Nucleocapsid assembly in pneumoviruses is regulated by conformational switching of the N protein.

Non-segmented, (-)RNA viruses cause serious human diseases. Human metapneumovirus (HMPV), an emerging pathogen of this order of viruses (Mononegavirales) is one of the main causes of respiratory tract illness in children. To help elucidate the assembly mechanism of the nucleocapsid (the viral RNA genome packaged by the nucleoprotein N) we present crystallographic structures of HMPV N in its assembled RNA-bound state and in a monomeric state, bound to the polymerase cofactor P. Our structures reveal molecular details of how P inhibits the self-assembly of N and how N transitions between the RNA-free and RNA-bound conformational state. Notably, we observe a role for the C-terminal extension of N in directly preventing premature uptake of RNA by folding into the RNA-binding cleft. Our structures suggest a common mechanism of how the growth of the nucleocapsid is orchestrated, and highlight an interaction site representing an important target for antivirals.

Introduction

Viruses possessing a non-segmented, single-strand, negative-sense RNA genome are the causative agents of many serious human illnesses. Notable members belonging to this group of viruses (Mononegavirales) include measles, rabies, Ebola, respiratory syncytial virus (RSV) and Human metapneumovirus (HMPV). HMPV (Paramyxoviridae, subfamily Pneumovirinae) is a leading cause of serious respiratory tract infections in children, the elderly, and immunocompromised individuals (Boivin et al., 2003; Osterhaus and Fouchier, 2003; van den Hoogen et al., 2001). In all members of the Mononegavirales, the RNA genome is packaged in the form of a nucleocapsid, a ribonucleoprotein complex consisting of polymerized viral nucleoproteins (N) and RNA (Ruigrok et al., 2011). Besides protecting the viral genome from host nucleases, the nucleocapsid serves as the template for transcription by the viral RNA-dependent RNA polymerase L. Nucleocapsid assembly necessitates a pool of monomeric, RNA-free N, termed N0, which is kept in an unassembled state through an interaction with an N-terminal portion of the polymerase cofactor P, until delivered to the sites of viral RNA synthesis (Ruigrok et al., 2011; Curran et al., 1995; Mavrakis et al., 2006). The P protein is a multifunctional, modular protein containing large intrinsically disordered regions and is found to be tetrameric in HMPV (Leyrat et al., 2013). In addition, P binds to the nucleocapsid via its C–terminus, and mediates the attachment of the RNA-dependent RNA polymerase L. Furthermore in pneumoviruses, P recruits the processivity factor M2-1 (Leyrat et al., 2014). A great deal of effort has been spent on understanding the functions of P and recent crystal structures of P bound to N proteins (N0-P) from vesicular stomatitis virus (VSV), Ebola virus, Nipah virus, and measles virus have highlighted its role in preventing assembly of N by blocking the C-terminal and N-terminal extensions of N (CTD-arm and NTD-arm) which facilitate N oligomerization (Leyrat et al., 2011; Guryanov et al., 2015; Leung et al., 2015; Yabukarski et al., 2014). However, there is still paucity in our understanding of the molecular details behind the proposed mechanisms, specifically regarding how P-bound N is released, attaches to the nucleocapsid and is loaded with RNA. To address these questions in the mechanism of N-chaperoning by P and nucleocapsid assembly we performed a structural analysis of assembled and unassembled N from HMPV. Our structure of N0-P reveals a conformational change, in which the negatively charged CTD-arm of N occupies the positively charged RNA binding site via specific and conserved interactions. Together with our RNA-bound structure of N these data imply a mechanism of how the growth of nucleocapsid filaments is coordinated in HMPV and related viruses.

Results and discussion

Biochemical studies of the nucleocapsid building block N are complicated by the fact that N proteins have a strong tendency to irreversibly oligomerize and bind host nucleic acids immediately upon recombinant expression (Gutsche et al., 2015; Tawar et al., 2009). One technique to mitigate this problem is to truncate regions of N that facilitate oligomerization (Yabukarski et al., 2014). To stabilize monomeric full-length N0 we fused the N-terminal domain of P to N, a strategy that has seen success with nucleoproteins from other viruses (Guryanov et al., 2015; Kirchdoerfer et al., 2015). We obtained crystals of RNA-free HMPV N in a monomeric state and bound to a P peptide at 1.9 Å resolution by adding trace amounts of trypsin (Dong et al., 2007) to prune flexible loops and promote crystallization (Figure 1—figure supplement 1 and Table 1). In the structure, the P peptide is firmly nestled into a hydrophobic surface of the C-terminal domain of N (CTD) primarily composed of α-helices αC1 and αC2 (Figure 1A,B). Ile9, Leu10 and Phe11 of P occupy key positions and insert into this hydrophobic groove (Figure 1B). Unlike the N0-P structure recently reported for measles (Guryanov et al., 2015), we find that the linker connecting N and P in our chimeric construct has been cleaved prior to crystal growth. The P peptide wraps around the CTD and residues 12–28 form an alpha helix that lies atop N (Figure 1B). This helix is initiated at Gly12 and pinned to the CTD through an aromatic side-to-face interaction of Phe23 with Tyr354 of N, both residues belonging to the so-called mir motif which is conserved within Pneumovirinae (Karlin and Belshaw, 2012). This result is consistent with an earlier study, in which alanine mutations of the corresponding residues in respiratory syncytial virus resulted in a drop of polymerase activity by more than 75% in a minireplicon system (Galloux et al., 2015).

