Structural basis for pan-coronavirus inhibition of 3CL protease.

The severe acute respiratory syndrome (SARS) coronavirus (CoV) main protease represents an attractive target for the development of novel anti-SARS agents. The tertiary structure of the protease consists of two distinct folds. One is the N-terminal chymotrypsin-like fold that consists of two structural domains and constitutes the catalytic machinery; the other is the C-terminal helical domain, which has an unclear function and is not found in other RNA virus main proteases. To understand the functional roles of the two structural parts of the SARS-CoV main protease, we generated the full-length of this enzyme as well as several terminally truncated forms, different from each other only by the number of amino acid residues at the C- or N-terminal regions. The quaternary structure and Kd value of the protease were analyzed by analytical ultracentrifugation. The results showed that the N-terminal 1–3 amino acid-truncated protease maintains 76% of enzyme activity and that the major form is a dimer, as in the wild type. However, the amino acids 1–4-truncated protease showed the major form to be a monomer and had little enzyme activity. As a result, the fourth amino acid seemed to have a powerful effect on the quaternary structure and activity of this protease. The last C-terminal helically truncated protease also exhibited a greater tendency to form monomer and showed little activity. We concluded that both the C- and the N-terminal regions influence the dimerization and enzyme activity of the SARS-CoV main protease.

