Slowly folding surface extension in the prototypic avian hepatitis B virus capsid governs stability

Hepadnaviruses (hepatotropic DNA viruses) are a family of small enveloped retrotranscribing DNA viruses. In humans, chronic infections with HBV cause ~800,000 deaths per year (Report, 2017). Current therapies can control but rarely cure infection (Fanning et al., 2019; Martinez et al., 2019; Nassal, 2015). Most fundamental aspects of the hepadnaviral lifecycle such as replication of the tiny ~3 kb DNA via reverse transcription of a pregenomic (pg) RNA (Summers and Mason, 1982), or the use of a covalently closed circular (ccc) DNA form (Tuttleman et al., 1986) as episomal persistence reservoir (Nassal, 2015), were uncovered using more tractable model viruses, mostly duck HBV (DHBV) (Schultz et al., 2004), prototype of the genus avihepadnaviridae.

Despite basal similarities in genome size, genetic organization and replication strategy the avian viruses exhibit characteristic differences to human and the other mammalian HBVs (genus orthohepadnaviridae). They lack an X protein, a regulator of cccDNA transcription (Livingston et al., 2017), and a medium sized (M) envelope or surface protein besides the small (S) and large (L) forms; in addition, the avian virus surface proteins are smaller, with only three predicted transmembrane helices versus four in the orthohepadnaviruses. Particularly striking is the much larger size (~260 amino acids) of their single capsid or core proteins (CPs) which in mammalian HBVs comprise only ~180 aa. Recently discovered HBV-like viruses in distant hosts (Lauber et al., 2017) maintain this division. Most employ the small type CP but amphibian and reptilian viruses, e.g. Tibetan frog HBV (Dill et al., 2016) and lizard HBVs (Lauber et al., 2017), encode the large type CP. As the small CPs can obviously perform all essential CP functions (see below, and Venkatakrishnan and Zlotnick, 2016) the biology behind the large type CPs is enigmatic. A key step would be knowledge of their structure but, ironically, here DHBV lags much behind HBV for which various structures are available for recombinant capsid-like particles (CLPs) (Böttcher and Nassal, 2018; Böttcher et al., 1997; Conway et al., 1997; Crowther et al., 1994; Wynne et al., 1999), serum-derived nucleocapsids (Roseman et al., 2005) and virions (Dryden et al., 2006; Seitz et al., 2007).

CPs are multifunctional (Zlotnick et al., 2015; Diab et al., 2018), making the HBV CP (HBc) a prime novel antiviral target (Lahlali et al., 2018; Schlicksup et al., 2018). Most obvious is their role as the building block of the icosahedral viral capsid. Production of progeny virions during infection involves transcription from nuclear cccDNA, translation in the cytoplasm of CP, polymerase and the other viral proteins, CP phosphorylation-regulated encapsidation of a complex of pgRNA and polymerase (Heger-Stevic et al., 2018b; Zhao et al., 2018), and reverse transcription of pgRNA into relaxed circular (RC plus some double-stranded linear (dsL)) DNA. Such nucleocapsids can interact with the surface proteins to be secreted as enveloped virions. Alternatively, they may interact with nuclear import receptors to be redirected into the nuclear pore where they disintegrate (Gallucci and Kann, 2017), releasing the rcDNA for conversion into cccDNA (Nassal, 2015; Schreiner and Nassal, 2017).

How hepadnaviral CPs integrate the different interaction cues to follow one path or the other is poorly understood but must somehow relate to their three-dimensional (3D) structure. Typical small type CPs comprise an N-terminal ~140 aa assembly domain (Birnbaum and Nassal, 1990) and an arginine-rich ~34 aa C-terminal domain (CTD) whose nucleic acid binding capacity is modulated by phosphorylation (Heger-Stevic et al., 2018b; Zhao et al., 2018). Both domains are connected by an ~10 aa linker (Watts et al., 2002).

The assembly domains feature five α-helices (Figure 1—figure supplement 1). α3 and α4 form an antiparallel hairpin providing the interface for dimerization with a second subunit. The resulting four-helix bundles protrude as spikes from the capsid surface. The helix α5 plus the downstream sequence (the ‘hand region’) provide the inter-dimer contacts, mostly arranged in T = 4 icosahedral symmetry which comprises 120 Cp dimers (the minor T = 3 class encompasses 90 dimers). The asymmetric unit is composed of two dimers in spatially different surroundings with four quasi-equivalent monomer conformations, A to D (Figure 1—figure supplement 1). The first residues of the CTD are capsid-internal (Zlotnick et al., 1997) but residues past position 150 usually lack sufficient order for visualization (Böttcher and Nassal, 2018).