Figure 1—figure supplement 1.

Construct design and purification of the HMPV N0-P hybrid.

(A) Schematic of the N0-P hybrid construct. The N-terminal (NTD) and C-terminal (CTD) domains of N are coloured in dark and light blue, respectively. The N-terminal and C-terminal arms are indicated. The first 40 residues of the HMPV P protein (shown in orange) were cloned at the C-terminus of N, immediately following the CTD-arm. (B) size exclusion chromatogram (Superdex 75) and (C) accompanying SDS-PAGE analysis of the last purification step of N0-P. Protein elution was monitored by the absorbance at 280 nm.

DOI: http://dx.doi.org/10.7554/eLife.12627.004

Table 1.

Data collection and refinement statistics.

DOI: http://dx.doi.org/10.7554/eLife.12627.005

	N0-P	N-RNA
Data collection
Space group	P 1	C 2 2 21
Cell dimensions
a, b, c (Å)	40.9, 62.8, 86.7	202.0, 233.2, 203.6
α, β, γ (°)	91.0, 96.4, 109.0	90, 90, 90
Wavelength (Å)	0.979	0.917
Resolution (Å)	28.42-1.86 (1.91-1.86)	101.19-4.17 (4.28-4.17)
CC (1/2)	1.00 (0.47)	1.00 (0.38)
Rmerge	0.055 (0.590)	0.220 (2.924)
I / σI	9.2 (1.1)	9.2 (1.0)
Completeness (%)	94.8 (75.0)	99.9 (100)
Redundancy	1.7 (1.6)	13.5 (13.8)
Refinement
Resolution (Å)	28.42-1.86	101.19-4.17
No. reflections	64451 (3743)	36125 (2617)
Rwork / Rfree	17.1/20.53	19.1/23.0
No. atoms
Protein	5707	27957
Non-protein	588	1400
B-factors
Protein	34.54	216.06
Non-protein	42.56	215.08
R.m.s. deviations
Bond lengths (Å)	0.007	0.010
Bond angles (°)	1.000	1.120
Ramachandran plot quality
Favoured (%)	99.72	95.01
Allowed (%)	0.28	4.96
Outliers (%)	0.00	0.03

Numbers in parentheses refer to the highest resolution shell.

Rfree was calculated as per Rwork for a 5% subset of reflections that was not used in the crystallographic refinement.

Molprobity scores are included in the Methods section.

Figure 1.

Structure of the HMPV N0-P complex.

(A) Crystal structure of RNA-free HMPV N0 bound to P1-28. The C-terminal domain (CTD) of N is colored in light blue and the N-terminal domain (NTD) in dark blue. Secondary structure elements involved in the interaction with P are indicated. The P peptide is colored in orange. (B) Residues that are important in facilitating the interaction between P and N are shown in stick representation. Conserved hydrophobic residues of the P binding site are colored in yellow. (C) Multiple sequence alignment of N proteins from Paramyxoviridae members. Conserved residues of the P-binding site are highlighted in yellow and correspond to those in B. Virus name abbreviations are given in Methods. (D) N0-P complexes throughout Mononegavirales. Surface representations of N-CTDs of HMPV, Nipah virus (PDB ID:4CO6), Ebola virus (PDB ID:4YPI) and Vesicular stomatitis virus (PDB ID:3PMK), colored by electrostatics. CTDs are shown in the same orientation. Bound P proteins (VP35, in the case of Ebola virus) are colored in orange. The red dotted circle indicates a P-binding sub-region which is shared in all structures. Arrows are explained in the accompanying text.

DOI: http://dx.doi.org/10.7554/eLife.12627.003

(A) Schematic of the N0-P hybrid construct. The N-terminal (NTD) and C-terminal (CTD) domains of N are coloured in dark and light blue, respectively. The N-terminal and C-terminal arms are indicated. The first 40 residues of the HMPV P protein (shown in orange) were cloned at the C-terminus of N, immediately following the CTD-arm. (B) size exclusion chromatogram (Superdex 75) and (C) accompanying SDS-PAGE analysis of the last purification step of N0-P. Protein elution was monitored by the absorbance at 280 nm.

DOI: http://dx.doi.org/10.7554/eLife.12627.004

Alignment of Paramyxoviridae N sequences revealed that many hydrophobic residues lining the P-binding surface of αC1 through αC2 are shared within the family (Figure 1C). For all known N0-P complexes (Leyrat et al., 2011; Leung et al., 2015; Yabukarski et al., 2014), P binds to the CTD of N (Figure 1D). Interestingly, although the specific interaction sites diverge (Figure 1D, indicated by white and black arrows), a sub-region of the CTD (Figure 1D, indicated by dotted circle) is bound by P in all structures, indicating that it is widely conserved throughout Mononegavirales.