Article Figures and data Abstract Editor's evaluation Introduction Results Discussion Materials and methods Data availability References Decision letter Author response Article and author information Metrics Abstract Paclitaxel (Taxol) is a taxane and a chemotherapeutic drug that stabilizes microtubules. While the interaction of paclitaxel with microtubules is well described, the lack of high-resolution structural information on a tubulin-taxane complex precludes a comprehensive description of the binding determinants that affect its mechanism of action. Here, we solved the crystal structure of baccatin III the core moiety of paclitaxel-tubulin complex at 1.9 Å resolution. Based on this information, we engineered taxanes with modified C13 side chains, solved their crystal structures in complex with tubulin, and analyzed their effects on microtubules (X-ray fiber diffraction), along with those of paclitaxel, docetaxel, and baccatin III. Further comparison of high-resolution structures and microtubules’ diffractions with the apo forms and molecular dynamics approaches allowed us to understand the consequences of taxane binding to tubulin in solution and under assembled conditions. The results sheds light on three main mechanistic questions: (1) taxanes bind better to microtubules than to tubulin because tubulin assembly is linked to a βM-loopconformational reorganization (otherwise occludes the access to the taxane site) and, bulky C13 side chains preferentially recognize the assembled conformational state; (2) the occupancy of the taxane site has no influence on the straightness of tubulin protofilaments and; (3) longitudinal expansion of the microtubule lattices arises from the accommodation of the taxane core within the site, a process that is no related to the microtubule stabilization (baccatin III is biochemically inactive). In conclusion, our combined experimental and computational approach allowed us to describe the tubulin-taxane interaction in atomic detail and assess the structural determinants for binding. Editor's evaluation Here Prota et al. compare the action of the microtubule-stabilizing agent, taxol, with that of closely-related analogues and, as a result, successfully dissect the interactions and roles of different regions of the taxol molecule. The overall story is solid, providing new molecular insights, including defining and separating the lattice expansion effect from the lattice stabilization effect upon taxane binding. This important work will be of interest to the microtubule cytoskeleton and structural biology communities. https://doi.org/10.7554/eLife.84791.sa0 Decision letter eLife's review process Introduction The taxane paclitaxel is a drug included in the World Health Organization’s List of Essential Medicines (World Health Organization, 2021). Taxanes, either alone or in combination with other chemotherapeutic agents, are important drugs for the treatment of several solid tumors, such as ovarian, lung, and breast cancer, as well as advanced Kaposi’s sarcoma (Ettinger, 1993; Arbuck et al., 1994; Saville et al., 1995; Lindemann et al., 2012). The three taxanes in clinical use, paclitaxel (Taxol), docetaxel (Taxotere), and cabazitaxel (Jevtana), are part of a large family of chemically diverse compounds that bind to the so-called ‘taxane site’ of the αβ-tubulin heterodimer (Field et al., 2013; Steinmetz and Prota, 2018; Figure 1A and B), the building block of microtubules. However, the appearance of peripheral sensory neuropathy and other side effects caused by taxanes compromises treatment efficacy in the long term (Gornstein and Schwarz, 2014). Thus, understanding the underlying mechanism of microtubule stabilization by this class of antitubulin agents is an important requirement for future and safer drug development efforts. Figure 1 Download asset Open asset Structures of tubulin and ligands employed in the work. (A) Tubulin heterodimer (α-tubulin in gray and β-tubulin in white) in ribbon representation, where nucleotide binding sites have been highlighted in sphere representation (B) Structural features of the tubulin β-subunit. (C) Structures of taxanes used in this study. Because taxane-site ligands stabilize microtubules and suppress their dynamics, they are collectively called microtubule-stabilizing agents. Several structures of microtubules in complex with taxane-site agents have been recently analyzed and solved by cryo-electron microscopy (cryo-EM) to resolutions ranging between ~3 and ~10 Å. For paclitaxel, it was initially suggested that the drug acts on longitudinal tubulin contacts along protofilaments in microtubules by allosterically expanding the microtubule lattice in the direction of its long filament axis (Vale et al., 1994; Arnal and Wade, 1995; Alushin et al., 2014), a notion that is also consistent with X-ray fiber diffraction data (Estévez-Gallego et al., 2020). However, more recent studies suggest that paclitaxel enhances lattice flexibility and acts on lateral tubulin contacts between protofilaments in microtubules through interactions with the M-loop of the β-tubulin subunit (βM loop) (Kellogg et al., 2017; Debs et al., 2020; Manka and Moores, 2018). Besides directly acting on microtubules, taxane-site ligands also have the capacity to bind to unassembled tubulin dimers and promote their assembly into microtubules (Schiff and Horwitz, 1981; Carlier and Pantaloni, 1983; Howard and Timasheff, 1988; Díaz et al., 1993; Buey et al., 2004). Several structures of non-taxane agents bound to the taxane site of tubulin have been solved to resolutions ranging from 2.4 to 1.8Å by X-ray crystallography (Prota et al., 2013a; Trigili et al., 2016; Prota et al., 2017; Balaguer et al., 2019). These data suggested that one mode of action of some taxane-site ligands such as zampanolide (PDB ID 4I4T) or epothilone A (PDB ID 4I5O) on unassembled tubulin is to stabilize lateral tubulin contacts between protofilaments within microtubules by structuring and stabilizing the βM loop into a short α-helix (Prota et al., 2013a). In contrast, the absence of a helical structure for this segment in the presence of the taxane-site ligands dictyostatin (PDB ID 4MF4) and discodermolide (PDB ID 5LXT) (Trigili et al., 2016; Prota et al., 2017) suggests a different, still poorly understood mechanism of microtubule stabilization for these two classes of non-taxane agents. In the case of taxanes, one hypothesis is that they preferentially bind to a specific conformation of tubulin. It is well established that tubulin displays two prominent conformations that are related to its assembly state (reviewed in Knossow et al., 2020): a ‘straight’ conformation present in assembled microtubules (denoted ‘straight tubulin’ hereafter) and a ‘curved’ conformation observed in unassembled tubulin (denoted ‘curved tubulin’ hereafter). The ‘curved-to-straight’ conformational transition is required for the formation of lateral tubulin contacts between protofilaments in the main shaft of microtubules. Some data suggest that the activation mechanism of taxanes facilitates the curved-to-straight conformational transition by preferentially binding to the straight conformation of tubulin (Nogales et al., 1998; Elie-Caille et al., 2007; Benoit et al., 2018). Structural information of a taxane in complex with unassembled tubulin is currently unavailable. With the aim of providing insight into the mechanism of action of this important class of anticancer drugs and into the tubulin-taxane interaction, we solved the high-resolution structures of three different taxanes bound to curved tubulin by X-ray crystallography. We further analyzed the effects of different taxanes on the microtubule lattice by X-ray fiber diffraction. These studies were complemented with molecular dynamics (MD) simulations that shed light on issues that were not amenable to experimental verification. Taken together, our results suggest that the main reason for the differential affinity of taxane-site ligands for assembled tubulin and unassembled tubulin arises from two terms. First, the stabilization of the βM loop in an ‘out’ conformation compatible with the formation of specific lateral contacts in microtubules and second, the selectivity of the bulky C13 side chain for the assembled, straight conformational state of tubulin. Finally, we found that the occupancy of the taxane site results in a displacement of the S9-S10 loop of β-tubulin that accounts for the observed microtubule expansion with no influence, however, on the straightness of tubulin protofilaments. Results High-resolution crystal structure of a tubulin-taxane complex To determine the high-resolution structure of a taxane bound to curved tubulin, we performed both soaking and co-crystallization experiments using the previously described protein complexes termed T2R-TTL and TD1. The former complex is composed of two αβ-tubulin heterodimers bound head-to-tail, the stathmin-like protein RB3, and the tubulin tyrosine ligase (PDB ID 4IIJ) (Prota et al., 2013a; Prota et al., 2013b); the latter complex contains one αβ-tubulin heterodimer and the DARP in D1 (PDB ID 4DRX) (Pecqueur et al., 2012). We did not succeed in procuring any valuable structural information from these two crystal ensembles using a first series of taxanes comprising paclitaxel, docetaxel, the more soluble 3’-N-m-aminobenzamido-3’-N-debenzamidopaclitaxel (N-AB-PT) (Li et al., 2000), and the engineered, high-affinity taxanes Chitax 40 (Matesanz et al., 2008) and Chitax 68 (Ma et al., 2018). We thus decided to approach the issue from a different angle and started off with baccatin III, a precursor in the biosynthesis of paclitaxel that contains both the C2-benzoyloxy ring C and the C10 acetate ester, but lacks the C13 side chain with both the 3’-N-benzamido phenyl ring A and the 3’-phenyl ring B moieties (Samaranayake et al., 1993; Figure 1C). Notably, baccatin III is largely biologically inactive despite displaying micromolar affinity for microtubules (Parness et al., 1982; Lataste et al., 1984; Kingston, 2000; Andreu and Barasoain, 2001). We found that baccatin III shows detectable affinity (Kb 25°C 3.0±0.5 × 103 M–1) to unassembled tubulin, which is in the same range as for other compounds that have been co-crystallized with tubulin, such as epothilone A 8±3 × 103 M–1 (Canales et al., 2014) and discodermolide 2.0±0.7 × 104 M–1 (Canales et al., 2011). Therefore, we hypothesized that the presence of the C13 side chain of the aforementioned taxanes might preclude the binding to the curved tubulin form present in both the T2R-TTL and the TD1 complexes. Subsequently, we succeeded in obtaining a T2R-TTL-baccatin III complex structure that was solved at 1.9 Å resolution (PDB ID 8BDE) (Figure 2A and D; Table 1). We found that the ligand binds to the taxane site of curved tubulin with its C2-benzoyloxy ring C stacked between the side chains of βH229 and βL275 in the leucine-rich β-tubulin pocket lined by the side chains of βC213, βL217, βL219, βD226, βH229, βL230, and βL275 (Figures 3A and 4A). Its carbonyl oxygen forms a weak hydrogen bond to the main chain amide of βR278. The C10 acetate is exposed to the solvent and, together with the C12 methyl, is within van der Waals distance to βG370 of the βS9-βS10 loop. Furthermore, the oxetane oxygen and the C13 hydroxyl accept hydrogen bonds from the main chain amide nitrogen of βT276 and the βH229 imidazole NE2, respectively. The C4 acetate is buried in the hydrophobic pocket made up by βL230, βA233, βF272, βP274, βL275, βM302, βL371, and the aliphatic portion of the βR369 side chain. Figure 2 Download asset Open asset T2R-TTL structures in complex with baccatin III, 2a, and 2b. Overall view of the T2R-TTL-baccatin III (PDB ID 8BDE) (A), the T2R-TTL-2a (PDB ID 8BDF) (B), and the T2R-TTL-2b (PDB ID 8BDG) crystal structures. The α- and β-tubulin chains are colored in dark and light gray, respectively. The TTL chains (cyan) and the RB3 (yellow-orange) are shown in ribbon representation. The tubulin-bound ligands are displayed as spheres and are colored following the same color scheme as in the main figures. (D–F) Electron-density maps highlighting the bound baccatin III, 2a, and 2b. The SigmaA-weighted 2mFo − DFc (dark blue mesh) and mFo − DFc (light green mesh) omit maps are contoured at +1.0σ and +3.0σ, respectively. The map calculations excluded the atoms of the corresponding ligands. (G) Anomalous density peaks detected in both the binding sites in chains B and D of T2R-TTL for the bromine moiety of compound 2b. Figure 3 Download asset Open asset Crystal structure of T2R-TTL-baccatin III (PDB ID 8BDE) and T2R-TTL-2a (PDB ID 8BDF) complexes. (A) Close-up view of the interaction network observed between baccatin III (lemon) and β-tubulin (light gray). Interacting residues of tubulin are shown in stick representation and are labeled. Oxygen and nitrogen atoms are colored red and blue, respectively; carbon atoms are in lemon (baccatin III) or light gray (tubulin). Hydrogen bonds are depicted as black dashed lines. Secondary structural elements of tubulin