Early cryo-EM reconstructions of DHBc suggested a T = 4 arrangement similar to HBc CLPs but with laterally widened spikes (Kenney et al., 1995). Extensive mutagenesis led to a DHBc model (Nassal et al., 2007; Vorreiter et al., 2007) predicting an HBc-like body of dimeric assembly domains comprising the N-terminal ~180 aa, an ~80 aa CTD with a more profound morphogenetic impact than in HBc, and a much extended ~45 aa proline-rich loop (residues 78–122) connecting helices α3 and α4. However, all attempts to directly verify this model by cryo-EM reconstructions fell short of sufficient resolution. The broader distribution of DHBc vs. HBc CLPs during sedimentation in sucrose gradients and native agarose gel electrophoresis (NAGE) suggested that morphological heterogeneity obscured the data. Some DHBc variants such as R124E produced more homogeneous CLPs (termed class2 Nassal et al., 2007) but did not yield better resolved cryo-EM reconstructions. In particular, the predicted loop at the spike tips remained invisible, in line with complete disorder.

Here we discovered that over several months of storage this loop, now termed extension domain, adopted a defined fold. This enabled us to determine the structure of whole DHBc CLPs at a resolution of 3.7 Å enabling de novo model building. Functional studies revealed that the extension domain´s ability to fold is dispensable for recombinant CLP assembly but correlates with capsid stability in hepatoma cells. We propose that plasticity of the extension domain represents a large-type CP-specific evolutionary adaptation.

DHBV CP (DHBc) expressed in E. coli self-assembles into CLPs which package bacterial RNA (Kenney et al., 1995; Nassal et al., 2007). Here, improved expression was achieved using vectors (Heger-Stevic et al., 2018b) with an E. coli optimized DHBc gene encoding the DHBV16 CP sequence (Uniprot accession: P0C6J7.1). CLPs for cryo-EM analysis were enriched by two sequential ammonium sulfphate precipitations (Birnbaum and Nassal, 1990) followed by sucrose gradient sedimentation (Heger-Stevic et al., 2018a; Figure 1—figure supplement 2).

After two weeks of storage at 4°C we exchanged the sucrose-containing buffer against a cryo-EM compatible buffer with low solute concentration and vitrified the samples. Micrographs showed distinct particles of spherical shape, aggregates of such particles and some proteo-liposomes (Figure 1—figure supplement 2). 2D-class averages revealed particles with protruding spikes but also particles that lacked distinct spikes or were deformed (Figure 1—figure supplement 2), indicating that only a fraction of particles had assembled into regular icosahedral capsids. The resulting 3D-map had a nominal resolution of 6.1 Å and showed capsids with an arrangement of spikes and holes resembling that of HBc-capsids with T = 4 triangulation (Crowther et al., 1994) but with 15 Å broader and 5 Å longer spikes (Figure 1—figure supplement 3). The characteristic broad spikes of the DHBc CLPs were only visible at lower density thresholds but disappeared at higher density thresholds, suggesting a markedly higher structural variability in the extensions than in the core of the spikes.

To test this hypothesis, we analysed the individual asymmetric units by 3D-classification. The class averages (Figure 1—figure supplement 3) showed 42% of asymmetric units without proper spikes and lower average density consistent with dimers either being misfolded, largely displaced or completely missing. Another third of the asymmetric units grouped into classes in which both spikes were broad, suggesting that all extension domains in the asymmetric unit were folded. The remaining 25% of asymmetric units grouped into classes with one narrow and one broad spike, consistent with folded extension domains in the broad spikes and still unfolded or flexible ones in the narrow spikes. Furthermore, the radial position of the spikes in the CLPs varied by up to 7 Å, indicating local deformations of the capsid shell.

