Extensive Mutagenesis of the Conserved Box E ... - Journal of Virology

Extensive Mutagenesis of the Conserved Box E Motif in Duck Hepatitis B Virus P Protein Reveals Multiple Functions in Replication and a Common Structure with the Primer Grip in HIV-1 Reverse Transcriptase Yong-Xiang Wang,a,b Cheng Luo,c Dan Zhao,c Jürgen Beck,a and Michael Nassala University Hospital Freiburg, Internal Medicine II/Molecular Biology, Freiburg, Germanya; Key Laboratory of Medical Molecular Virology, Institute of Medical Microbiology, Shanghai Medical College, Fudan University, Shanghai, Chinab; and State Key Laboratory of Drug Research, Shanghai Institute of Materia Medica, Chinese Academy of Sciences, Shanghai, Chinac

Hepadnaviruses, including the pathogenic hepatitis B virus (HBV), replicate their small DNA genomes through protein-primed reverse transcription, mediated by the terminal protein (TP) domain in their P proteins and an RNA stem-loop, ⑀, on the pregenomic RNA (pgRNA). No direct structural data are available for P proteins, but their reverse transcriptase (RT) domains contain motifs that are conserved in all RTs (box A to box G), implying a similar architecture; however, experimental support for this notion is limited. Exploiting assays available for duck HBV (DHBV) but not the HBV P protein, we assessed the functional consequences of numerous mutations in box E, which forms the DNA primer grip in human immunodeficiency virus type 1 (HIV-1) RT. This substructure coordinates primer 3=-end positioning and RT subdomain movements during the polymerization cycle and is a prime target for nonnucleosidic RT inhibitors (NNRTIs) of HIV-1 RT. Box E was indeed critical for DHBV replication, with the mutations affecting the folding, ⑀ RNA interactions, and polymerase activity of the P protein in a position- and amino acid side chain-dependent fashion similar to that of HIV-1 RT. Structural similarity to HIV-1 RT was underlined by molecular modeling and was confirmed by the replication activity of chimeric P proteins carrying box E, or even box C to box E, from HIV-1 RT. Hence, box E in the DHBV P protein and likely the HBV P protein forms a primer grip-like structure that may provide a new target for anti-HBV NNRTIs.

H

epadnaviruses are small hepatotropic DNA viruses that infect humans and select mammals and birds. Hepatitis B virus (HBV), one of the most relevant viral pathogens of humans (20), is their prototypic member. All hepadnaviruses replicate their ⬃3.0-kb genomes by chaperone-assisted protein-primed reverse transcription (10), executed by their P proteins. These are unusual reverse transcriptases (RTs), which, beyond the common RNA-dependent and DNA-dependent DNA polymerase and RNase H (RH) domains, contain a unique terminal protein (TP) domain at their N termini (Fig. 1A). To initiate reverse transcription, the phenolic OH group of a specific Tyr residue in TP fills the role that conventionally is taken by the 3=-hydroxyl end of a nucleic acid primer (30). P proteins are translated from a greater-than-genome-length transcript, the pregenomic RNA (pgRNA), which also acts as mRNA for the viral core protein. The interaction of the P protein with an RNA stem-loop, ⑀, on the pgRNA is crucial for viral replication; it triggers the coencapsidation of pgRNA and the P protein into newly forming nucleocapsids and the synthesis of a short DNA oligonucleotide, which is templated by the bulge in ⑀ and, via its 5=-terminal nucleotide, becomes covalently attached to the Tyr residue in TP (“protein priming”). Upon transfer to a 3=-proximal acceptor site on pgRNA, the oligonucleotide is extended into fulllength minus-strand DNA, and the pgRNA template is concurrently degraded by the P protein’s RH activity. Some 15 to 18 residues from the RNA 5= end are spared and, upon another template switch, serve as primers for plus-strand DNA. The final product is a capsid-borne relaxed circular DNA (RC-DNA), which still carries P protein covalently bound to the 5= end of the minus strand (reviewed in reference 10).