are labeled in blue. (B) Close-up view of the interaction of 2a (violet) with β-tubulin in the same view and representation as in (A). (C) The same close-up view as in (A) and (B) with the superimposed baccatin III (lemon) and 2a (violet) complex structures. Water molecules belonging to the baccatin III structure are represented as lemon spheres. Figure 4 Download asset Open asset Comparison of taxane binding to unassembled curved versus assembled straight tubulin. (A) Close-up view of the superimposed baccatin III bound (ligand in lemon; protein in gray ribbon and sticks) to curved tubulin (PDB ID 8BDE) and paclitaxel bound to straight tubulin as found in a microtubule (PDB ID 6WVR; ligand in dark green; protein in slate ribbon and sticks) structures. Interacting residues of tubulin are shown in stick representation and are labeled. Oxygen and nitrogen atoms are colored red and blue, respectively. Hydrogen bonds are depicted as black dashed lines. Secondary structural elements of tubulin are labeled in blue. Water molecules belonging to the baccatin III structure are represented as lemon spheres. The structures were superimposed onto their taxane sites (residues 208–219+225–237+272–276+286–296+318–320+359–376); root-mean-square deviations (rmsd) 0.894 Å (52 Cα atoms). (B) Close-up view of superimposed 2a bound to curved tubulin (PDB ID 8BDF) (ligand in violet; protein in gray ribbon and sticks) and paclitaxel bound to straight tubulin (PDB ID 6WVR; ligand in dark green; protein in slate ribbon and sticks) structures (rmsd 0.826 Å over 52 Cα atoms) using the same settings as in (A). (C) Conformational changes on β-tubulin induced by paclitaxel upon binding to straight tubulin in microtubules (PDB ID 6WVR). The α-tubulin and β-tubulin chains are in ribbon representation and are colored in dark and light gray, respectively. The rmsd differences between unbound and paclitaxel-bound straight tubulin are represented as dark (backbone rmsd) blue spheres. Only the rmsd differences above a threshold of average ± standard deviation are displayed. The sphere radii correspond to the average-subtracted rmsd values displayed in panel (D). (D) Rmsd plots of backbone positions between the paclitaxel bound (PDB ID 6WVR) and the apo (PDB ID 6DPV) straight tubulin in microtubules. The gray error bar represents the average rmsd ± standard deviation. The top bar is colored according to the following domain assignment: N-terminal domain (N-domain., marine blue), intermediate domain (I-domain, orange), central helix βH7 (lemon), and C-terminal domain (C-domain, red). The β-tubulin chains of the corresponding structures were superimposed onto their β-tubulin N-terminal β-sheets (rmsd 0.304 Å over 30 Cα). Table 1 X-ray data collection and refinement statistics. T2R-TTL-BacIIIT2R-TTL-2aT2R-TTL-2bData collectionSpace groupP212121P212121P212121Cell dimensionsa, b, c (Å)104.1, 157.2, 179.2104.8, 157.9, 179.1105.3, 158.6, 179.2Resolution (Å)49.2–1.9 (1.95–1.90)49.3–1.95 (2.00–1.95)49.4–2.35 (2.41–2.35)Rmerge(%)10.7 (491.9)13.3 (516.6)17.4 (403.5)Rmeas (%)11.1 (513.1)13.6 (526.1)17.7 (410.8)Rpim (%)3.3 (147.5)2.9 (102.9)2.6 (57.7)I/σI16.5 (0.5)20.1 (0.7)20.1 (0.9)CC half100 (17.8)100 (31.4)99.9 (46.6)Completeness (%)100 (99.8)100 (100)100 (100)Redundancy13.5 (12.4)27.3 (27.8)28.5 (28.3)RefinementResolution (Å)49.2–1.949.3–1.9549.4–2.35No. unique reflections229654215774125168Rwork/Rfree19.2/21.818.9/21.618.3/21.4No. atomsProtein175551738917227Ligand42120Water861883166Average B-factors (Å2)Protein59.062.976.1Ligand (chain B/D)n.a. / 109.2111.4/102.8146.6/144.9Water56.260.359.4Wilson B-factor41.743.156.9R.m.s. deviationsBond lengths (Å)0.0030.0030.002Bond angles (°)0.6420.6550.550Ramachandran statisticsFavored regions (%)98.198.198.0Allowed regions (%)1.81.82.0Outliers (%)0.10.10 For each structure, data were collected from a single crystal. Values in parentheses are for highest-resolution shell. Generation of paclitaxel analogs that bind to tubulin crystals Aiming to understand the implication on tubulin activation of the paclitaxel’s bulky and hydrophobic C13 ring A moiety (or its equivalent tert-butyl in docetaxel) and to elucidate the reason why it apparently precludes binding to T2R-TTL and TD1 crystals (see above), we devoted a synthetic effort to obtaining new taxane ligands with modified C13 side chains. We produced a series of modified taxanes bearing smaller groups than paclitaxel at the 3’-N position, namely, acrylamide 2a, haloacetamides 2b, and 2c, and isothiocyanate 2d (Figure 1C). We could measure binding of 2a to unassembled tubulin dimers (Kb25°C 0.8±0.3 × 103 M–1), but not of N-AB-PT (Li et al., 2000), Chitax 40 (Matesanz et al., 2008), or Chitax 68 (Ma et al., 2018), thus indicating that the modification of the paclitaxel structure increased the binding affinity for unassembled tubulin. In fact (Figure 2B, C, E and F), we found unequivocal difference electron densities at the taxane site of β-tubulin in T2R-TTL crystals soaked with 2a (PDB ID 8BDF) and 2b (PDB ID 8BDG) and refined the corresponding structures to 1.95 and 2.35 Å resolution, respectively (Table 1). Interestingly, the electron densities of compounds 2a and 2b displayed a continuity between the 3’-N-attached moieties of both ligands and the side chain of His 229 of β-tubulin (βH229), suggesting the possible formation of a covalent adduct. For further validation, we collected additional X-ray diffraction data on T2R-TTL crystals soaked with the haloacetamide derivative 2b at the bromine peak wavelength of 0.91501 Å. After rigid body and restrained refinement, we detected two clear anomalous difference peaks in electron densities at the taxane sites of the two tubulin dimers in the T2R-TTL crystals soaked with 2b (Figure which did not covalent bond Furthermore, refinement performed in with 2a in both the covalent and the in clear electron density for the red difference peaks for the covalent form were present refinement we the electron density observed in the T2R-TTL-2a structure as a hydrogen bond between the βH229 and the carbonyl of the ligand side chain than a covalent bond (Figure The T2R-TTL-2a complex structure that 2a in interactions to curved tubulin by of both its C2-benzoyloxy ring C and its oxetane as found for baccatin III (Figure 3A and However, the core ring of 2a is helix and by between the two of Å for core a that is to that observed for paclitaxel bound to straight tubulin in microtubules (PDB ID 6WVR; of Å for core of Å for core Figure to paclitaxel bound to straight tubulin, the carbonyl of the moiety of 2a forms a hydrogen bond to the βH229 in curved tubulin (Figures and The moiety of 2a is exposed to the solvent and it forms hydrogen bonds to the side chains of and βR369 of it within a that is by molecules in the curved III This hydrogen bond network be established by the 3’-N-benzamido phenyl ring A of paclitaxel in the curved tubulin both the molecules and the hydrophobic of the and side chains on helix are for to to stabilize the ring In the of paclitaxel-bound microtubules the helix helix these three side chains to form a hydrophobic that stabilizes the A which suggest a structural mechanism for the affinity of paclitaxel observed for the straight tubulin the helix the side chain of to the of the βR369 side which a This additional stabilization through a interaction to the amide nitrogen of paclitaxel and a more binding of paclitaxel to microtubules (Figures and The absence of the C10 acetate in 2a to baccatin III has on the conformation of the structural elements that the taxane site (Figure these structural data for the first a high-resolution structural description of the interaction of taxanes a C13 side chain with curved tubulin. that the main interaction of this class of antitubulin agents is by their baccatin III core further that the taxane in both curved and straight tubulin is however, structural why paclitaxel binds more to straight tubulin. The of these structural determinants the development of taxanes to better their mechanism of a new to side our results suggest that the structure is an to the interaction of paclitaxel with curved tubulin at resolution and that X-ray crystallography is a valuable to the molecular mechanism of action of microtubule-stabilizing agents binding to the taxane Conformational changes upon taxane binding to curved and straight tubulin we the conformational changes induced by binding of baccatin III and 2a to curved tubulin. To this we first superimposed the crystal structures of apo tubulin (PDB ID III (PDB ID and (PDB ID 8BDF) onto the N-terminal β-sheets of β-tubulin (residues and and the root-mean-square deviations (rmsd) between the apo and the two Å of Å of Cα). These rmsd values were also and onto the corresponding structures to the regions of conformational shown in Figure and conformational changes were observed for backbone atoms of the loop and the N-terminal segment of the βM loop in both the III and complex structures. Interestingly, the loop that is in longitudinal tubulin contacts along protofilaments is in the ‘out’ conformation in both structures et al., 2011). This an between the taxane site and the loop the central helix βH7 and the nucleotide bound to In the case of the βM we found electron densities for its N-terminal up to the portion of the loop in both complex structures. This βM loop structuring has been observed previously in tubulin complexes with the taxane-site ligands dictyostatin and discodermolide (Trigili et al., 2016; Prota et al., 2017; that the taxane-site ligands zampanolide and epothilone A promote the structuring of the βM loop into a helical conformation Prota et al., 2013a). A effect of taxanes on the βM loop is consistent with the notion that paclitaxel stabilizes this structural in two conformations to lateral contacts et al., 2020). We also found conformational changes in the βS9-βS10 which were more prominent in than in III. This be by the presence of a C13 side chain in 2a that more for accommodation the taxane site to baccatin III, which lacks a C13 side chain. Finally, we observed a conformational of the helix in the III structure, which was in Figure Download asset Open asset Conformational changes induced by taxane binding to curved tubulin. (A) Conformational changes on the backbone atoms (dark of the β-tubulin chain induced by baccatin III upon binding to curved tubulin. The tubulin chains are in ribbon representation and are colored in dark and light gray, respectively. The root-mean-square deviation (rmsd) values of the superimposed unbound and baccatin curved tubulin are represented as dark blue (backbone rmsd) respectively. Only the rmsd values above a threshold of average standard deviation are displayed. The sphere radii correspond to the average-subtracted rmsd values displayed in panel (B) Rmsd plots of the backbone positions between the baccatin bound (PDB ID 8BDE) and the apo (PDB ID curved tubulin The gray error bar represents the average rmsd ± standard deviation. The top bar is colored according to the following domain assignment: N-terminal domain marine blue), intermediate domain (I-domain, orange), central helix (lemon), C-terminal domain (C-domain, red). The β-tubulin chains of the corresponding structures were superimposed onto their β-tubulin N-terminal (rmsd Å over Cα). (C) Conformational changes on the backbone atoms (dark of the β-tubulin chain induced by 2a upon binding to curved tubulin. (D) Rmsd plots of the backbone positions between the 2a bound (PDB ID 8BDF) and the apo (PDB ID curved tubulin state (rmsd Å over Cα). The same settings as in (B) are To the effect of the observed conformational changes on the domain in β-tubulin of the we further superimposed the β-tubulin chains of apo tubulin, III, and onto their central βH7 (residues For III, a between the N-terminal and the intermediate was observed (Figure 1 and binding of 2a caused both the N-terminal and intermediate of β-tubulin to (Figure 3 and Thus, taxane binding to tubulin but conformational Figure Download asset Open asset representation of domain observed from apo to baccatin to curved tubulin. The three structures were superimposed onto their central βH7 to better the domain to each The are colored according to their domain and their are contoured using the same color N-terminal domain marine blue), intermediate domain (I-domain, orange), central helix βH7 (lemon), C-terminal domain (C-domain, red). The of the are highlighted with black 1 Download asset This be in because still the for Download as Download as Download as Conformational transition from apo to baccatin unassembled tubulin view on β-tubulin the in the of a 2 Download asset This be in because still the for Download as Download as Download as Conformational transition from apo to baccatin unassembled tubulin view on β-tubulin from the in the of a 3 Download asset This be in because still the for Download as Download as Download as Conformational transition from apo to unassembled tubulin view on β-tubulin the in the of a 4 Download asset This be in because still the for

