). The reaction is initiated by the formation of a guide (g)RNA/pre-edited mRNA duplex (Blum et al, 1990). This duplex functions as substrate for an endonuclease that cleaves the pre-mRNA at the first mismatch 5′ of the duplex. During insertion editing, a 3′-terminal uridylyl-transferase (TUTase) adds a number of U-nucleotides to the 3′-end of the 5′ pre-mRNA fragment, whereas for deletion editing, a 3′ → 5′ exoribonuclease removes terminal uridylates from the 5′ pre-mRNA fragment. Finally, an RNA ligase joins the two mRNA fragments. However, editosomes do not only contain endoribonuclease (Brecht et al, 2005; Carnes et al, 2005; Trotter et al, 2005), TUTase (Ernst et al, 2003), exoribonuclease (Brecht et al, 2005; Mian et al, 2006; Rogers et al, 2007) and RNA ligase activities (McManus et al, 2001), but also various additional polypeptides whose functions are poorly defined (Aphasizhev et al, 2003; Panigrahi et al, 2003a, 2003b, 2006). Although significant progress has been made to unravel the protein inventory of editing complexes (Stuart et al, 2005), little is known as to the structural organisation and dynamics of this molecular machine. Especially, even though several editing complexes have been described, their relationship in the context of the catalytic cycle is not understood. Editing complexes include a 5–10S insertion subcomplex and a 5–10S deletion subcomplex each of which comprising three different proteins, an ∼20S complex that contains up to 20 proteins including those of both the 5–10S insertion and deletion subcomplexes as well as a 35–40S complex with biochemically poorly defined composition (Pollard et al, 1992; Schnaufer et al, 2003; Panigrahi et al, 2006). The presence of specific RNA classes has been shown for the 35–40S complex, which harbours both, gRNA and pre-mRNA (Pollard et al, 1992), whereas gRNAs are found only in a subset of ∼20S editing complexes (Rusché et al, 1997; Madison-Antenucci et al, 1998). In addition, it has been suggested that an 11S gRNA complex may be generated by the interaction of a 5S complex with gRNA serving as gRNA storage (Goulah and Read, 2007). The understanding of the assembly of the RNA editing machinery is far from complete. A current model involves a stepwise pathway (Madison-Antenucci et al, 2002). An ∼20S gRNA maturation complex is formed by the interaction of polycistronic gRNAs with a pre-assembled core complex. This pre-assembled complex may be derived by the interaction of the 5–10S insertion and deletion subcomplexes with other proteins (Schnaufer et al, 2003). Polycistronic gRNAs are suggested to be cleaved in the ∼20S gRNA maturation complex (Grams et al, 2000). Ultimately, this ∼20S complex may interact with pre-edited/partially edited pre-mRNAs, resulting in the formation of an ∼35–40S complex, which was suggested to perform the catalytic reactions (Pollard et al, 1992). Together, this assembly model suggests that the 20S complex may be converted into the ∼35–40S particle and thus, both complexes may share a common core structure. However, the described data would also be consistent with an alternative model in which both complexes may represent functionally separate particles with individual assembly pathways in vivo. Due to their different functional roles, some proteins of the ∼20S complex may be exchanged for other components during the transition to the ∼35–40S particle resulting in different architectures of both complexes. In any of these models, various intermediates can be discerned, each of which represents independent challenges for biochemical studies and structural biology. The structural analysis of such diverse assemblies is currently a major task for electron microscopy (EM), especially as the existing purification protocols can only incompletely separate mixtures of different functional states. Here, we have purified functional RNA editing complexes by a multistep purification approach. Endogenous complexes were analysed by biochemical, functional and structural means. We describe the structural relationship between the ∼20S and the ∼35–40S complexes and suggest how the ∼20S complex may be integrated into the larger complex, thereby providing a picture of the architecture of the RNA editing machinery. By variability analysis, we present a comprehensive view of the endogenous RNA editing machinery in trypanosomes at different assembly stages. Results Purification of native RNA editing complexes To isolate RNA editing complexes at native conditions, we used the tandem-affinity purification (TAP) protocol (Rigaut et al, 1999). To this end, we generated a Trypanosoma brucei cell line (29-13-MP42/TAP) that expresses a C-terminally TAP-tagged version of TbMP42. TbMP42 is a 42-kDa protein that