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Duck HBV (DHBV) has as yet provided the deepest insights into the mechanism of hepadnaviral replication. Beyond providing a feasible in vivo infection system (41, 42), DHBV is the only hepadnavirus for which replication initiation has successfully been reconstituted in vitro. The DHBV P protein in vitro translated in rabbit reticulocyte lysate (RRL) (51) or expressed as a fusion protein in Escherichia coli and supplemented with RRL or purified chaperones (Hsc70 and Hsp40, with further stimulation by Hsp90 and Hop [46]) displays authentic protein-priming activity when supplied with the cognate DHBV ⑀ (D⑀) RNA and deoxynucleoside triphosphates (dNTPs), as manifested by the covalent labeling of the protein if ␣-32P-labeled dNTPs are used (8, 25, 46). A simpler system exploits severely truncated P proteins (miniPs) lacking part of TP, the spacer, the RH domain, and the C-terminal part of the RT domain, which exert chaperone-independent priming activity (7, 8, 12, 52). In any such in vitro assays, the HBV P protein shows, at most, specific binding to its cognate ⑀ RNA (19, 24) but no enzymatic activity. The structural correlates to the diverse activities of hepadnaviral P proteins are not well defined, because it has been impossible

Received 4 January 2012 Accepted 4 April 2012 Published ahead of print 18 April 2012 Address correspondence to Michael Nassal, [email protected]. Supplemental material for this article may be found at http://jvi.asm.org/. Copyright © 2012, American Society for Microbiology. All Rights Reserved. doi:10.1128/JVI.00011-12

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June 2012 Volume 86 Number 12

DNA Primer Grip in DHBV Reverse Transcriptase

FIG 1 Relationship of hepadnaviral P proteins to other reverse transcriptases. (A) Domain structure, relative positions, and primary sequence of the conserved motifs box A to box E in P proteins versus HIV-1 RT. Beyond the RT (DNA polymerase) and RNase H (RH) domains found in all RTs, P proteins contain an extra terminal protein (TP) domain, in which a specific Tyr residue (Y96 for DHBV and Y63 for HBV) acts as an acceptor for the first nucleotide of minus-strand DNA. Numbers are amino acid positions for the DHBV P protein. Of the conserved boxes, box E constitutes the primer grip in HIV-1 RT. The alignment (determined by ClustalW) comprises the sequences in HIV-1 RT and the DHBV and HBV P proteins from boxes A to E; the position of the first amino acid is given in each line; the fourth line (rt no.) refers to HBV P positions in the unified RT numbering system (47). For HIV-1 RT, residues contributing the palm and finger subdomains are shown in green and blue lettering, respectively; the ␤-strands ␤9/␤10 (box C with the YMDD motif) and ␤12/␤13 (box E with the primer grip) are highlighted by yellow boxes. HIV-1 RT sequences transplanted into the DHBV P protein are indicated as chimera 2 and chimera 1. For an extended version of the alignment, see Fig. S1 in the supplemental material. (B) Conservation of box E. Shown is an alignment of box E from HIV-1 RT and Moloney murine leukemia virus (MuLV) RT with the corresponding regions of the P proteins of various hepadnaviruses (WMHBV, woolly monkey HBV; WHV, woodchuck hepatitis virus; GSHV, ground squirrel hepatitis virus; HHBV, heron hepatitis B virus). Residues forming the loop between ␤12 and ␤13 in the HIV-1 RT primer grip are boxed. For comparison, box E positions have been numbered from ⫹1 to ⫹8. (C) The primer grip as a central hub in HIV-1 RT. The primer grip coordinates the relative positions of the palm, finger, and thumb subdomains; its loop interacts with the active-site YMDD motif and with the second-last residue of the primer, thereby properly positioning the primer 3= end toward the incoming dNTP (red pentangle). Green spheres, bivalent metal ions.