To provide a rationale for the molecular switching between the monomeric, P-bound state and the assembled, RNA-bound state, a direct comparison at the atomic level is necessary. To this end, we purified and crystallized assembled HMPV N in the form of a decameric N-RNA ring (Figure 2—figure supplement 1 and Table 1). By exploiting the ten-fold non-crystallographic symmetry in the rings, we were able to obtain excellent electron density maps at 4.2-Å resolution (Figure 2—figure supplement 2A–C) and build a reliable model (Karplus and Diederichs, 2012) (Figure 2—figure supplement 2D). Assembled HMPV decameric N-RNA rings are ~0.5 MDa in molecular mass and 160 Å in diameter and 70 Å in height (Figure 2A). The observed RNA binding mode is similar to that seen in the related RSV N-RNA structure (Tawar et al., 2009). The RNA wraps around the N ring and wedges tightly in the cleft between the NTD and CTD of N, which is lined by positively charged residues (Figure 2A and Figure 2—figure supplement 3). In members of the Paramyxovirinae, the number of nucleotides in the viral genome is required to be a multiple of six (Calain and Roux, 1993) and the structural basis for this so-called rule of six has been elucidated recently (Gutsche et al., 2015). In members of the Pneumovirinae, however, this rule is not observed (Tawar et al., 2009). Our structure further highlights this difference; with each N subunit contacting seven RNA nucleotides (Figure 2—figure supplement 2C and Figure 2—figure supplement 3B).

Figure 2—figure supplement 1.

Purification of HMPV N-RNA and characterisation of oligomeric state.

(A) Size exclusion chromatogram (Superose 6) of N-RNA after purification from E. coli. The fractions containing N-RNA are indicated by a red bar. The broad peak centred around 10 mL (indicated with white arrow) constitutes nucleic acid co-purified from the expression host as indicated by the ratio of absorption at 260 and 280 nm. (B) SDS-PAGE analysis of the fractions marked with red bar in a. (c) Purified N-RNA was analysed by transmission electron cryomicroscopy, showing oligomeric rings. (D) 2D-class averages of N-RNA rings reveal three oligomeric states: 9-mers, 10-mers and 11-mers (as indicated with white labels). The population distribution of the different oligomeric states is indicated in the accompanying pie chart.

DOI: http://dx.doi.org/10.7554/eLife.12627.007

Figure 2—figure supplement 2.

Electron density maps of N-RNA.

(A–C) Samples of electron density of the N-RNA crystal at 4.2 Å. A 2Fo-Fc map contoured at 1.0 σ after density modification with Parrot and B-factor map sharpening is shown. (A) zoomed-out overview of the density, (B) close-up view of two consecutive helices and (C) density for the bound RNA. (D) Data and model quality. Comparison of the correlation of the true signal CC* with CCfree and CCwork. The CC* plot shows that there is useful information up to a resolution of 4.2 Å. CCwork and CCfree values below CC* show that the model is not overfitting the data.

DOI: http://dx.doi.org/10.7554/eLife.12627.008

Figure 2.

Comparison of N in assembled RNA-bound and monomeric RNA-free states.

(A) top- and side-views of RNA-bound HMPV subnucleocapsid rings. N protomers and RNA are shown as surfaces with RNA rendered in brown. The diameter and height of the ring are indicated. (B) Three adjacent protomers of assembled RNA-bound N are shown viewed looking outwards from the centre of the ring, with the middle subunit rendered as surface. The exchange subdomains (NTD- and CTD-arm) that facilitate assembly of N are indicated. (C) The overlay with P1-28 (orange) bound to the middle protomer shows that the P-binding site overlaps with that of the NTD- and CTD-arms and that binding is mutually exclusive. (D) Hinge-motion of NTD and CTD of N. Monomeric N0 is superposed onto a single protomer of assembled, RNA-bound N (N-RNA, shown in grey). The NTD pivots by 10 degrees relative to the CTD (indicated). For clarity, only the NTD and CTD of the two states are shown. (E) showing N0, and (F) showing N-RNA, close-up of the pivot point facilitating the hinge-motion of N. The white arrow in F indicates where the hinge region uncoils, allowing pivoting. For clarity, the P-peptide and the CTD-arm are omitted in E and F.

DOI: http://dx.doi.org/10.7554/eLife.12627.006

(A) Size exclusion chromatogram (Superose 6) of N-RNA after purification from E. coli. The fractions containing N-RNA are indicated by a red bar. The broad peak centred around 10 mL (indicated with white arrow) constitutes nucleic acid co-purified from the expression host as indicated by the ratio of absorption at 260 and 280 nm. (B) SDS-PAGE analysis of the fractions marked with red bar in a. (c) Purified N-RNA was analysed by transmission electron cryomicroscopy, showing oligomeric rings. (D) 2D-class averages of N-RNA rings reveal three oligomeric states: 9-mers, 10-mers and 11-mers (as indicated with white labels). The population distribution of the different oligomeric states is indicated in the accompanying pie chart.

DOI: http://dx.doi.org/10.7554/eLife.12627.007

(A–C) Samples of electron density of the N-RNA crystal at 4.2 Å. A 2Fo-Fc map contoured at 1.0 σ after density modification with Parrot and B-factor map sharpening is shown. (A) zoomed-out overview of the density, (B) close-up view of two consecutive helices and (C) density for the bound RNA. (D) Data and model quality. Comparison of the correlation of the true signal CC* with CCfree and CCwork. The CC* plot shows that there is useful information up to a resolution of 4.2 Å. CCwork and CCfree values below CC* show that the model is not overfitting the data.

DOI: http://dx.doi.org/10.7554/eLife.12627.008

(A) External side view showing RNA inserted into three neighbouring protomers of assembled N. The two outer protomers are shown as surface representation coloured by electrostatics, highlighting the basic nature of the RNA-binding cavity. The central N subunit is shown in cartoon representation with the NTD coloured in dark blue and the CTD in light blue. RNA is shown in stick representation and coloured in brown. (B) close-up of the RNA-binding site of one N protomer. NTD and CTD are coloured as in A. The seven bound nucleotides are numbered, counting from 3’-end to 5’-end. Important residues interacting with RNA are shown in stick representation.