was identified as an integral component of editosomes (Panigrahi et al, 2001b; Brecht et al, 2005; Niemann et al, 2008). Expression of TbMP42/TAP in the 29-13-MP42/TAP strain relies on the tetracycline (tet) repressor/operator system and, thus, can be conditionally regulated (Wirtz et al, 1999). Tet-induced insect stage 29-13-MP42/TAP trypanosomes showed a cell-doubling time identical to the parental 29-13 strain and were morphologically indistinguishable from wild-type cells (data not shown). Expression of TbMP42/TAP was verified by Western blotting (Figure 1A) and RNA editing complexes of tet-induced 29-13-MP42/TAP trypanosomes were enriched from non-ionic detergent lysates of mitochondrial vesicles isolated at isotonic conditions (Göringer et al, 1994; Hauser et al, 1996). Figure 1.Biochemical characterisation of endogenous TAP-tagged RNA editing complexes. (A) Western blot verification of the expression of the 64.3 kDa TAP-tagged version of TbMP42 (arrow). The whole cell protein extract was prepared from 29-13-MP42/TAP insect stage trypanosomes cultivated in the absence (−) or presence (+) of tet for 74 h. (B) Silver-stained protein pattern (2 μg each) of a mitochondrial lysate (MiL), an IgG column eluate (IgG) and a calmodulin column(CaM) eluate. Proteins were identified by mass spectrometry. Contaminants (β-tubulin, TEV protease) are marked by *. (C) Identification of the RNA editing ligases TbMP52 and TbMP48 through auto-adenylation in the presence of α-[32P]-ATP. (D) CaM eluates were radioactively labelled by tyrosin iodination (125I) and fractionated in linear 10–40% (v/v) glycerol gradients. The ∼20S (green) and ∼35–40S (blue) peak fractions were analysed for the presence of RNA by radioactive postlabelling methods and spotted onto PEI cellulose plates. The presence of gRNAs was identified by labelling with guanylyl transferase and α-[32P]-GTP. (E) RNA editing in vitro activity (Ins, insertion; Del, deletion) of TAP-tagged and glycerol gradient-purified ∼20S and ∼35–40S editing complexes. The electrophoretic mobilities of editing products (arrows), ligation products and of various reaction intermediates are given on the right. '*' indicates the position of the radioactive label. Download figure Download PowerPoint Protein composition of native RNA editing complexes The electrophoretic analysis showed a decrease in the complexity of the protein pattern from the mitochondrial lysate (MiL) to the calmodulin column (CaM) eluate, ultimately identifying 14 bands ranging in size from 18 to 100 kDa (Figure 1B). Mass spectrometry identified 12 of them as previously characterised editing proteins (TbMP100, TbMP99, TbMP90, TbMP67, TbMP63, TbMP61, TbMP57, TbMP52, TbMP46, TbMP44, TbMP42/TAP, TbMP24; nomenclature of Panigrahi et al, 2001a; Supplementary Tables 1 and 2). Two proteins, identified as β-tubulin and TEV protease, were considered contaminants. Of the two RNA editing ligases (TbMP48, TbMP52), only TbMP52 was identified by mass spectrometry. However, the presence of TbMP48 was verified by self-adenylation (Sabatini and Hajduk, 1995) in the presence of α-[32P]-ATP (Figure 1C). Purified RNA editing complexes sediment as ∼20S and ∼35–40S particles CaM eluates were further analysed by isokinetic ultracentrifugation in glycerol gradients using non-radioactive material and 125I labelled eluates to increase the detection limit (Figure 1D). In line with previously published studies (Pollard et al, 1992; Corell et al, 1996), the data show the presence of two complexes with apparent Svedberg (S) values of ∼20S and ∼35–40S. Using radioactive 5′ and 3′ post-labelling methods, we confirmed the presence of endogenous RNA, including gRNA, for the ∼35–40S complexes (Pollard et al, 1992; Corell et al, 1996), whereas ∼20S complexes did not contain detectable amounts of pre-bound RNA (Rusché et al, 1997). To test whether the two complexes were functionally active, we performed U-deletion and U-insertion RNA editing in vitro assays (Igo et al, 2000, 2002). The ∼20S complexes were fully competent to correctly edit synthetic pre-edited mRNAs in a gRNA-dependent fashion (Figure 1E). By contrast, ∼35–40S complexes only showed background activity (⩽2%) likely because the RNA-binding site is occupied with endogenous RNA. Electron microscopy of the ∼35–40S RNA editing complex Using the TAP protocol (Rigaut et al, 1999) in combination with the GraFix method (Kastner et al, 2008), we purified ∼35–40S complexes for EM. The raw EM images of the ∼35–40S complexes displayed a monodisperse population of an asymmetric particle of up to ∼26 nm in size (Figure 2A). Imaged complexes revealed a compact globular shape with distinct structural features, suggesting that intact complexes had been purified. Figure 