to generate sufficient amounts of homogeneous P protein for direct analyses. Due to its importance as a drug target (all five currently approved chemotherapeutics for hepatitis B virus are nucleos[t]idic RT inhibitors [NRTIs] [31]), several models for the RT domain of the HBV P protein have been calculated by using HIV-1 RT as the template (5, 16, 17, 50). The rationale is the presence in the RT domains of all P proteins of short motifs (boxes A to E plus box F [or box II] and box G [or box I] [Fig. 1A]) that


are universally conserved in RTs; for an easier comparison of different HBV isolates, a unified numbering system accounts exclusively for the RT/DNA polymerase domain (47, 61); individual residues are identified as rtXn, where X denotes the specific amino acid and n its position number. In retroviral RTs, the conserved boxes constitute distinct structure elements that together form the catalytic core. Experimental support for the modeled HBV P protein structures comes mainly from the similar locations of muta-

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tions conferring resistance to nucleoside analogs in and around the conserved boxes as those in HIV-1 RT (61) and from cell culture studies in which mutations, e.g., in the YMDD motif (36) of the catalytic center (in box C) and in the proposed helix clamp motif (see Fig. S1 in the supplemental material) downstream of box E (53–55, 60), reduced or abolished viral replication. However, the limited experimental repertoire applicable to the HBV P protein’s function often does not allow one to distinguish whether such inhibitory mutations act specifically or simply induce global P protein misfolding. The DHBV P protein shares about 30% sequence identity as well as susceptibility to several NRTIs with its HBV counterpart but offers broader experimental options. We therefore previously began a more systematic analysis of the functional consequences of mutations in the conserved boxes. For instance, the replacement of F451 (in box A, corresponding to rtF88 in HBV and homologous to Y115 in HIV-1 RT) rendered the DHBV P protein capable of using ribonucleotide triphosphates as substrates, demonstrating the involvement of box A in dNTP versus nucleoside triphosphate (NTP) discrimination, as in HIV-1 RT (11). Here we focus on box E, the most C-proximal motif. Based on various crystal structures of HIV-1 RT-nucleic acid complexes, it has been termed the “DNA primer grip” (48). The sequence forms a hairpin (␤12-␤13) that acts as a connecting hub for all other subdomains (fingers, palm, and thumb); its loop also contacts the second-last residue from the primer 3= end (mostly via M230 and G231) and thus helps in positioning the 3=-terminal OH group close to the catalytic site for a nucleophilic attack on the incoming dNTP (Fig. 1C). Moreover, the primer grip forms part of a binding pocket for nonnucleosidic HIV-1 RT inhibitors (NNRTIs), which cause a misalignment of important components at the polymerase active site and/or prevent the dynamic intersubdomain movements required during polymerization (reviewed in references 38 and 43). Proper geometry can also be disturbed by mutations in the primer grip, explaining the multiple phenotypes reported for such mutants, including impacts on polymerase activity, primer/template utilization (21, 29), dNTP binding (57), the fidelity of DNA synthesis (14, 22, 56), and (via the thumb domain that connects to the RH domain) also RNase H activity (21, 34). Characteristic residues for box E in retroviral RTs and in P proteins from diverse hosts (Fig. 1B) are an aromatic residue at position ⫹3, a hydrophobic residue at position ⫹4, and a GY dipeptide at positions ⫹5 and ⫹6, as exemplified by DHBV P protein residues 556 to 563. Moreover, the box E motif (rt247 to rt254 in the unified HBV RT numbering system) is almost invariably conserved in the ⬎3,000 sequences in the HBV database (http://hivdb.stanford.edu/HBV /DB/cgi-bin/MutPrevByGenotypeRxHBV.cgi). More directly, box E residue Y561 in the DHBV P protein must be close to the active site, because this residue can replace Y96, although less efficiently and only in truncated DHBV miniP, in the in vitro protein-priming reaction (7). Therefore, we used the DHBV P protein as a model to reveal the potential presence in hepadnaviral P proteins of an actual primer grip element that is functionally and structurally related to that in HIV-1 RT. To this end, we extensively mutagenized the box E motif and subjected the variant P proteins to numerous functional assays, from replication in transfected cells to in vitro D⑀ RNA binding. Together with HIV-1 RT homology-based modeling and the functionality of chimeric P proteins carrying box E, or even the sequence encompassing the catalytic YMDD motif in box