Decision letter: Structural insight into the stabilization of microtubules by taxanes

The infectious SARS-CoV-2 causes COVID-19, which is now a global pandemic. Aiming for effective treatments, we focused on the key drug target, the viral 3C-like (3CL) protease. We modeled a big dataset with 42 SARS-CoV-2 3CL protease-ligand complex structures from ∼98.7% similar SARS-CoV 3CL protease with abundant complex structures. The diverse flexible active site conformations identified in the dataset were clustered into six protease pharmacophore clusters (PPCs). For the PPCs with distinct flexible protease active sites and diverse interaction environments, we identified pharmacophore anchor hotspots. A total of 11 "PPC consensus anchors" (a distinct set observed in each PPC) were observed, of which three "PPC core anchors" EHV2, HV1, and V3 are strongly conserved across PPCs. The six PPC cavities were then applied in virtual screening of 2122 FDA drugs for repurposing, using core anchor-derived "PPC scoring S" to yield seven drug candidates. Experimental testing by SARS-CoV-2 3CL protease inhibition assay and antiviral cytopathic effect assays discovered active hits, Boceprevir and Telaprevir (HCV drugs) and Nelfinavir (HIV drug). Specifically, Boceprevir showed strong protease inhibition with micromolar IC50 of 1.42 μM and an antiviral activity with EC50 of 49.89 μM, whereas Telaprevir showed moderate protease inhibition only with an IC50 of 11.47 μM. Nelfinavir solely showed antiviral activity with a micromolar EC50 value of 3.28 μM. Analysis of binding mechanisms of protease inhibitors revealed the role of PPC core anchors. Our PPCs revealed the flexible protease active site conformations, which successfully enabled drug repurposing.

L-arginine deiminase (ADI) catalyzes the irreversible hydrolysis of L-arginine to citrulline and ammonia. In a previous report of the structure of apoADI from Pseudomonas aeruginosa, the four residues that form the catalytic motif were identified as Cys406, His278, Asp280, and Asp166. The function of Cys406 in nucleophilic catalysis has been demonstrated by transient kinetic studies. In this study, the structure of the C406A mutant in complex with L-arginine is reported to provide a snapshot of the enzyme.substrate complex. Through the comparison of the structures of apoenzyme and substrate-bound enzyme, a substrate-induced conformational transition, which might play an important role in activity regulation, was discovered. Furthermore, the position of the guanidinium group of the bound substrate relative to the side chains of His278, Asp280, and Asp166 indicated that these residues mediate multiple proton transfers. His278 and Asp280, which are positioned to activate the water nucleophile in the hydrolysis of the S-alkylthiouronium intermediate, were replaced with alanine to stabilize the intermediate for structure determination. The structures determined for the H278A and D280A mutants co-crystallized with L-arginine provide a snapshot of the S-alkylthiouronium adduct formed by attack of Cys406 on the guanidinium carbon of L-arginine followed by the elimination of ammonia. Asp280 and Asp166 engage in ionic interactions with the guanidinium group in the C406A ADI. L-arginine structure and might orient the reaction center and participate in proton transfer. Structure determination of D166A revealed the apoD166A ADI. The collection of structures is interpreted in the context of recent biochemical data to propose a model for ADI substrate recognition and catalysis.