2.Electron microscopy of the ∼35–40S RNA editing complex. (A) Raw RT EM image of the ∼35–40S complex. (B) 2D class averages displaying a particle comprising an elongated platform density (oriented to the right, upper panel and to the left, lower panel) packed against a semispherical element (oriented to the left, upper panel and to the right, lower panel). (C) Structural landmarks of the complex. (D) Consensus structure of the ∼35–40S complex. Characteristic structural features are labelled. (E) Euler angle plot of the 3D RCTs included in the consensus map coloured according to weight as shown on the right. Download figure Download PowerPoint Two-dimensional (2D) class averages revealed a characteristic distribution of densities (Figure 2B and C). Typical class averages displayed a particle composed of an elongated, straight to slightly convex platform density (oriented to the right in Figure 2B, upper row and to the left in the lower row) packed against a semispherical element (oriented to the left and right side in the upper and lower row). On both ends, the platform extends into a small head-like element oriented to the top and a larger foot-like density extending to the bottom of the structure. The majority of visualised particles adopted this morphology in several independent purifications. A small number of particles, however, differed in being composed of the platform density only suggesting that these particles may differ in composition. Consensus structure of the ∼35–40S RNA editing complex On the basis of 2D class averages, it is often difficult to distinguish whether given views belong to a common three-dimensional (3D) structure or represent broken or contaminating particles. We therefore applied a 3D averaging approach that combines the random conical tilt (RCT) (Radermacher, 1988) method with weighted averaging (Sigworth, 1998). This approach can be performed free of user-bias with heterogeneous data sets and can be expanded to a detailed variability analysis. The obtained consensus structure displayed all structural characteristics described in the 2D class averages and visualised in the raw images (Figure 2D). Specifically, an elongated platform is seen to which small head-like and foot-like elements are attached on opposite sides. A small protrusion is visible in the lower part of the complex. A semispherical back is packed against the platform and forms a tight density network to it. The interface between both elements is marked by incisions in the upper and lower part. Notably, the semispherical back is asymmetric in its appearance: on one side a large protruding 'shoulder' density is seen, whereas on the opposite side a smaller inclined element is visible. A plot shows the presence of a wide range of Euler angles (Figure 2E). Variability analysis of the ∼35–40S RNA editing complex The EM analysis suggested structural heterogeneity of the ∼35–40S complex, as some particles appeared to be composed of the platform density only. As the purified particles represent endogenous RNA editing complexes enriched at steady state conditions, these complexes may differ not only in the assembly stage, but also in their RNA and protein content. To address this aspect, we performed a 3D multivariate statistical analysis (MSA) (Liu et al, 2004) on the aligned 3D RCTs (Sander et al, 2006). Variability analysis revealed at least six morphologically different 3D subtypes (Figure 3A). Four of these subtypes (rows II–V) showed all the structural characteristics seen in the consensus structure (compare Figures 3A and 2D, first view). However, these subtypes differed considerably in width of the semispherical back (indicated as dashed line), ranging from small (row II) to extensive (row V). Notably, the first two structures shown in row III are very similar to the consensus structure. In contrast, structures shown in the row I were only composed of an elongated density that adopts different curvatures, while lacking the semispherical back. This confirms that the elongated 2D class averages described above miss this density. Furthermore, we also found a small subset of structures that did not show the typical structural features (row VI). We suggest that this population may either represent contaminating complexes or different assembly stages. Figure 3.Variability analysis of the ∼35–40S RNA editing complex. (A) 3D-MSA of individual aligned 3D RCT volumes indicated six subtypes of particle populations within the ∼35–40S sample. Landmarks are labelled in representative examples (h, head-like protuberance; p, platform; f, foot-like protuberance; the semispherical back is indicated as a dashed line). (B) Refined maps of representative 3D class averages subsequent to cMRA. The