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C to box E, from HIV-1 RT, these data demonstrate a common catalytic core architecture of the RT domains of hepadnaviral P proteins and retroviral RTs, in particular the formation of a primer grip-like structure by the box E residues in P proteins. Because the primer grip in HIV-1 RT is the target for multiple approved and experimental NNRTIs (13, 18), the presence of a primer grip equivalent in P proteins suggests that related compounds might be developed into a new class of HBV inhibitors. MATERIALS AND METHODS Cell culture and transfection. LMH cells were cultured in Iscove’s modified Dulbecco’s medium (IMDM; Invitrogen) supplemented with 10% fetal bovine serum, 100 U/ml penicillin G, 100 ␮g/ml streptomycin, and 0.2 mM L-glutamine at 37°C in a humidified atmosphere of 5% CO2. Cells were transfected by using TransIT-LT1 reagent (Mirus), as recommended by the manufacturer. Compounds. Lamivudine (LAM) and phosphonoformic acid (PFA) were purchased from Sigma-Aldrich, and MG132 was purchased from Axxora Platform Biochemicals. Plasmid constructs. All variant DHBV expression vectors were based on plasmid pCD16, which harbors a 1.1-fold DHBV16 genome (GenBank accession no. K01834) under the control of the cytomegalovirus immediate-early (CMV-IE) promoter (33), followed by a T7 promoter in the reverse orientation. Mutations were introduced by standard mutagenic PCR. In the priming-defective variant pCD16-Y96D, the codon for Y96 was changed from TAT to GAT; in the replication-defective variant pCD16-YMHA, the codons for the two essential Asp residues in the YMDD motif were changed to codons for His and Ala (15). In pCD16-Pnull, the P open reading frame (ORF) was prematurely terminated by the replacement of the codon for Y96 with TGA. Box E mutations (amino acids [aa] 556 to 563 in the P ORF) introduced into pCD16 or pCD16Y96D included I556A (ATA⬎GCT), R557A (AGA⬎GCT), F558A (TTC⬎GCT), F558L (TTC⬎CTC), F558W (TTC⬎TGG), L559A (CTC⬎GCT), L559M (CTC⬎ATG), L559I (CTC⬎ATC), G560A (GGT⬎GCT), Y561A (TAC⬎GCT), Q562A (CAG⬎GCT), and I563A (ATT⬎GCT). Expression vectors for the wild-type (wt) and mutant DHBV P proteins with an N-terminal 3⫻ FLAG tag were based on plasmid pcDNA3.1/ Hygro(⫺) (Invitrogen). For the chimeric P proteins (chimeras 1 and 2), the DHBV sequences encoding P protein aa 556 to 563 and 505 to 563, respectively, were replaced by fragments encoding aa 227 to 234 and 177 to 234 from HIV-1 RT (GenBank accession no. AAK08484.2), respectively. Vectors for the in vitro transcription and translation of the DHBV P protein were based on plasmid pT7AMVpol16H6 (6), which encodes a His6 tag between P protein aa 2 and 3. pAAV-Dcore contains the DHBV core ORF between the ClaI and XhoI sites in pAAV-MCS (Stratagene). All HBV vectors were based on plasmid pCH-9/3091, encoding a genotype D, subtype ayw, wt HBV genome (32). All plasmid constructs were confirmed by DNA sequencing. Extraction of capsid-associated viral DNA from transfected cells and Southern blotting. Cells were lysed with NP-40 lysis buffer (10 mM Tris-HCl [pH 7.5], 1 mM EDTA, 50 mM NaCl, and 0.5% Nonidet P-40) and incubated with 10 mM Mg acetate, 100 units/ml DNase I, and 100 ␮g/ml RNase A to remove free nucleic acids. Extraction and detection by Southern blotting of intracellular capsid-associated viral DNA were conducted as described previously (54). Densitometry was performed by using MultiGauge V2.2 software (Fujifilm). RNA encapsidation assay. RNA encapsidation efficiency was determined as reported previously (54). In brief, transfected cells were lysed with NP-40 lysis buffer. Lysates were centrifuged at 12,000 ⫻ g for 2 min, and 40-␮l supernatants were electrophoresed on a native agarose gel and transferred onto a nitrocellulose membrane by using TNE buffer (10 mM Tris-HCl [pH 7.5], 1 mM EDTA, and 50 mM NaCl). Capsids were detected by using anti-DHBV core monoclonal antibody (MAb) 2B9-4F8 (49), followed by peroxidase-conjugated anti-mouse secondary antibody (Jackson ImmunoResearch Laboratories) and ECL reagent (GE Healthcare). For the detection of encapsidated viral RNA, the same membrane