DOI: http://dx.doi.org/10.7554/eLife.12627.009

(A) Monomeric N0 (blue) is superposed onto RNA-bound N (grey). The dotted arrow indicates the tilting of α-helix αC3 during the transition from N0 to N-RNA. Tyr252 is thereby pushed upwards, facilitating the hinge motion. (B–G) panel of N proteins throughout Mononegavirales for which an aromatic residue (shown in red) can be observed at the same position and orientation, indicating a conserved function despite low overall sequence identity. (H and I) in Paramyxovirinae the aromatic residue located before the hinge region is flipped in the opposite direction in respect to other mononegaviruses.

DOI: http://dx.doi.org/10.7554/eLife.12627.010

Figure 2—figure supplement 3.

RNA-binding cleft of HMPV N.

(A) External side view showing RNA inserted into three neighbouring protomers of assembled N. The two outer protomers are shown as surface representation coloured by electrostatics, highlighting the basic nature of the RNA-binding cavity. The central N subunit is shown in cartoon representation with the NTD coloured in dark blue and the CTD in light blue. RNA is shown in stick representation and coloured in brown. (B) close-up of the RNA-binding site of one N protomer. NTD and CTD are coloured as in A. The seven bound nucleotides are numbered, counting from 3’-end to 5’-end. Important residues interacting with RNA are shown in stick representation.

DOI: http://dx.doi.org/10.7554/eLife.12627.009

Similar to N proteins from other members of Mononegavirales (Tawar et al., 2009; Alayyoubi et al., 2015; Albertini et al., 2006; Green et al., 2006), the NTD- and CTD-arms grasp the neighbouring protomers, thus facilitating assembly of polymeric N (Figure 2B). The NTD-arm packs against the flank of the previous protomer (Figure 2B, the NTD-arm of Ni+1 packs against Ni). The CTD-arm in turn latches onto the top of the CTD of the next protomer (Figure 2B, CTD-arm of Ni-1 latches onto CTD of Ni). We observed that the binding site of the P peptide overlaps with the binding sites of the NTD- and CTD-arms (Figure 2C). Our structures thus provide conclusive evidence that P hampers subdomain exchange between adjacent proteins in Pneumovirinae. This mechanism has also been proposed for a range of viruses thoughout Mononegavirales (Leyrat et al., 2011; Guryanov et al., 2015; Yabukarski et al., 2014; Alayyoubi et al., 2015) and there is mounting evidence that it may be universal throughout the entire viral order.

A hinge-like motion has been proposed by which N alternates between an open, RNA-free conformation (N0) and a closed RNA-bound (N-RNA) conformation (Guryanov et al., 2015; Yabukarski et al., 2014). Comparison of these two states for HMPV reveals a rigid body movement of the NTD relative to the CTD (Figure 2D). The conformational change rotates the NTD towards the CTD by 10°, the interface between the two domains acting as a hinge. At the interface, hinge residues Thr257 and Ala254 play a particularly crucial role. In the open, RNA-free state the hinge is maintained in a helical conformation by stabilization of Ala254 through the side chain of Thr257 and an additional backbone interaction with Thr175 (Figure 2E). Upon RNA binding, Thr257 contacts the backbone of a nucleotide instead of stabilizing Ala254 (Figure 2F). In addition, the loop containing Thr157 retracts to sterically accommodate the RNA chain. Having lost the stabilizing contacts of Thr257 and Thr157, the helical hinge region around Ala254 unravels and becomes flexible (Figure 2F, indicated by white arrow), allowing the relative domain motions of NTD and CTD. Furthermore, we propose that Tyr252 is important in facilitating the hinge motion. Tyr252 is positioned just before the pivot point and packs tightly against αC3 (Figure 2—figure supplement 4A). An aromatic residue at this position is found packing against the same helix in most known structures of N (Figure 2—figure supplement 4B–G). Transition from the RNA-free to RNA-bound state induces a rotation of αC3, exerting upwards pressure on Tyr252 that is conferred onto the NTD (Figure 2—figure supplement 4A). Intriguingly, in structures of Paramyxovirinae N, which obey the rule-of-six, this aromatic is flipped in the opposite direction (Figure 2—figure supplement 4H,I) and contacts RNA (Gutsche et al., 2015), suggesting a similar coupling of RNA-binding and hinge-motion in these viruses.

Figure 2—figure supplement 4.

Role of a conserved aromatic residue in N hinge motion.

(A) Monomeric N0 (blue) is superposed onto RNA-bound N (grey). The dotted arrow indicates the tilting of α-helix αC3 during the transition from N0 to N-RNA. Tyr252 is thereby pushed upwards, facilitating the hinge motion. (B–G) panel of N proteins throughout Mononegavirales for which an aromatic residue (shown in red) can be observed at the same position and orientation, indicating a conserved function despite low overall sequence identity. (H and I) in Paramyxovirinae the aromatic residue located before the hinge region is flipped in the opposite direction in respect to other mononegaviruses.