numbers of particles used to calculate the refined 3D structures are listed on the right. (C) Shape profiling of the ∼35–40S complex by colour-scale encoded overlays showing that the majority of complexes possesses a platform density packed against a semispherical back (red). (D) Histogram of the particle width suggesting at least three differently sized subpopulations. (E) cMRA of three independently purified samples revealing that the majority of particles belong to the subset shown in (A), row III (colour coding as in (A)) consistent with the consensus structure (Figure 2D). Download figure Download PowerPoint Quantification of ∼35–40S editosomal subpopulations To quantify the subpopulations, three different types of analysis were performed, (1) shape profiling (Figure 3C), (2) a width distribution analysis (Figure 3D) and (3) a competitive multireference alignment (cMRA) (Figure 3B and E). For a description of the overall architecture, all 3D class averages were colour-scale encoded according to their abundance and an overlay was created (Figure 3C). The majority of complexes (>80%) is composed of the described densities (red) in line with the consensus structure, whereas variable regions (blue) are found in the periphery of the semispherical back and in the size and position of the foot-like domain. Quantification of the particles' width—a prominent individual feature of the 3D class averages—suggested an overlay of 3–5 Gaussian distributions (Figure 3D). First, a small subpopulation of complexes possesses a width of ∼15±1 nm (green curve) and represents the elongated particles as seen in Figure 3A, row I. The largest subpopulation (rows II–IV) has a width of ∼18.5±2 nm (blue curve). The third subpopulation (purple curve) comprises particles similar to the consensus structure except for the larger width of ∼21.5±1 nm (row V). Finally, cMRA using the 3D class averages to create reference projections was performed. Thereby, each single-particle image is competitively aligned to reference projections of the 3D class averages so that the number of particles that fit best to each of the six subtypes can be depicted (Figure 3E) and refined 3D maps are obtained (Figure 3B). Significantly, we found no major differences between the three samples derived from independent editosome preparations, suggesting that the distribution of particle subtypes was preparation independent. In line with the shape profile (Figure 3C) and the distribution of the particle width (Figure 3D), we found that ∼40% of the particles adopted a conformation similar to the consensus structure (Figure 3E), whereas all other subpopulations occurred roughly equally in frequency (∼10–20%, each). Thus, the major particle subpopulation is composed of an elongated platform packed against a semispherical density, whereas the minor subpopulations lack the semispherical back or possess additional densities. Structure refinement of the ∼35–40S RNA editing complex by cryo negative staining EM The 3D map of the best-defined 3D class average (Figure 4A) was used as start-up reference to refine another data set of the ∼35–40S complex taken at liquid nitrogen temperature to a resolution of ∼12.7–19.0 nm using cMRA (Figure 4B and C). The Euler angle plot of all included particles indicated the presence of a wide variety of angular views with two more frequently found angular regions (Figure 4D). The initial consensus and the refined maps are very similar except for the higher resolution of the refined structure, resulting in more fine structural details in the refined map (Figure 4C). All described elements (compare landmarks in Figures 2D and 4C) were also visible in the refined reconstruction. The surface representation of the ∼35–40S complex is estimated to enclose a molecular mass of ∼1.45±0.15 MDa. By using the 3D map, the sedimentation coefficient can be predicted (Garcia de la Torre et al, 2001). Our calculation predicts a value of 35–41S, which is in agreement with the apparent sedimentation behaviour observed in the glycerol gradients. Figure 4.Refinement of the 3D map of the ∼35–40S RNA editing complex by cryo negative staining EM. (A) Surface representation of the best-defined 3D class average showing similar structural elements as seen in Figure 2D. (B) FSC suggested a resolution of 1.27–1.90 nm for the refined map. (C) Surface representation of the ∼35–40S RNA editing complex showing a wealth of fine structural details in the refined 3D map. The labelling of landmarks is similar to Figure 2D. (D) Euler angle plot of the refined data set. Relative particle numbers per unit area are colour coded as shown on the right. Download figure Download PowerPoint Electron microscopy of the ∼20S RNA editing complex The