Journal of Virology


was subsequently treated with 0.2 M NaOH–1.5 M NaCl for 30 s, followed by neutralization with 0.2 M Tris-HCl (pH 7.4)–1.5 M NaCl for 5 min. RNAs were then detected by using a 32P-labeled antisense riboprobe. Western blotting. SDS-PAGE and Western blotting were conducted according to standard procedures. Primary antibodies used were antiFLAG MAb M2 or MAbs against tubulin, actin, or lamin (all Sigma); a MAb against green fluorescent protein (GFP) (Roche); and mAb9 against the DHBV P protein (58). For detection, the blots were incubated with peroxidase-conjugated secondary antibodies, followed by ECL⫹ reagent (GE Healthcare). Chemiluminescent signals were recorded on X-ray film or by using a Fuji LAS 3000 instrument. Endogenous polymerase assay. For the enrichment of cytoplasmic nucleocapsids, NP-40 lysates were treated with 10 mM Mg acetate, 100 units/ml DNase I, and 100 ␮g/ml RNase A and then loaded onto a cushion solution (NP-40 lysis buffer plus 20% sucrose). After centrifugation with a TLS55 rotor at 55,000 rpm for 90 min at 20°C, the pellets were suspended in capsid buffer (20 mM Tris-HCl [pH 7.5], 50 mM NaCl, 1 mM EDTA, 0.01% Triton X-100, 0.1% NP-40, and 0.05% ␤-mercaptoethanol) and then incubated overnight at 4°C. The suspension was briefly sonicated and centrifuged at 15,000 ⫻ g for 5 min to remove insoluble material. Cleared supernatants were treated with 15 U micrococcal nuclease and 5 mM CaCl2 and then incubated with reaction buffer containing 50 mM Tris-HCl (pH 7.5), 75 mM NH4Cl, 1 mM EDTA, 10 mM EGTA, 20 mM MgCl2, 0.1% (vol/vol) ␤-mercaptoethanol, 0.5% (vol/vol) Nonidet P-40, 0.4 mM dATP, 0.4 mM dCTP, 0.4 mM dGTP, and 10 ␮Ci [␣-32P]dTTP (3,000 Ci/mmol) at 37°C for various times. Labeled viral DNA was released by the addition of 20 mM EDTA, 0.5% SDS, and 0.5 ␮g/␮l proteinase K and incubation at 50°C for 2 h. After extraction by phenol-chloroform and precipitation by ethanol, labeled DNAs were separated on 1% agarose gels in 0.5⫻ Tris-borate-EDTA (TBE) buffer. Radioactive signals on the dried gels were recorded by using a Typhoon imager and quantitated by using ImageQuant software (both GE Healthcare). DHBV P protein trans-complementation. One microgram of pCD16-P-null plasmid DNA was mixed with either 1 ␮g pCDNA3⫻FLAG-Dpol or its derivative plasmids and then transfected into LMH cells. Four days later, capsid-associated viral DNA was extracted and determined by Southern blotting. In vitro transcription. D⑀ RNA as the template for in vitro priming was obtained from ClaI-linearized pD⑀1 (6), by using the Ampliscribe T7 high-yield transcription kit (Epicentre Biotechnologies). 32P-labeled D⑀ RNA and antisense riboprobes were generated by using the Riboprobe T7 system (Promega) from ClaI-linearized pD⑀1 and NheI-linearized pCD16 plasmids, respectively. D⑀ RNA binding assay. A D⑀ RNA binding assay was performed as previously described (6). In brief, using plasmid pT7AMVpol16H6 as the template, His6-tagged P proteins were in vitro translated in the presence of [35S]Met by using the TNT T7 quick coupled transcription/translation system (Promega). Three microliters of 32P-labeled D⑀ RNA (approximately 8 ⫻ 105 cpm/␮l) was added to the translated products. The mixture was incubated at 30°C for 30 min to allow the formation of P protein-D⑀ RNA complexes. Complexes were immobilized on Ni-nitrilotriacetic acid (NTA) agarose beads (Qiagen) in 300 ␮l binding buffer (0.1 M sodium phosphate [pH 7.5], 150 mM NaCl, 0.1% NP-40, 20 mM imidazole, 100 ␮g/ml yeast total RNA). After shaking for 2 h at 4°C, the beads were washed twice individually with 1 ml binding buffer and 1 ml TMK buffer (50 mM Tris-HCl [pH 7.5], 40 mM KCl, 10 mM MgCl2, 100 ␮g/ml yeast total RNA). Bound complexes were released by the addition of 2⫻ SDS loading buffer (100 mM Tris-HCl [pH 6.8], 4% [wt/vol] SDS, 20% [vol/vol] glycerin, 2% [wt/vol] ␤-mercaptoethanol, and 0.2% [wt/ vol] bromophenol blue) containing 100 mM EDTA and subsequently denatured at 100°C for 5 min. 35S-labeled P protein and 32P-labeled RNA were analyzed by SDS-PAGE, followed by phosphorimaging. In vitro protein-priming assay. In vitro protein priming was performed as described previously (11). In brief, wt and mutant P proteins were in vitro translated in the presence of [35S]Met. Five microliters of the