DOI: http://dx.doi.org/10.7554/eLife.12627.010

The most profound changes between assembled and unassembled states, however, involve the CTD-arm of N, a region that has been little characterized in pneumoviruses. In the polymeric, RNA-bound state of N (N-RNA) the CTD-arm flips upwards and latches onto the next protomer, whilst in the monomeric state (N0) it packs down against the core of N (Figure 3A). The downward, monomeric conformation is stabilized by specific salt-bridges linking the CTD-arm with the core of N (Figure 3B). In this position the negatively charged CTD-arm folds into the positively charged RNA binding cleft, occupying it and directly blocking the binding of RNA (Figure 3A). It is interesting to note, that whilst the CTD-arm blocks the RNA site in HMPV, it is the P peptide that inserts itself there in VSV (Leyrat et al., 2011). Because this is not observed in paramyxoviral N0-P complexes (Guryanov et al., 2015; Yabukarski et al., 2014) we hypothesize that, in Rhabdoviridae, a different strategy has evolved to block off the RNA binding cleft. The question arises how the interactions that hold the downwards-positioned CTD-arm in place are broken when assembly of N-RNA necessitates it flipping into the upwards position. In the RNA-free state, Arg260 and Trp261 contact Glu375, while Arg186 forms a salt-bridge with Asp373 of the CTD-arm (Figure 3B). In the assembled, RNA-bound state these interactions are broken, with Arg186 and Trp261 now positioning RNA nucleotides in the cleft, whilst Arg260 instead fastens onto the NTD-arm of the neighbouring Ni+1 (Figure 3C). The shift from initial stabilization of the inhibitory (downwards) CTD-arm conformation to stabilization of bound RNA and neighbouring N subunit implies that attachment of a new N protomer and insertion of nascent RNA occur concomitantly. This makes sense in the context of viral replication sites, where tetrameric P proteins act as molecular chaperones attaching to the nucleocapsid template, polymerase and free N0, leading to high local concentrations of nucleoprotein and RNA.

Figure 3.

Role of the CTD-arm in inhibiting premature RNA uptake.

(A) Conformational switch of the CTD-arm. The CTD-arm (red) is shown in a upward conformation assumed in the N-RNA state and downward conformation of the N0 state (indicated). (B) Polar interactions fastening the CTD-arm (red) in the downward conformation. Involved residues are shown as sticks. (C) in the assembled state, the CTD-arm is displaced by RNA (shown in brown). The NTD-arm of the neighboring Ni+1 protomer is colored in green. (D) Schematic model of nucleocapsid filament growth. Nascent RNA and the active RdRP complex are indicated. Binding of emerging RNA to Ni primes the displacement of P (colored in orange) and attachment of incoming Ni+1 by liberating the CTD-arm (colored in red). The dotted arrows indicate that CTD-arms switch to the upward conformation and latch onto incoming N during attachment of the next N protomer. (E) Multiple sequence alignment of CTD-arms from Paramyxoviridae family members. Residues are colored using the ClustalX color scheme. The consensus secondary structure is indicated below the alignment. Virus name abbreviations are given in Materials and methods.

DOI: http://dx.doi.org/10.7554/eLife.12627.011

(A) Conformational change of the CTD-arm in HMPV as in Figure 3A. (B and C) negative charges within the CTD-arm are topologically conserved in Borna virus (Bornaviridae) Rudolph et al., 2003 and Ebola virus (Filoviridae). Downwards motion of the CTD-arms could position them into the RNA-binding cleft (indicated by dotted arrows), analogous to what is observed in HMPV.

DOI: http://dx.doi.org/10.7554/eLife.12627.012

Based on the comparison of our N-RNA and N0-P structures we suggest a model for nucleocapsid growth (Figure 3D). Upon delivery of fresh N0-P to the growth site, addition of the next N protomer (Ni+1) to the filament necessitates that the CTD-arm of the terminal Ni unbinds and flips upwards (Figure 3D, indicated by dotted arrow), latching onto Ni+1 and displacing P. In our model, this is driven by the formation of new interactions to the NTD- and CTD arms and, importantly, the concerted insertion of nascent RNA into the RNA binding cleft of Ni, with the CTD-arm switching into the upward conformation. In this model the growth of the filament is reminiscent of a zipper closing up with one row of teeth corresponding to nascent viral RNA and the other to newly delivered N subunits which interdigitate in a fluid, concerted motion. The notion that concerted RNA insertion is required for the hand-over of N subunits from P lends additional specificity to the nucleocapsid polymerization reaction.

We hypothesized that the role of the CTD-arm in inhibiting premature RNA binding may be conserved and therefore compared sequences throughout Paramyxoviridae (Figure 3E). We find a semi-conserved LGLT-motif within the CTD-arms which is followed by a stretch of residues with helical propensity. The beginning of this stretch preferentially features negatively charged residues at positions equivalent to HMPV which may in turn pack against the complementary charges of the RNA binding cleft. Indeed, analysis of structures of more distantly related members of Mononegavirales shows that these negatively charged residues are topologically conserved and that a switch to the downward conformation would position these residues into the RNA binding cleft (Figure 3—figure supplement 1).

Figure 3—figure supplement 1.

CTD-arms in other Mononegavirales family members.

(A) Conformational change of the CTD-arm in HMPV as in Figure 3A. (B and C) negative charges within the CTD-arm are topologically conserved in Borna virus (Bornaviridae) Rudolph et al., 2003 and Ebola virus (Filoviridae). Downwards motion of the CTD-arms could position them into the RNA-binding cleft (indicated by dotted arrows), analogous to what is observed in HMPV.