The analysis of the individual asymmetric units required accurate determination of the orientations of an ordered asymmetric unit of only some 80 kDa, which is at the lower size limit that can be confidently analysed. We speculated that our analysis might have been limited by the imaging conditions (integrating mode, exposure <40 e/Ų), which were chosen for entire capsids (Song et al., 2019) rather than for the individual asymmetric units. Therefore, we acquired another data set at a higher dose in counting mode (Table 1). For this we prepared new grids of the CLPs that had now been stored in sucrose at 4°C for 10 months.

Much to our surprise 2D-class averages showed crisper projection averages (Figure 1—figure supplement 2) and the 3D-maps of the CLPs were better resolved than before. The 3D-map of the CLPs had a resolution of 3.7 Å (Figure 1, Figure 1—figure supplement 4), but more importantly, the extension domains were clearly visible at the same threshold as the core of the spikes, indicating that during long-term storage most extension domains had become folded. To investigate the folding of individual extension domains, we analysed the asymmetric units as described above. We observed three well resolved classes with broad spikes comprising 74% of the asymmetric units (Figure 1, Figure 1—figure supplement 4), one class with broad spikes and low resolution (9% of the asymmetric units), and one class with a very low average density (18% of the asymmetric units), in line with a still significant number of missing or grossly displaced asymmetric units in the capsids. The three well-resolved classes varied in their radial position by only 2 Å with no major structural differences. Local refinement of the orientations of these particles further improved the overall resolution of the asymmetric unit to a nominal resolution of 3 Å (Figure 1—figure supplement 4) with the extension domains still less well resolved than the rest of the asymmetric unit.

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We were able to build a de novo model of DHBc’s assembly domain into the 3D-map of the asymmetric unit of DHBc (Figure 1, Figure 1—figure supplement 4, Table 2). All four subunits of the asymmetric unit had the same overall fold with adaptations to the different spatial surroundings close to the N-termini and in the C-terminal hand regions (Figure 2a–c). The core of the spikes was formed by four helices, with each DHBc monomer contributing an ascending helix (α3; residues 44–78, 35 aa) and a descending helix (α4; residues 120–157, 38 aa). The ascending helix α3 was kinked in the center between F60 and W61 and tilted away from the dimer axis in the upper part of the spike. The helices were considerably longer than in HBc (Figure 3 and Figure 3—figure supplement 1), where α3 is straight (25 aa) and α4 is kinked (31 aa). The four helices of the spike of HBc and DHBc had a very similar packing below the kink (Figure 3) but diverged above the kink with a different twist of the helices around each other (Figure 3). At the tips of the spikes the positions of the helices were dramatically different, basically swapping the positions of DHBc α3 with HBc α4 in the same subunit and of DHBc α4 with HBc α3 between different subunits of the dimer. Moreover, DHBc helices α3 and α4 were connected through a 41 aa long extension domain (residues 79–119, Figure 2g) whereas in HBc this connection is just a short loop of 3–4 aa. The extension with two short helical segments (αe1: 92–99; αe2: 103–117), was positioned at the side of the spike close to α4 of the same subunit and in contact with α3 of the other subunit (Figure 2e). As independently confirmed below, the packing of the extension domain to the flank of the spike was stabilized by a salt bridge between R124 in α4 of the core spike and E109 in αe2 of the extension domain (Figure 2—figure supplement 1).

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Analysis with Intersurf (Ray et al., 2005) showed that the extension domain increased the inter-dimer surface area from ~1,900 Ų to ~3,400 Ų, highlighting its important contribution to the intra-dimer contact. Nonetheless, the absence of folded extension domains from many spikes in freshly prepared CLPs implied that its contribution is not strictly required for capsid formation and stability, as directly addressed below.

In HBc which lacks an extension domain a disulphide-bridge between Cys61 on opposite monomers (Figure 1—figure supplement 1) can add to dimer stabilization (Nassal et al., 1992). DHBc lacks a homologous Cys residue; instead at a similar position in the spike Phe60 made a π-stacking interaction with Phe60 on the other monomer, and so did His52 with His52 (Figure 2—figure supplement 2). Another difference between HBc and DHBc was the position of the N-termini. In HBc the N-terminus is located at the dimer interface at the base of the spikes and contributes to the intra-dimer contact (Wynne et al., 1999). In DHBc the seven N-terminal residues pointed towards the lumen of the capsid, without contributing to the intra-dimer contact (Figure 2c). Interestingly, the two N-termini within one dimer had different orientations, with those of chains B and C directed towards the base of the spike and those of chains A and D pointing away (Figure 2).