translation mixture products was analyzed by SDS-PAGE to control translation efficiency. For priming, 10 ␮l of translated products was mixed with 0.6 ␮M D⑀ RNA and 10 ␮Ci [␣-32P]dGTP (3,000 Ci/mmol; PerkinElmer) in a 15-␮l volume containing 10 mM Tris-HCl (pH 7.5), 2 mM MnCl2, and 6 mM MgCl2. The mixture was incubated at 30°C for 1 h and analyzed by SDS-PAGE. After drying, the gels were covered with or without 3 layers of Inkjet films and exposed to a phosphorimage plate to separately monitor the 32P and 35S signals. Molecular modeling. Molecular models of the RT domains of the wt DHBV P protein and chimeras 1 and 2 were constructed similarly to methods reported previously for the HBV P protein (17, 50). BLASTP (1) was used to search for homologs; multiple-sequence alignments were performed by using ClustalW from EBI Tools (http://www.ebi.ac.uk/). Based on the results, chain A from the crystal structure (Protein Data Bank [PDB] accession number 1RTD) of the HIV-1 RT and DNA complex (28) was used as the template to model the three-dimensional (3D) structures of RT domains by using Insight II software (Accelrys, Inc., San Diego, CA). Energy minimizations were performed by using molecular mechanics in the SYBYL software package (version 6.8; Tripos Associates, St. Louis, MO), with the Kollman all-force field. All protein atoms were assigned Kollman all-atom charges, and the energy convergence gradient value of the simulation systems was set to 0.005 kcal · mol⫺1 · Å⫺1. The geometrical reasonability of the modeled structures was tested by a Procheck analysis (http://nihserver.mbi.ucla.edu/SAVS/). DNA and dTTP were superimposed onto the RT structures in Pymol (http://www.pymol .sourceforge.net/), followed by energy minimization to remove unreasonable contacts. Graphic representations were generated by using DS ViewerPro 5.0 (Accelrys). Distances between atoms were calculated by using 3D-Mol Viewer (a module of Vector NTI 8.0; InforMax, Inc.). Drug sensitivity assays. LMH cells were transfected with equal amounts of plasmid pCD16, pCD16-F558A, or pCD16-L559A. Five hours later, different concentrations of LAM or PFA were added to the media. Four days later, cells were lysed, and intracellular capsid-associated viral DNA was determined by Southern blotting. All experiments were performed five times. The 50% inhibitory concentrations (IC50s) for LAM and PFA were calculated by sigmoidal curve fitting, as implemented in Origin 8.0 software (OriginLab, Northampton, MA). Statistical significance was evaluated by a one-way analysis of variance (ANOVA) using Origin 8.0.