Article Figures and data Abstract Editor's evaluation eLife digest Introduction Results Discussion Materials and methods Data availability References Decision letter Author response Article and author information Metrics Abstract Actin isoforms organize into distinct networks that are essential for the normal function of eukaryotic cells. Despite a high level of sequence and structure conservation, subtle differences in their design principles determine the interaction with myosin motors and actin-binding proteins. Therefore, identifying how the structure of actin isoforms relates to function is important for our understanding of normal cytoskeletal physiology. Here, we report the high-resolution structures of filamentous skeletal muscle α-actin (3.37 Å), cardiac muscle α-actin (3.07 Å), ß-actin (2.99 Å), and γ-actin (3.38 Å) in the Mg2+·ADP state with their native post-translational modifications. The structures revealed isoform-specific conformations of the N-terminus that shift closer to the filament surface upon myosin binding, thereby establishing isoform-specific interfaces. Collectively, the structures of single-isotype, post-translationally modified bare skeletal muscle α-actin, cardiac muscle α-actin, ß-actin, and γ-actin reveal general principles, similarities, and differences between isoforms. They complement the repertoire of known actin structures and allow for a comprehensive understanding of in vitro and in vivo functions of actin isoforms. Editor's evaluation This study presents four high quality cryo-EM structures of ADP-actin filaments formed from skeletal α-, cardiac α-, cytoplasmic β- and cytoplasmic γ-actin. These structures are important for understanding the functional differences among these actin isoforms. This work is of significant general interest, because actin filaments, composed of different actin isoforms, have a critical role in a number of physiological processes from muscle contraction to cell migration and division. https://doi.org/10.7554/eLife.82015.sa0 Decision letter eLife's review process eLife digest The protein actin is important for many fundamental processes in biology, from contracting muscle to dividing a cell in two. As actin is involved in such a variety of roles, human cells have slightly different versions of the protein, known as isoforms. For example, alpha-actin is vital for contracting muscle, while beta- and gamma-actin drive cellular processes in non-muscle cells. In order to carry out its various functions, actin interacts with many other proteins inside the cell, such as myosin motors which power muscle contraction. These interactions rely on the precise chain of building blocks, known as amino acids, that make up the actin isoforms; even subtle alterations in this sequence can influence the behavior of the protein. However, it is not clear how differences in the amino acid sequence of the actin isoforms impact actin’s interactions with other proteins. Arora et al. addressed this by studying the structure of four human actin isoforms using a technique called cryo-electron microscopy, where the proteins are flash-frozen and bombarded with electrons. These experiments showed where differences between the amino acid chains of each isoform were located in the protein. Arora et al. then compared their structures with previous work showing the structure of actin bound to myosin. This revealed that the tail-end of the protein (known as the N-terminus) differed in shape between the four isoforms, and this variation may influence how actin binds to others proteins in the cell. These results are an important foundation for further work on actin and how it interacts with other proteins. The structures could help researchers design new tools that can be used to target specific isoforms of actin in different types of laboratory experiments. Introduction Actin isoforms are among the most ubiquitous and abundant structural proteins that facilitate the functional organization of the cytoplasm of eukaryotic cells (Blanchoin et al., 2014; Pollard, 2016). Humans express six actin genes in a tissue-specific and developmentally regulated manner (Kashina, 2020). The gene products are structurally and functionally highly conserved among vertebrates and can be grouped into four muscle actins: skeletal muscle α-actin, smooth muscle α-actin (vascular), cardiac muscle α-actin, smooth muscle γ-actin (enteric), and two nonmuscle actins (ß-actin and γ-actin; Otey et al., 1987; Vandekerckhove and Weber, 1978). Most cells maintain a defined ratio of actin isoforms with muscle and nonmuscle actins representing the main isoforms in muscle and nonmuscle cells, respectively (Kashina, 2020; Otey et al., 1987; Vandekerckhove and Weber, 1978; Kee et al., 2009; Tondeleir et al., 2009; Patrinostro et al., 2017). Actin isoforms have specific and redundant roles in cells and display different biochemistries, cellular localization, and interactions with myosin motors and actin-binding proteins (ABPs; Pollard, 2016; Kashina, 2020; Perrin and Ervasti, 2010; Vedula et al., 2021; Varland et al., 2019; Bunnell et al., 2011; Tondeleir et al., 2012; Baranwal et al., 2012; Diensthuber et al., 2011; Müller et al., 2013; Lee and Dominguez, 2010; Harris et al., 2020; Lappalainen, 2016). Driven by the dominating action of ABPs and signaling proteins, differences between actin isoforms may facilitate the formation of diverse cellular actin networks with distinct compositions, architectures, dynamics, and mechanics that enable fundamental cell functions including adhesion, migration, and contractility (Blanchoin et al., 2014; Tondeleir et al., 2009; Vedula et al., 2021). The altered expression and mutation of the genes encoding for actin isoforms have been linked to human diseases (Tondeleir et al., 2009; Chaponnier and Gabbiani, 2004; Parker et al., 2020). Actin isoforms share high sequence identity at the protein level (~93–99%) and the propensity to self-assemble into helical, polarized filaments (F-actin) from monomers (G-actin; Pollard, 2016; Perrin and Ervasti, 2010; Dominguez and Holmes, 2011; Arnesen et al., 2018). Since the publication of the first crystal structure of G-actin in complex with DNaseI ~30 years ago, extensive studies have advanced our understanding of the structure of monomeric and filamentous actin, polymerization mechanisms, post-translational modifications (PTMs), interaction with drugs, myosin motors, and ABPs at ever-increasing resolution (Dominguez and Holmes, 2011; Holmes et al., 1990; Kabsch et al., 1990; von der Ecken et al., 2016; Chou and Pollard, 2019; Zsolnay et al., 2020; Chou and Pollard, 2020; Belyy et al., 2020; Merino et al., 2018; Mentes et al., 2018; Mei et al., 2020; Ducka et al., 2010; Lee et al., 2007; Otomo et al., 2005; Oda et al., 2009; Egelman et al., 1982; Ali et al., 2022; Oda et al., 2020; Gong et al., 2022). Although isoform-specific mechanisms with myosin motors and ABPs that drive functional distinction are widely described, they are poorly understood at the structural level. To address how the structure contributes to the functional distinction of actin isoforms, we employed a combination of recombinant post-translationally modified actins and actins purified from native source to obtain pure, single-isotype preparations of individual actin isoforms to perform cryo-electron microscopy (cryo-EM) analyses. Specifically, we used our previously established Pick-ya actin method to recombinantly produce human ß-actin and γ-actin in an engineered Pichia pastoris strain that expresses the human N-acetyl transferase NAA80 and histidine methyl transferase SETD3 to ensure uniform Nt-acetylation and methylation of H72/H73, a conserved PTM profile of vertebrate actins (Hatano et al., 2020). Skeletal muscle α-actin and cardiac muscle α-actin were purified from rabbit skeletal muscle and the left ventricle of a bovine heart, respectively. At the protein level, all actin isoforms are conserved across vertebrates, allowing us to compare our structures to previous structures of filamentous actin from other vertebrate species in the correct physiological context (Figure 1—figure supplement 1; Pollard, 2016; Perrin and Ervasti, 2010; Dominguez and Holmes, 2011). Our 2.99–3.38 Å resolution structures of filamentous actin isoforms show that the N-termini of bare muscle and nonmuscle actins have different orientations that contribute to distinct binding interfaces for myosin motors and likely other ABPs. Results High-resolution structures and general principles of actin isoforms To determine the structural characteristics and differences between actin isoforms, we solved the high-resolution structures of single-isotype skeletal muscle α-actin (3.37 Å), cardiac muscle α-actin (3.07 Å), ß-actin (2.99 Å), and γ-actin (3.38 Å) in the filamentous state using cryo-EM (Figure 1A–D, Figure 1—figure supplement 1, Table 1). The structures show local resolutions ranging from 2.1 to 3.5 Å for skeletal muscle α-actin, 1.9–3.3 Å for cardiac muscle α-actin, 1.5–3.0 Å for ß-actin, and 2.1–4.4 Å for γ-actin (Figure 1—figure supplement 2). For all actin