The pockets at the base of the spikes close to the N-termini of chains A and D contained density that was not accounted for by the so far described model (Figure 1e,f magenta). This density resembled a short peptide and was modelled with the 13 very C-terminal residues 250–262. In the other two pockets similar albeit weaker density accounted for the 6 C-terminal residues (257-262). As the connecting residues to the assembly domain were unresolved the C-termini could not be assigned to a specific subunit. Overall, the C-termini appear to take a similar structural role in DHBc as the N-termini do in HBc (Figure 3).

While the intra-dimer contacts differed significantly between HBc and DHBc, the inter-dimer packing in the C-terminal hand-region was largely conserved. The last resolved residues in our model agreed well with the biochemically derived C-terminal border of the DHBc assembly domain around position 187 (Nassal et al., 2007). At the given resolution, the fold of the hand-region was similar to that of HBc (Figure 2d) as it maintained the length of the α5 helix and the distance to the adjacent extended stretch. Overall, the RMSD between HBc and DHBc of the hand-region and the part of the spike below the kink and the fulcrum is 1.9 Å, further highlighting the preservation of the whole shell-region of the capsids (Figure 3—figure supplement 1). Consistent with the preserved inter-dimer contacts, the radius of inner and outer surface of the protein shell and the distance to a diffuse density shell in the interior of the capsid that is attributed to RNA and unresolved parts of the CTDs (Figure 1b) was the same in DHBc and HBc.

The DHBc extension domain is dispensable for recombinant CLP assembly

The low folding propensity of the extension domain questioned a direct role in capsid assembly. We therefore replaced residues 78–122 by a short G₂SG₂ linker and evaluated the consequences on assembly and particle morphology. The corresponding protein, DHBc_Δ78–122, was virtually insoluble; however, solubility was drastically improved by the additional mutation R124E (DHBcR124EΔ). Sedimentation, NAGE and negative staining EM data all indicated the formation of orderly CLPs, prompting a closer look by cryo-EM image (Figure 1—figure supplement 2) reconstruction. The resulting density maps at 3 Å resolution (Figure 4, Figure 4—figure supplement 1, Figure 1—figure supplements 4 and 5) revealed the same architecture as in the wt DHBc particles, except that all spikes remained narrow as also confirmed by the asymmetric analysis (Figure 4—figure supplement 1). Also, the fold of the subunits in the asymmetric unit was nearly superimposable on that of the full length protein subunits with an RMSD of 0.6 Å, except for the missing sideward pointing density. This missing density was accounted for by the 45 amino acids of the deleted extension domain whereas the flexible G₂SG₂ linker was not resolved. Difference mapping confirmed that all density differences between full length and internal deletion variant localized to the extension domain which therefore behaves much like an independent internal appendix.

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Folding competence of the DHBc extension domain is critical for capsid stability in eukaryotic cells and hence viral replication

To reveal a potential virological relevance of the extension domain we examined the impact of extension domain-affecting mutations on capsid expression, viral nucleic acid packaging and reverse transcription in transfected LMH cells, an avian hepatoma line supporting formation of infectious DHBV (Condreay et al., 1990; Dallmeier et al., 2008). Most variants, including the additional mutant R124Q, were tested by trans-complementation of a cloned CP-deficient DHBV16 genome (pCD16_core^-) with the respective DHBc protein expressed from a separate plasmid vector. This excludes inadvertent effects on the virus genome. Mutant E109R_R124E was also tested in the cis-context. Controls included the non-complemented core^- genome and a variant with a mutated YMDD motif in the polymerase active site (pCD16_YMhD), allowing pgRNA encapsidation but not reverse transcription (Radziwill et al., 1990). By NAGE assay of cytoplasmic lysates and subsequent anti-DHBc immunoblotting (Figure 5a) the pCD16 constructs wt, YMhD and E109R_R124E, but not the core-deficient vector, produced similarly intense capsid signals whose mobilities matched those of the recombinant CLPs (lanes 16–19). Comparable signals were generated in the trans-format yet only with wt DHBc and mutants R124K, E109R_R124E and E110R. Variants R124Q, E109R, R124E and i95 gave very weak or no signals at all. Intriguingly, all these variants had exhibited the more homogeneous class2 phenotype when expressed in bacteria. Of the extension-less variants only Δ78–122_R124E produced specific though weak signals (Figure 5—figure supplement 1). To get more insight into the low or lacking capsid signals we repeated the pCD16_core^- cotransfections with variants R124E, Δ78–122_R124E, and E109R_R124E (including at a five-fold reduced plasmid amount) versus wt-DHBc, which reproduced the previously seen NAGE blot data for capsids and capsid-borne DNA (Figure 5—figure supplement 2a,b). This time, however, we additionally addressed non-assembled CP subunits, employing immunoprecipitation (IP) with a polyclonal anti-DHBc antiserum (12/99, Vorreiter et al., 2007) to enrich DHBc-related proteins from lysates of the transfected cells, followed by SDS-PAGE separation of the immunopellets and immunoblotting, again with the polyclonal antiserum; lysate from cells transfected with a GFP expression vector served as control. All DHBc transfected samples produced specific signals with the same mobility as the recombinant full-length proteins (Figure 5—figure supplement 2c); the Δ78–122_R124E band was weak, and thus consistent with the weak NAGE blot capsid signal. Strikingly, however, the R124E sample presented with a whole ladder of smaller products which in sum by far exceeded the left-over full-length band. Hence, the 124E mutation markedly enhances proteolytic susceptibility of the variant and concomitantly impedes its ability to form stable capsids in avian hepatoma cells.