DOI: http://dx.doi.org/10.7554/eLife.12627.012

In conclusion, the reported structures of a paramyxoviral N protein reveal two distinct conformational states, N bound either to the polymerase cofactor P or to RNA. A direct comparison of these two structures provides a molecular level rationale for how nucleocapsid assembly is controlled through P by sterically blocking the binding sites of the NTD- and CTD-arms. In addition, this work elucidates a key role of the CTD-arm in hindering premature RNA insertion into the binding cleft, thus presenting a mechanistic explanation of how premature RNA uptake is directly inhibited in Paramyxoviridae. Peptides of the N0-binding region of P have previously been shown to inhibit replication activity in RSV (Galloux et al., 2015), Nipah virus (Yabukarski et al., 2014), and rabies virus (Castel et al., 2009). The characterization of P-binding surfaces on N proteins is therefore of biomedical importance as these surfaces constitute genuine targets for the development of antivirals.

Materials and methods

Expression and purification of N-RNA rings

The full-length N gene from human metapneumovirus (strain NL1-00, A1) was cloned into the pOPINE expression vector, which includes a C-terminal His-tag, using the In-Fusion system (Takara Clontech, Mountain View, CA) following standard procedures. The construct was verified by sequencing. Rosetta2 E.coli cells harboring the expression plasmid were grown at 37°C in terrific broth containing appropriate antibiotics and expression was induced at an OD600 of 0.8 by adding isopropyl β-D-1-thiogalactopyranoside to 1 mM. The temperature was then lowered to 18°C and after further 18 hrs the cells were harvested by centrifugation (18°C, 20 min, 4000 x g). Cell pellets were resuspended in 40 mL of 25 mM Tris, pH 8, 1 M NaCl per L of culture and lysed by sonication. The lysate was centrifuged (4°C, 45 min, 50000 x g) and the supernatant was filtered and loaded on a column containing pre-equilibrated Ni2+-nitrilotriacetic (NTA) agarose (Qiagen, Netherlands). The column was washed and the protein was eluted in 25 mM Tris, pH 8, 1 M NaCl, 400 mM imidazole. The eluate was further purified by size exclusion chromatography using a Superose6 10/300 column (GE Healthcare, United Kingdom) equilibrated in 25 mM Tris, pH 8, 1 M NaCl. The protein was buffer exchanged into 25 mM Tris, pH 8, 150 mM NaCl, 500 mM NDSB201, 50 mM Arginine using a PD10 column (GE Healthcare) and then concentrated to ~4 mg/mL for crystallization.

Expression and purification of the N0-P hybrid

The N0-P hybrid gene was generated by fusing the sequence corresponding to the first 40 residues of HMPV P (strain NL1-00, A1) to the 3’ end of the full-length N gene using overlapping primer PCR. The resulting hybrid construct was cloned into POPINE as described above and verified by sequencing. Protein expression was carried out as described for N, above. Cell pellets were resuspended in 20 mM Tris, pH 7, 1M NaCl, lysed by sonication and the lysate was subsequently centrifuged (4°C, 45 min, 50000 x g). The supernatant was purified using a column containing pre-equilibrated Ni2+-NTA agarose and elution was carried out using 20 mM Tris, pH 7, 1M NaCl, 300 mM imidazole. The protein was then buffer exchanged into 20 mM Tris, pH 7, 100 mM NaCl and loaded onto a HiTrap Heparin HP column (GE Healthcare) for further purification using a stepwise NaCl gradient. Finally, the N0-P hybrid was gel-filtrated using a Superdex 75 column (GE Healthcare) equilibrated with 20 mM Tris, pH 7, 100 mM NaCl, and concentrated to ~7 mg/mL for crystallization.

Crystallization and data collection

Sitting drop, vapor diffusion crystallization trials were set up in 96-well Greiner plates using a Cartesian Technologies robot (Walter et al., 2005). A diamond-like, diffraction quality N-RNA crystal was obtained after 132 days in mother liquor containing 100 mM Tris/Bicine, pH 8.5, 90 mM NPS (NaN03, Na2HPO4, (NH4)2SO4), 37.5% methyl-2 4-pentanediol, polyethylene glycol 1000 and polyethylene glycol 3350 of the MORPHEUS crystal screen. The crystal was frozen in liquid nitrogen and diffraction data up to 4.2 Å were recorded at 100 K on the I04-1 beamline at Diamond Light Source, Didcot, UK.

For the N0-P hybrid, crystals were obtained via in-situ proteolysis (Dong et al., 2007) using 1 µg of trypsin per 1000 µg of sample. The trypsin was added to the concentrated N0-P preparation just before setting up the crystallization trials. Initial crystals formed in mother liquor containing 100 mM PCB System, pH 7, 25% polyethylene glycol 1500 and improved crystals could be grown with additives of the Hampton Silver Bullet screen (9 mM 1,2-diaminocyclohexane sulfate, 6 mM diloxanide furoate, 17 mM fumaric acid, 10 mM spermine, 9 mM sulfaguanidine and 20 mM HEPES, pH 6.8). The crystals were cryoprotected in 25% glycerol and frozen in liquid nitrogen. Diffraction data up to 1.9 Å were recorded at 100 K on the I04 beamline at Diamond Light Source, Didcot, UK. All data were processed and scaled with XIA2 (Winter, 2010).