isoforms, the highest local resolutions were obtained in the central core region. Lower local resolutions were obtained in the most surface-exposed and flexible regions such as the N-terminus and the D-loop as expected. All actin structures were solved in the Mg2+·ADP state (Figure 1E, Figure 2) and have been obtained without the use of stabilizing drugs that may interfere with the filament structure (Diensthuber et al., 2011; Isambert et al., 1995; Zimmermann et al., 2015). Figure 1 with 8 supplements see all Download asset Open asset Cryo-electron microscopy (cryo-EM) filament structures of actin isoforms. (A) Helical reconstruction of skeletal muscle α-actin, (B) cardiac muscle α-actin, (C) β-actin, and (D) γ-actin. Views in (B–D) are according to (A). Four individual actin protomers in the filament are shown and denoted with italic numbers. The pointed (−) and barbed (+) ends are indicated. (E) Representative key regions of actin isoforms with corresponding cryo-EM densities in transparent surface representation are shown. The protein backbone and amino acid side chains are shown in licorice and stick representation, respectively. Throughout this work, amino acids are numbered according to the sequence of mature actin isoforms (Figure 1—figure supplement 1C). Figure 2 with 1 supplement see all Download asset Open asset Conserved nucleotide-binding cleft active site in actin isoforms. (A–D) Coordination of Mg2+·ADP in the nucleotide-binding cleft of skeletal muscle α-actin (A), cardiac muscle α-actin (B), β-actin (C), and γ-actin (D). Underlines indicate locations of amino acid substitutions between actin isoforms. The protein backbone and side chains are shown in licorice and stick representation, respectively. ADP is shown in cyan-colored stick representation. Electron densities for key amino acids in the nucleotide-binding cleft active site of actin isoforms are shown. Schematic representations of key interactions in the nucleotide-binding cleft active sites of the respective actin isoforms are shown in the right panel. The schematics are not drawn to scale. Table 1 Data collection, image processing, and structure characteristics summary. MapSkeletal muscle α-actinCardiac muscle α-actinβ-actinγ-actinData collectionMicroscopeFEI Titan Krios G3iFEI Titan Krios G3iFEI Titan Krios G3iFEI Titan Krios G3iVoltage (kV)300300300300DetectorGatan K3Gatan K3Gatan K3Gatan K3Automation softwareEPUEPUEPUEPUEnergy filter slit width (eV)20202020Recording modeSuper-resolutionSuper-resolutionSuper-resolutionSuper-resolutionMagnification (nominal)81,00081,00081,00081,000Movie micrograph pixel size (Å)0.8910.8910.8910.891Total Dose rate (e−/Å2)65605065Defocus range (µm)–0.5 to –2.5–0.5 to –2.5–0.5 to –2.5–0.5 to –2.5Spherical aberration (mm)0.010.010.010.01Movies2046144413522952Total extracted particles261,195657,300279,1201,249,379Total # of refined particles185,406657,041263,9111,009,372ReconstructionEMDB codeEMD-27548EMD-27549EMD-27572EMD-27565Box size350256256256SymmetryhelicalhelicalC1C1Map sharpening B-factor (Å2)–90–149–81–201Resolution (global) (Å)3.373.072.993.38Structure building and validationPDB ID8 DMX8DMY8DNH8DNFModel buildingCootCootCootCootRefinement programPhenixPhenixPhenixPhenixRefinement targetReal-spaceReal-spaceReal-spaceReal-spaceRMSD from ideal valuesBond length (Å)0.020.020.040.03Bond Angles (0)0.4930.4940.7610.711Ramachandran favored (%)97.6196.6896.2196.20Ramachandran allowed (%)2.393.323.793.46Ramachandran outliers (%)0000.34MolProbity Score1.421.421.721.72Structures CharacteristicsSpeciesRabbitBovineHumanHumanAmino acid resolved4–3752–3751–3741–374PTMs resolvedH73H73D1/H72E1/H72 Our cryo-EM maps allowed us to build unambiguous models of actin isoforms in which secondary structure information including the side chains, the nucleotide and associated cation (Mg2+·ADP), and PTMs were apparent from the densities (Figure 1E, Figure 1—figure supplement 2). This allowed us to resolve the N-terminus of actin isoforms that is often disordered or missing in prior structures (Kudryashov and Reisler, 2013). For ß-actin and γ-actin, we could resolve the entire N-terminus starting from amino acids D1 and E1, respectively. For skeletal muscle α-actin and cardiac muscle α-actin, we could resolve the N-terminus starting from amino acids E4 and D2, respectively. The lack of resolvable density for the very first amino acid of cardiac muscle α-actin and the first three amino acids of skeletal muscle α-actin may be attributed to a nonuniform PTM pattern of native actin isoforms prepared from muscle compared to our recombinant nonmuscle actin isoforms with uniform PTM pattern in that the entire N-terminus region could be resolved (Figure 1E, Figure 1—videos 1–5). Consistent with the high sequence conservation across actin isoforms (Figure 1—figure supplement 1B–C), our reconstructions show the characteristic double-stranded actin helix with a helical rise of ~27.6 Å to 28 Å and a helical twist of ~–166.5° to –168° (Figure 1A–D, Figure 1—figure supplement 1A) that has been observed in numerous previous structural studies, including previous high-resolution cryo-EM studies (Holmes et al., 1990; Chou and Pollard, 2019; Mei et al., 2020; Oda et al., 2009; Egelman et al., 1982; Ali et al., 2022; Fujii et al., 2010; Galkin et al., 2015). The actin filament itself is composed of G-actin (42 kDa) protomers that are oriented in the same direction (Holmes et al., 1990). Each protomer folds into four subdomains that are referred to as SD1–SD4 (Figure 1, Figure 1—figure supplement 1A; Kabsch et al., 1990). SD1 and SD2 form the outer domain, and SD3 and SD4 form the inner domain (Figure 1). This domain arrangement results in the formation of two clefts – the nucleotide-binding cleft and the barbed end groove (Figure 1, Figure 1—figure supplement 1A; Pollard, 2016; Dominguez and Holmes, 2011; Merino et al., 2020). The nucleotide-binding cleft between SD2 and SD4 harbors the active site that is occupied by Mg2+·ADP in our structures (Figure 1E, Figure 2, Figure 1—figure supplement 1A). The barbed end groove between SD1 and SD3 represents a major binding interface for myosins and ABPs (von der Ecken et al., 2016). It further mediates longitudinal interfaces within the actin filament. SD2 and SD4 of an actin protomer are at the pointed end, and SD1 and SD3 are at the barbed end (Figure 1—figure supplement 1A; Pollard, 2016; Dominguez and Holmes, 2011; Merino et al., 2020). The longitudinal interface between two adjacent actin protomers involves the extended D-loop located in SD2 of one actin protomer that interacts with amino acids located in SD1 and SD3 of another protomer. Both the N- and C-terminus of the actin protomer are in SD1 (Pollard, 2016; Dominguez and Holmes, 2011; Merino et al., 2020). Similarities and differences between actin isoforms The superposition of all actin isoform structures shows a root-mean-square deviation (RMSD) between 0.83Å to 1.04Å for Cα atoms, indicating an overall similar topology. No significant differences in the pitch of the actin helix were observed between our structures of actin isoforms, emphasizing their overall conserved filamentous structure in the absence of myosin motors or ABPs. Actin isoforms differ by conservative and nonconservative substitutions (Figure 1—figure supplement 1C) that contribute to their distinct biochemical and in vivo functions (Blanchoin et al., 2014; Tondeleir et al., 2009; Vedula et al., 2021). Overall, the amino acid sequence is more conserved among muscle actins than between muscle and nonmuscle actins (Figure 1—figure supplement 1C). A structural comparison of our cryo-EM reconstructions shows amino acid substitutions across isoforms with the positions of substituted amino acids highlighted (Figure 3). Actin isoforms show the largest divergence at the acidic N-terminus within SD1 (Figure 1E, Figure 1—figure supplement 1C, Figure 3A). Of note, amino acids 1–3 in our structure of skeletal muscle α-actin and the first amino acid in our structure of cardiac muscle α-actin are not resolved and therefore not shown in Figure 3. Other substitutions are within SD1 (Figure 3B, Figure 3—figure supplement 1), SD3 (Figure 3C, Figure 3—figure supplement 1), and SD4 (Figure 3D, Figure 3—figure supplement 1). There are no substitutions in SD2, the smallest and most flexible subdomain (Kudryashov and Reisler, 2013). Figure 3 with 1 supplement see all Download asset Open asset Similarities and differences between actin isoforms. (A) Sequence variations at the N-terminus located in SD1 of actin isoforms. (B) Sequence variations in SD1 of actin isoforms. (C) Sequence variations in SD3 of actin isoforms. (D) Sequence variations in SD4 of actin isoforms. SD2 is conserved between actin isoforms. The identical and nonidentical amino acids at sites of substitutions within the actin protomer across isoforms are shown for skeletal muscle α-actin (orange), cardiac muscle α-actin (yellow), β-actin (purple), and γ-actin (teal) as spheres. Note that the first three amino acids of skeletal muscle α-actin and the first amino acid of cardiac muscle α-actin are