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To reveal capsid-borne viral nucleic acids a parallel NAGE blot was hybridized with a ³²P-labeled DHBV DNA probe, after breaking up the blotted capsids by brief exposure to 0.1 N NaOH; this leaves both RNA and DNA intact. All capsid-positive samples, except the recombinant non-viral RNA containing CLPs, scored positive in this assay (Figure 5b, top). Longer NaOH treatment ablated the signals for sample pCD16_YMhD and reduced it for DHBc_E110R (Figure 5b, bottom), in line with an exclusive and partial RNA content, respectively. Persistence of the other signals indicated they were DNA. A more sensitive Southern blot assay revealed in all these variants a wt-typical pattern of rc-DNA and dsL-DNA (Figure 5c). Such signals were also faintly visible for R124Q and the triple variant E109,110R_R124E but not for variant Δ78–122_R124E. In sum, only variant R124K preserving the positive charge, and the charge-polarity reversing variant E109R_R124E maintained an in all aspects wt-like replication phenotype (Figure 5c and Figure 5—figure supplement 1). Hence opposite charges at positions 109 and 124 and the extension domain´s ability to fold are critical for virus replication.

Despite DHBV's long history as hepadnaviral model virus, knowledge of its structural biology has lagged behind that of HBV. Our EM-map of the DHBc CLP at 3.7 Å and of the DHBc_Δ78–122_R124E CLP at 3 Å resolution allowed us for the first time to build de novo models of DHBc’s assembly domain. Confirming selective aspects of evolutionary conservation between avi- and orthohepadnaviruses (Kenney et al., 1995), the new data also reveal unique structural modules, including a direct contribution of the very C-terminal residues to the capsid shell and a centrally inserted extension of ~45 aa that behaves like an independent folding entity.

A temporarily mobile spike extension - defining feature of hepadnaviruses employing large type CPs?

The most distinctive feature in DHBc vs. HBc are the large extensions at the tips of the spikes that comprise residues 79–119, in line with the biochemical prediction (Nassal et al., 2007) and directly supported by the structure of the extension-less Δ78–122_R124E CLPs (Figure 4). The extension sequence is unusually rich in P (8 of 45 aa, Figure 2) and E residues (6 of 45 aa), the two most disorder-promoting amino acids (Theillet et al., 2013; Uversky, 2013). Notably, P79/P80 and P117/P120 at the borders of the extension domain are ideally located to insulate it from the spike-forming four-helix bundle (Figure 2). These features are preserved in the CPs of diverse other avian hosts (Figure 1—figure supplement 1). Non-avian hosts with large-type CPs such as the 266 aa sequence of the Tibetan frog HBV CP diverge at many positions. Yet, the sequence aligning to the DHBc extension domain also has a very high proline content (10 of 48 aa; Figure 1—figure supplement 1), indicating evolutionary conservation of this feature over millions of years. We therefore expect that the fundamental structural aspects of the DHBV capsid revealed in this study are prototypical for all hepadnaviruses employing the large-type CPs. The plasticity of the extension domain is also reflected by local resolution analysis and high local B-factors in the model (Figure 1—figure supplement 4). Also the spike tips in HBc have high local B-factors (Böttcher and Nassal, 2018), perhaps in both cases facilitating induced-fit interactions, e.g. with the envelope proteins, which for DHBV might be more akin to a folding-by-binding process (Weng and Wang, 2020). Notably, the structure of one HBc spike tip fits well into the DHBc extension domain emerging from the core-spike which provides two such sites per spike (Figure 3—figure supplement 2). Thus, large-type CPs could have evolved two surface protein binding sites per spike in the extension domains, compared to only one in small-type CPs. This coincides with a generally smaller membrane part of the surface proteins in hepadnaviridae with large-type CPs allowing for more surface proteins per area of envelope.