Structure determination and refinement

The structure of N0-P was solved by molecular replacement using PHASER (McCoy et al., 2007) with the structure of RSV N (Tawar et al., 2009) as a search model. Iterative rounds of refinement using PHENIX (Adams et al., 2010) with TLS parameters and manual building in COOT (Emsley and Cowtan, 2004) resulted in a model for HMPV N starting at residue 30 and ending at residue 383 of the total 394. Residues 101 to 111 were found to be disordered and were not included in the model. Of the 40 P residues contained in our N0-P construct the first 28 were well-resolved.

The structure of the RNA-bound subnucleocapsid ring was solved with PHASER (McCoy et al., 2007) using a decameric model of our high-resolution HMPV N structure as a search model. Initially, we performed iterative rounds of manual building with COOT (Emsley and Cowtan, 2004) and refinement using PHENIX (Adams et al., 2010) with non-crystallographic symmetry (NCS) constraints to lower the parameter to observations ratio. To aid model building we made use of density modified maps obtained with PHENIX RESOLVE (Adams et al., 2010) and Parrot of the CCP4 suite (Winn et al., 2011) in combination with B-factor sharpening. Later stages of refinement were performed with autoBuster (Smart et al., 2012), applying NCS restraints, TLS parameters and using our high-resolution N0-P structure to generate reference model restraints. Structures were validated with MolProbity (Chen et al., 2010) resulting in overall MolProbity scores of 0.95 and 2.22 for N0-P (at 1.9 Å) and N-RNA (at 4.2 Å), respectively. Refinement and geometry statistics are given in Table 1.

Multiple sequence alignment

Multiple sequence alignments (MSA) were carried out with PROMALS3D (Pei and Grishin, 2014) and figures were prepared with Jalview. Nucleoprotein sequences of the following viruses were used: HMPV, Human metapneumovirus, AMPV, Avian metapneumovirus, RSV, Respiratory syncytial virus, MPV, Murine pneumonia virus, BRSV, Bovine respiratory syncytial virus, CPV, Canine pneumonia virus, MeV, Measles virus, MuV, Mumps virus, RPV, Rinderpest virus, HPIV5, Human parainfluenza virus 5, SeV, Sendai virus, HPIV2, Human parainfluenza virus 2, SV41, Simian virus 41, NiV, Nipah virus, HeV, Hendra virus, CDV, Canine distemper virus, MENV, Menangle virus.

Electron microscopy

N-RNA rings were analysed via electron cryomicroscopy (cryo-EM). Aliquots (3 µl) of N-RNA preparations were pipetted onto glow-discharged Cflat holey carbon grids (Protochips, Raleigh, NC) and excess liquid was blotted with filter paper for 3 s. Grids were then plunge-frozen in an ethane-propane mixture at liquid nitrogen temperature using a CP3 plunging device (Gatan). Cryo-EM data were acquired using a 300-kV Polara transmission electron microscope (FEI) equipped with a K2 Summit direct electron detector (Gatan) and using defocus values ranging from -2.0 to -6.0 μm at a calibrated magnification of 37,000x, resulting in a pixel size of 1.35 Å. The contrast transfer function (CTF) parameters were determined using CTFFIND3 (Mindell and Grigorieff, 2003) and 2D-classification was carried out with RELION (Scheres, 2012).

In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.

[Editors’ note: this article was originally rejected after discussions between the reviewers, but the manuscript was accepted after an appeal against the decision.]

Thank you for submitting your work entitled "Nucleocapsid assembly in pneumoviruses is regulated by conformational switching of the N protein" for consideration by eLife. Your article has been reviewed by two peer reviewers, and the evaluation has been overseen by a Reviewing Editor and John Kuriyan as the Senior Editor. Our decision has been reached after consultation between the reviewers and the Reviewing Editor. Based on these discussions, we regret to inform you that your work will not be considered further for publication in eLife. The decision was made not because of the quality of the work, which both reviewers judged to be fine, but rather because of the criteria for novelty and breadth of interest that the journal and its Board of Reviewing Editors have set for the journal.

Renner and colleagues provide evidence that nucleocapsid assembly in pneumoviruses involves a conformational switch of the N protein. That conclusion is based on the structures of an N-protein RNA complex and a soluble form of N, referred to as No, both for human metapneumovirus (HMPV) N. The No structure was obtained in complex with a small N-terminal fragment of P. The structures are of interest and suggest that a conformational change occurs during the process of encapsidation of the nascent RNA. The authors propose, based on the structures, that the C-terminal arm of the last N protein molecule in the growing filament flips up to allow engagement of the new incoming No-P, displacing P concomitant with the insertion of the nascent RNA chain into the newly added N. The process of encapsidation of the nascent RNA lacks a detailed molecular model, and this manuscript may provide some important new clues as to how the process occurs not just for HMPV, but perhaps other viruses in the order mononegavirales (MNV). There are, however, some concerns regarding the No-P structure, and several statements of novelty and precedent in the manuscript that do not seem justified. Moreover, other structures of RNA free, soluble No and N-RNA complexes have been determined for other viruses in the MNV, diminishing the novelty of present work.

1) The No-P structure was solved using an artificial construct in which the N-terminal 40 residues of P were fused to the C-terminal arm of full-length N, and crystals obtained by in situ proteolysis with trypsin. It is uncertain to what extent the structure obtained represents a physiologically relevant form of soluble N or is in fact a form that is simply obtained using the in situ proteolytic approach used here.

2) The conformational switch model proposed suggests contacts that can be tested experimentally using HMPV reverse genetics – this would go a long way to alleviate the concern of point 1.