unresolved in our structures. The protein backbone is shown in licorice representation, and the substituted amino acids are shown in spheres representation. Amino acid substitutions at subdomain interfaces, such as the nucleotide-binding cleft active site of actin isoforms, are likely to influence protein function. To evaluate their possible impact on nucleotide coordination and the structural organization of the active site, we performed a comparative structural analysis. Our cryo-EM reconstructions show that the nucleotide-binding cleft active site is conserved between actin isoforms (Figure 1E and Figure 2). The densities for Mg2+ and ADP were assigned without ambiguity and revealed interactions with amino acids located in SD2 (Q58/Q59 and Y68/Y69) and SD4 (E206/E207, R209/R210, K212/K213, and E213/E214) but also with amino acids located in SD1 (M15/L16, K17/K18, Q136/Q137, and Y336/Y337) and SD3 (D156/D157, M304/M305, Y305/Y306, and K335/K336; Figure 2). While most of the interactions are polar and electrostatic, amino acid Y305/Y306 forms π-π interactions with the adenine ring of ADP. The superimposition of the nucleotide cleft active sites of our four structures of actin isoforms shows small differences in the positions of the nucleotide (RMSD ~0.44–0.47 Å) and the bound Mg2+ (RMSD ~0.5–1.3 Å), especially in the position of the β-phosphate group relative to the α-phosphate group (Figure 2, Figure 2—figure supplement 1). The position of the Mg2+ moves relative to the position of the β-phosphate group (Figure 2—figure supplement 1). The superimposition of the nucleotide-binding cleft active sites further shows small local rearrangements of side chains of conserved amino acids (Figure 2—figure supplement 1), including R182/R183 and K335/K336. Amino acids M15/L16 and M304/M305 form hydrophobic interactions in the nucleotide-binding cleft active site (Figure 2). Amino acid substitution M15/L16 between nonmuscle and muscle actin does not alter the overall topology of the nucleotide-binding site. Instead, the longer side chain of M15 in nonmuscle actins, located in a loop that protrudes into the nucleotide-binding cleft, acts as an extended lid that flanks the active site and shields the phosphate groups of ADP (Figure 2, Figure 2—figure supplement 1). Near the nucleotide-binding cleft active site, amino acids C10 and V17 in muscle actins are substituted with V9/I9 and C16 in β/γ-actin (Figure 3, Figure 1—figure supplement 1C). These reciprocal amino acid substitutions maintain the overall oxidation-reduction environment within filamentous actin isoforms which is important for its dynamic properties and the interaction with some regulatory proteins (Farah et al., 2011; Lassing et al., 2007; Wilson et al., 2016; Terman and Kashina, 2013). The analysis of interprotomer interfaces in our four structures of actin isoforms (Figure 4, Figure 5, Figure 4—figure supplement 1) showed that longitudinal interactions are mainly mediated by hydrophilic amino acids that are likely to enable interactions with water molecules that were recently shown to mediate interprotomer contacts within the filament core (Figure 4, Figure 4—figure supplement 1; Reynolds et al., 2022). Amino acid substitutions at the longitudinal interprotomer interface (also called long pitch helix interface) include L175/M176, T200/V201, Q224/N225, C271/A272, F278/Y279, and V286/I287. These substitutions may influence the stability of the promoters based on their ability to interact with the solvent and other protomer residues (Reynolds et al., 2022). The transverse interprotomer interface (also called short pitch helix interface) is formed through hydrophilic and hydrophobic interactions (Figure 5, Figure 4—figure supplement 1). In contrast to interactions at the longitudinal interface, most of the transverse interprotomer interactions are direct and not mediated by solvent. A single amino acid substitution (V286/I287) is at the transverse interprotomer This amino acid substitution is located the of the longitudinal and transverse interprotomer interfaces (Figure 3—figure supplement 1). Amino acid is in hydrophobic with and in muscle actins and a surface of the interaction between and in nonmuscle actins is hydrophobic and a surface of in the transverse interprotomer interface (Figure 4—figure supplement and with the D-loop located in SD2 are conserved in our structures of actin isoforms, the critical and conserved role of the D-loop to mediate interprotomer At the of the longitudinal and transverse interprotomer interfaces, and with as central Figure with 1 supplement see all Download asset Open asset structural analysis of the longitudinal interprotomer (A–D) residues at the interprotomer interface of skeletal muscle α-actin (A), cardiac muscle α-actin (B), β-actin (C), and γ-actin (D). protomers in actin isoforms are oriented according to Figure 4—figure supplement Underlines indicate locations of amino acid substitutions between actin isoforms. The protein backbone and side chains are shown in licorice and stick representation, respectively. Figure Download asset Open asset structural analysis of the transverse interprotomer (A–D) residues at the interprotomer interface of skeletal muscle α-actin (A), cardiac muscle α-actin (B), β-actin (C), and γ-actin (D). protomers in actin isoforms are oriented according to Figure 4—figure supplement Underlines indicate locations of amino acid substitutions between actin isoforms. The protein backbone and amino acid side chains are shown in licorice and stick representation, respectively. PTMs are not essential for actin and but also for their interaction with myosin motors and et al., 2019; et al., et al., 2018). Here, we on two widely and highly conserved PTMs of mature vertebrate actins that are important for actin structure and the post-translational of the N-terminus by NAA80 and methylation of by domain protein 3 Kabsch et al., 1990; Terman and Kashina, 2013; et al., 2018; et al., 2019; et al., Both PTMs are in actin prepared from vertebrate and our preparations of recombinant human β- and γ-actin in an engineered Pichia pastoris strain (Hatano et al., 2020). The quality of our density maps allowed us to resolve key PTMs (Figure 1E, Figure 5, Figure 1—figure supplement 3). Specifically, the of resolvable density allowed us to the entire N-terminus including the D1 and the in the cryo-EM reconstructions of β- and γ-actin (Figure The site on the N-terminus is on the filament surface and an to the N-terminus (Figure 1—figure supplement 1C). in previous cryo-EM structures of muscle are no resolvable densities for the very including in our density maps of skeletal muscle α-actin and cardiac muscle α-actin (Figure to nonuniform PTM of muscle In to Nt-acetylation in β- and γ-actin, we could resolve the in all cryo-EM reconstructions of actin isoforms (Figure 1—figure supplement 3). The of key PTMs in our cryo-EM reconstructions that our recombinant human actins post-translationally mature nonmuscle actins (Hatano et al., 2020). actin filament structure motors actin in a manner to and and The myosin can be into with and and and actin and 2009; and In the a binding interface is established between actin and the myosin domain (von der Ecken et al., 2016; and 2021; and Holmes, To determine myosin binding to actin may actin filament we compared our structures of filamentous bare actin isoforms with previous high-resolution cryo-EM structures of compared the structure of nonmuscle bound to γ-actin, the structure of bound to skeletal muscle α-actin and the structure of bound to skeletal muscle α-actin with our structures of bare actin isoforms (Figure also compared the structure of cardiac bound to the cardiac filament muscle α-actin with and Figure supplement 1) with our structure of bare cardiac muscle The structures were they distinct binding and of myosin motors with different et al., 2018; and and 2011). Figure with 2 supplements see all Download asset Open asset The (A) of bare actin isoform structures in the Mg2+·ADP state superimposition of actin isoforms structures and of bare actin structures are shown. (B) of the interface at the D-loop region. For involved in the binding of myosins are highlighted in the respective The between the structures in the two is by a of SD2 in compared to bare actin structures. binding to actin filaments subtle in the filament (Figure of the actin structures showed that the Cα of bare and actin protomers with an of These are similar for myosins from different and of the nucleotide state of the respective domain (Figure The superposition of structures of bare actin isoforms and known structures of actins shows that SD2 a different (Figure and Figure supplement 1, Figure 1). The of the located in SD2, with a Cα of Å (Figure and of bare γ-actin with complex shows the subtle of the D-loop with a Cα of Å in the that subtle in the barbed end groove that not the between and The of the D-loop (RMSD Å) is also observed in the structure of cardiac myosin bound to the filament compared to our structure of bare cardiac muscle α-actin (Figure supplement 1),