Despite its disposition for disorder, the extension domain in wt DHBc can adopt a stably folded state, albeit very slowly in vitro. The time-scale in live cells is unknown but in sum our data favour that assembly of the capsid shell precedes folding of the extension domains. Assembly can occur in the complete absence of the extension domain, and CLPs with permanently disordered extension domains, such as R124E, present with more homogeneous morphologies than those from wt DHBc. Hence the large contact area to the intra-dimer interface that forms upon folding of the extension domain may even disfavour regular icosahedral assembly; conversely, the requirement for a folding helper would increase the time-span for undisturbed assembly. Thereafter, extension domain folding provides extra stability to the assembled structure. As a resolution of around 7 Å is sufficient to distinguish folded from unfolded extension domains, defining their states in various stages of the viral life-cycle, including in enveloped virions, appears worthwhile and feasible.

Based on the high density of prolines in the extension domain we expect the folding helper to be a peptidyl-prolyl cis-trans isomerase, as is supported by the higher frequency of DHBc CLPs with folded extension domains upon coexpression of E. coli FkpA. From such activity of a bacterial enzyme it appears unlikely that a specialized isomerase is required in DHBV host cells.

In contrast to wt DHBc CLPs, the extension domains in R124E CLPs remained invisible (Figure 4—figure supplement 1), suggesting the homogeneous appearance of class2 CLPs relates to mostly disordered extension domains, as supported by their similarity to the extension-less variant (Figure 4, Figure 4—figure supplement 1).

Intriguingly, despite their efficient expression in bacteria DHBc class2 variants were barely if at all detectable in hepatoma cells (Figure 5); also Δ78–122_R124E gave only faint signals in the NAGE immunoblot (Figure 5—figure supplement 1). As the same eukaryotic vectors led to efficient production of wt DHBc and other class1 particles, the class2 mutations probably interfered with some aspect of protein stability. In fact, decreased CP stability and lack of capsid formation have independently been reported for a fortuitously identified variant lacking H107 (Guo et al., 2006) which we now know to reside within the center of helix αe2. Deletion of H107 thus puts the remaining residues of the helix out of register and, specifically, ablates a hydrophobic interaction of H107 with I99 in αe1 (Figure 2—figure supplement 1) which contributes to the stability of the folded extension domain.Likely, long-term unfolded extension domains enhance accessibility of instability determinants, e.g. target sites for proteases, or for ubiquitylation which initiates proteasomal degradation. This view is strongly supported by the multiple proteolysis products seen with DHBc _R124E (Figure 5—figure supplement 2), and the simultaneous absence of stable capsids and/or nucleocapsids from this variant (Figure 5;.Figure 5—figure supplements 1 and 2). In addition, without the extra contribution of the folded extension domains to the intra-dimer interface the capsid shell may become too labile to withstand the replication process, in particular double-stranded DNA formation. While these are testable hypotheses the lack of stable capsids from these variants precluded statements on a specific role of the extension domain in DHBV replication. Most striking, however, is the complete rescue of wt DHBV-like replication by combining two per se detrimental mutations in the double mutant E109R_R124E (Figure 5), fully supporting our structural model as well as the importance of a salt-bridge between these two specific positions for the ordered state of the extension domain (Figure 6). Conversely, protonation of the glutamic acid carboxylate, realistically achievable at pH values of certain cell compartments (Huotari and Helenius, 2011), would be sufficient to break this bridge and thus create a reversible pH-responsive conformational switch for capsid stabilization and capsid disintegration.