3) Throughout the manuscript the authors make broad claims of precedent, stating that this is the first virus in the mononegavirales for which structures of both No and N-RNA have been determined. This is not the case – the authors show several previously published structures in Figure 1D. Such statements of precedent should be removed or clarified (see Abstract for example).

Renner and colleagues provide evidence that nucleocapsid assembly in pneumoviruses involves a conformational switch of the N protein. That conclusion is based on the structures of an N-protein RNA complex and a soluble form of N, referred to as No, both for human metapneumovirus (HMPV) N. The No structure was obtained in complex with a small N-terminal fragment of P. The structures are of interest and suggest that a conformational change occurs during the process of encapsidation of the nascent RNA. The authors propose, based on the structures, that the C-terminal arm of the last N protein molecule in the growing filament flips up to allow engagement of the new incoming No-P, displacing P concomitant with the insertion of the nascent RNA chain into the newly added N. The process of encapsidation of the nascent RNA lacks a detailed molecular model, and this manuscript may provide some important new clues as to how the process occurs not just for HMPV, but perhaps other viruses in the order mononegavirales (MNV). There are, however, some concerns regarding the No-P structure, and several statements of novelty and precedent in the manuscript that do not seem justified. Moreover, other structures of RNA free, soluble No and N-RNA complexes have been determined for other viruses in the MNV, diminishing the novelty of present work.

I would like to thank you and the referees for the comments regarding our manuscript “Nucleocapsid assembly in pneumoviruses is regulated by conformational switching of the N protein”. Two of the three main points brought forward by the referees are either unjustified or easy to address.

1) The No-P structure was solved using an artificial construct in which the N-terminal 40 residues of P were fused to the C-terminal arm of full-length N, and crystals obtained by in situ proteolysis with trypsin. It is uncertain to what extent the structure obtained represents a physiologically relevant form of soluble N or is in fact a form that is simply obtained using the in situ proteolytic approach used here.

Linking of binding partners as we have performed in this manuscript is a powerful technique to circumvent problems of unstable binding and increase naturally occurring interaction. The method (reviewed in Sivaraman et al., 2013) is well-established and has been successfully used on a variety of systems, including MHC-receptors and, indeed, viral nucleoproteins (e.g. Kirchdoerfer et al., 2015). Due to the inherent propensity to form oligomeric assemblies such tricks are absolutely necessary for the study of N. The referees categorically denote the method as being artificial but offer no specific criticisms to the structure at hand. Similarly, the use of limited proteolytic digestion is a very common technique in crystallography to facilitate the pruning of flexible loops and thus promote crystallization (Dong et al., 2007). It seems that the major criticism of the referees was that we utilized two common and established techniques which happen to be unfamiliar to them. Indeed, we can bring forward several points that confirm our N0-P structure is correctly folded, even though this should not be necessary: ( (1)) the binding site of P is in line with previous functional studies (Galloux et al., 2015), as mentioned in the manuscript, ( (2)) the P-site is also in line with structures of related viruses, ( (3)) the linker between N and P is cleaved by the in-situ proteolysis yielding non-covalently bound P which should completely negate this point of contention, ( (4)) the overall fold of N follows other N proteins. We therefore cannot understand why our structure is deemed artificial and would like to challenge the referees to bring forward specific criticisms. We addressed the criticisms in our manuscript and explain the rationale of using a chimeric protein and proteolytic digestion. We have also addressed the criticism of data/model quality for our N-RNA structure by including CC*, CCwork and CCfree statistics by resolution shell (Karplus and Diederichs, 2012).

2) The conformational switch model proposed suggests contacts that can be tested experimentally using HMPV reverse genetics – this would go a long way to alleviate the concern of point 1.

We fully agree that testing the proposed interactions using a reverse genetics system is key in further dissecting the mechanism of nucleocapsid assembly. Unfortunately, we lack the resources or necessary collaborations to perform these experiments. This is why our manuscript was written as a short report focused on biophysical and structural analysis of this system and we feel that such experiments would be beyond its scope. However, we could conceive of mutagenesis experiments that could test the interactions in-vitro, and these may provide valuable additional validation

3) Throughout the manuscript the authors make broad claims of precedent, stating that this is the first virus in the mononegavirales for which structures of both No and N-RNA have been determined. This is not the case – the authors show several previously published structures in

Our study aims at structurally characterising assembly of N. The transition between assembled N-RNA and monomeric N0 -P was analysed in-depth in our manuscript. At the point of review, to our best knowledge, there was no instance where an assembled N-RNA and monomeric N0 -P structure was available for the same virus. The only instance where this might be contended is for vesicular stomatitis virus (VSV). However, in this case both structures form oligomeric, assembled rings that are conformationally identical (Leyrat et al., 2011). We could thus, for the first time, analyse the structural transition without having to resort to qualitative comparisons with phylogenetically distant homologues. Similar to major comment #1 the referees unfairly dismissed our claims without specific criticism. We would like to challenge them to show us for which virus they think monomeric N0 -P and assembled N-RNA was available during review. However, during to the lengthy review process, the measles virus N0 -P was published (Guryanov et al., 2015, published online on December 30), meaning that we cannot make the contended claim anymore. To quote the manuscript by Guryanov et al: "Here, for the first time in Paramyxovirus research, our data allow direct comparison of the structure of the nucleoprotein from the same virus in two functional states: a P-bound naive state, and an RNA-bound helical assembly." Consequently, although they were true during the review stage, we have now removed these claims of precedence.