Author response: Structural insights into actin isoforms

Editor's evaluation: Structural insights into actin isoforms

Identification and characterization of Iflavirus 3C-like protease processing activities

Specificity and Affinity of Natural Product Cyclopentapeptide Inhibitors against A. fumigatus, Human, and Bacterial Chitinases

Novel hybrid antiviral VTRRT-13V2.1 against SARS-CoV2 main protease: retro-combinatorial synthesis and molecular dynamics analysis

The ongoing coronavirus infectious disease (COVID-19) pandemic caused by severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) still urgently requires effective treatments. The 3C-like (3CL) protease of SARS-CoV-2 is a highly conserved cysteine protease that plays an important role in the viral life cycle and host inflammation, providing an ideal target for developing broad-spectrum antiviral drugs. Herein, we describe the discovery of a large number of herbs mainly produced in Heilongjiang Province, China, that exhibited different inhibitory activities against SARS-CoV-2 3CL protease. We confirmed that Syringa reticulata, which is used for clinical treatment of chronic bronchitis and asthma, is a specific and potent inhibitor of 3CL protease. A 70 % ethanol extract of S. reticulata dose-dependently inhibited the cleavage activity of 3CL protease in a fluorescence resonance energy transfer assay with an IC50 value of 0.0018 mg/mL, but had minimal effect in pseudovirus-based cell entry and luciferase-based RNA-dependent RNA polymerase assays. These results suggest that S. reticulata will be a potential leading candidate for COVID-19 treatment.

Crystallographic and Functional Analysis of the ESCRT-I /HIV-1 Gag PTAP Interaction

Viral protease inhibitors are remarkably effective at blocking the replication of viruses such as human immunodeficiency virus and hepatitis C virus, but they inevitably lead to the selection of inhibitor-resistant mutants, which may contribute to ongoing disease. Protease inhibitors blocking the replication of coronavirus (CoV), including the causative agents of severe acute respiratory syndrome (SARS) and Middle East respiratory syndrome (MERS), provide a promising foundation for the development of anticoronaviral therapeutics. However, the selection and consequences of inhibitor-resistant CoVs are unknown. In this study, we exploited the model coronavirus, mouse hepatitis virus (MHV), to investigate the genotype and phenotype of MHV quasispecies selected for resistance to a broad-spectrum CoV 3C-like protease (3CLpro) inhibitor. Clonal sequencing identified single or double mutations within the 3CLpro coding sequence of inhibitor-resistant virus. Using reverse genetics to generate isogenic viruses with mutant 3CLpros, we found that viruses encoding double-mutant 3CLpros are fully resistant to the inhibitor and exhibit a significant delay in proteolytic processing of the viral replicase polyprotein. The inhibitor-resistant viruses also exhibited postponed and reduced production of infectious virus particles. Biochemical analysis verified double-mutant 3CLpro enzyme as impaired for protease activity and exhibiting reduced sensitivity to the inhibitor and revealed a delayed kinetics of inhibitor hydrolysis and activity restoration. Furthermore, the inhibitor-resistant virus was shown to be highly attenuated in mice. Our study provides the first insight into the pathogenicity and mechanism of 3CLpro inhibitor-resistant CoV mutants, revealing a low genetic barrier but high fitness cost of resistance. Importance: RNA viruses are infamous for their ability to evolve in response to selective pressure, such as the presence of antiviral drugs. For coronaviruses such as the causative agent of Middle East respiratory syndrome (MERS), protease inhibitors have been developed and shown to block virus replication, but the consequences of selection of inhibitor-resistant mutants have not been studied. Here, we report the low genetic barrier and relatively high deleterious consequences of CoV resistance to a 3CLpro protease inhibitor in a coronavirus model system, mouse hepatitis virus (MHV). We found that although mutations that confer resistance arise quickly, the resistant viruses replicate slowly and do not cause lethal disease in mice. Overall, our study provides the first analysis of the low barrier but high cost of resistance to a CoV 3CLpro inhibitor, which will facilitate the further development of protease inhibitors as anti-coronavirus therapeutics.

Structure of an Asymmetric Ternary Protein Complex Provides Insight for Membrane Interaction