Figure 6

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EM-maps are deposited in the EMDB. Where applicable models were deposited in the pdb DHBc capsid: 10800 (EMDB) 6ygh (pdb) DHBC co expressed with FkpA: 10801 (EMDB) DHBC R124E (mutant): 10802 (EMDB) DHBCR124E_del (deletion-mutant): 10803 (EMDB) 6ygi (pdb).

1. 1. Makbul C
   2. Nassal M
   3. Bottcher B

   (2020) RCSB Protein Data Bank

   ID 6ygh. DHBc capsid.

   https://www.rcsb.org/structure/6ygh
2. 1. Makbul C
   2. Nassal M
   3. Bottcher B

   (2020) RCSB Protein Data Bank

   ID 6ygi. DHBCR124E_del (deletion-mutant).

   https://www.rcsb.org/structure/6ygi
3. 1. Makbul C
   2. Nassal M
   3. Bottcher B

   (2020) Electron Microscopy Data Bank

   ID EMD-10800. DHBc capsid.

   https://www.ebi.ac.uk/pdbe/entry/emdb/EMD-10800
4. 1. Makbul C
   2. Nassal M
   3. Bottcher B

   (2020) Electron Microscopy Data Bank

   ID EMD-10801. DHBC co expressed with FkpA.

   https://www.ebi.ac.uk/pdbe/entry/emdb/EMD-10801
5. 1. Makbul C
   2. Nassal M
   3. Bottcher B

   (2020) Electron Microscopy Data Bank

   ID EMD-10802. DHBC R124E (mutant).

   https://www.ebi.ac.uk/pdbe/entry/emdb/EMD-10802
6. 1. Makbul C
   2. Nassal M
   3. Bottcher B

   (2020) Electron Microscopy Data Bank

   ID EMD-10803. DHBCR124E_del (deletion-mutant).

   https://www.ebi.ac.uk/pdbe/entry/emdb/EMD-10803

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   (1984) Uniprot

   ID P0C6J7. Nucleotide sequence of a cloned duck hepatitis B virus genome: comparison with woodchuck and human hepatitis B virus sequences.

   https://www.uniprot.org/uniprot/P0C6J7
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   (1984) NCBI Nucleotide

   ID K01834.1. Nucleotide sequence of a cloned duck hepatitis B virus genome: comparison with woodchuck and human hepatitis B virus sequences.

   https://www.ncbi.nlm.nih.gov/nuccore/K01834
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   ID AAC41459.1. Escherichia coli and other species of the Enterobacteriaceae encode a protein similar to the family of Mip-like FK506-binding proteins.

   https://www.ncbi.nlm.nih.gov/protein/AAC41459.1
4. 1. Böttcher B
   2. Nassal M

   (2018) RCSB Protein Data Bank

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   https://www.rcsb.org/structure/6HTX
5. 1. Leslie AGW
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   3. Crowther RA

   (1999) RCSB Protein Data Bank

   ID 1qgt. HUMAN HEPATITIS B VIRAL CAPSID (HBCAG).

   https://www.rcsb.org/structure/1QGT
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   (2019) Pfam

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   http://pfam.xfam.org/family/PF00906

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Author details

1. Cihan Makbul

   Julius Maximilian University of Würzburg, Department of Biochemistry and Rudolf Virchow Centre, Würzburg, Germany

   Contribution

   Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing - original draft, Writing - review and editing

   Competing interests

   No competing interests declared

2. Michael Nassal

   University Hospital Freiburg, Internal Medicine 2/Molecular Biology, Freiburg, Germany

   Contribution

   Conceptualization, Resources, Funding acquisition, Validation, Investigation, Visualization, Methodology, Writing - original draft, Project administration, Writing - review and editing

   Contributed equally with

   Bettina Böttcher

   For correspondence

   nassal2@ukl.uni-freiburg.de

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-2204-9158

3. Bettina Böttcher

   Julius Maximilian University of Würzburg, Department of Biochemistry and Rudolf Virchow Centre, Würzburg, Germany

   Contribution

   Conceptualization, Data curation, Formal analysis, Funding acquisition, Validation, Investigation, Visualization, Methodology, Writing - original draft, Project administration, Writing - review and editing

   Contributed equally with

   Michael Nassal

   For correspondence

   bettina.boettcher@uni-wuerzburg.de

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-7962-4849

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