concentration of ∼20S particles seen in the EM was markedly higher by a factor of 5–10 as compared with the ∼35–40S complex. The raw EM images showed a monodisperse population of an elongated particle with dimensions of up to ∼21–26 nm (Figure 5A). Some of the particles appeared to adopt a straight conformation: these particles were particularly slim and elongated (yellow box). Other particles were broader and appeared to be in a more bent conformation (green circles). Figure 5.Electron microscopy of the ∼20S RNA editing complex. (A) Raw RT EM image of the ∼20–24S fractions showing an elongated particle of variable width including globular (green) and elongated (yellow) complexes. (B) 2D class averages of the ∼20S complex. (C) Euler angle plot of the ∼20S complex RCTs used to calculate the consensus map coloured according to weight as shown on the right. (D) Consensus model of the ∼20S complex displaying a bipartite shape. Landmarks are labelled. Download figure Download PowerPoint These shapes were also reflected by the 2D class averages showing a variety of different views (Figure 5B). We found amongst others triangular (views 1–4), bent (views 5–8), semicircular (views 9–12) and elongated views (views 13–16), suggesting the sample to be heterogeneous. The majority of particles showed a bipartite appearance with two approximately equally sized subdomains connected by an interface. Both subdomains, however, differed in their structural details, suggesting that the particle is not a homodimer. Consensus structure of the ∼20S RNA editing complex Similar to the procedure used for the ∼35–40S complex, a consensus structure of the ∼20S complex was determined. The Euler angle plot disclosed preferential binding to the carbon with two preferred angular regions (Figure 5C). Overall, the complex has an elongated, slightly bent appearance (Figure 5D). This results in a concave–convex shape, displaying one concave and one convex contour on opposite sides. The particle is composed of two globular domains roughly equal in size, the upper domain of which being more roundish, and the lower one being somewhat thinner. Both subdomains interact extensively in an interface where a protruding arm is seen on one side (view 1) and a triangular protrusion emerges from the opposite side (view 3). Variability analysis of the ∼20S RNA editing complex To describe the structural variability suggested by the 2D class averages, 3D-MSA was performed using aligned RCTs. We identified four structural subgroups showing distinct structural features: representative 3D class averages are shown in Figure 6A and refined 3D maps subsequent to cMRA are shown in Figure 6B. The first subgroup comprises particles that show a clear separation into two approximately equally sized subdomains connected by an interface: the two subdomains adopt variable relative positions resulting in different curvatures of the particle (row I). The second subgroup (row II) also shows a slightly bipartite shape but displays, in addition, a small semispherical back similar to the ∼35–40S particles. The third and fourth subgroups (rows III and IV) differ from the former two in being more elongated but still exhibiting a bipartite shape. The fourth subtype possesses an additional domain attached to the body in the upper right region. Figure 6.Variability analysis of the ∼20S RNA editing complex. (A) Representative 3D class averages demonstrating the overall variability of ∼20S complexes. (B) Refined models of representative 3D class averages subsequent to cMRA. The numbers of particles used to calculate the refined 3D structures are listed on the right. (C) Shape profiling of the ∼20S complex by colour-scale encoded overlays revealing that the majority of complexes are composed of two subdomains connected by a broad interface (red). (D) Histogram of the overall abundance of structural subtypes within three data sets as determined by cMRA showing that about one third of the particles were assigned to the subgroups I and II (colour coding as in (A)). Download figure Download PowerPoint Quantification of ∼20S editosomal subpopulations An overlay of 3D shapes according to their proportion in the data set provides a graphical representation of particle shapes (Figure 6C) and also visualises variable regions. Notably, the majority of complexes has an elongated, slightly curved shape, whereas in some particles, additional densities are attached to the upper region. Using cMRA, we quantified the abundance of the 3D subtypes for three data sets. Indeed, we found that the relative proportions of the 3D subtypes were similar in these data sets, suggesting that the distribution of particles belonging to the different subtypes is representative (Figure 6D)