RESULTS

The box E motif in the DHBV P protein is crucial for viral replication. Box E in the DHBV P protein comprises residues I556 to I563 (Fig. 1B). To address a potential role in viral replication, all 8 residues were individually changed to alanines. All mutations were transferred into the DHBV expression vector pCD16, which harbors a 1.1-fold wt DHBV16 genome under the control of the CMV-IE enhancer/promoter and produces intact virions upon transfection into the chicken hepatoma cell line LMH (33). Viral DNAs from cytoplasmic nucleocapsids of cells transfected with the mutant genomes and the wt DHBV16 vector as a reference were analyzed by Southern blotting (Fig. 2A). The amounts of viral core protein and P protein, and of cellular ␤-actin, in the respective cytoplasmic lysates were assessed by Western blotting (Fig. 2A). For quantitation, all signals were first normalized to equal loading by using the ␤-actin signals. The signals for viral DNA (RC-DNA plus some double-stranded linear DNA [DLDNA], which is regularly formed as a by-product of reverse transcription) were correlated with the amounts of core protein in the same sample. Mean replication levels and standard deviations (from 5 to 7 experiments for the Ala-scanning mutants and 3 experiments for the other variants) relative to those of wt DHBV, set at 100%, are indicated at the bottom of Fig. 2A; a graphical

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FIG 2 Distinct impact of box E mutations on overall replication and pgRNA encapsidation of DHBV. (A) Relative replication levels. LMH cells were transfected with wt DHBV expression vector pCD16 and its indicated derivatives, and viral DNAs from cytoplasmic nucleocapsids were then analyzed by Southern blotting. RC, relaxed circular DNA; DL, double-stranded linear DNA; M, marker for DL-DNA. Aliquots from the same cell lysates were analyzed by immunoblotting for ␤-actin and the DHBV core protein (Dcore) to normalize for loading and transfection efficiency. The presence of soluble DHBV P protein was monitored by using an anti-DHBV P protein antibody (Dpol). DNA signal intensities were determined by phosphorimaging and, after normalization, were used to calculate replication efficiencies relative to that of pCD16, which was set at 100%. Mean values and standard deviations (SD) were obtained from 5 to 7 experiments (Ala-scanning mutants) and 3 experiments (non-Ala mutants). A graphical representation of interexperiment variation is shown in Fig. S6 in the supplemental material. (B) Relative pgRNA encapsidation efficiencies. LMH cells were transfected with the protein-priming-deficient but pgRNA encapsidation-proficient DHBV vector pCD16-Y96D or its indicated box E derivatives. Vectors for a polymerase-defective DHBV genome (pCD16-YMHA) and for DHBV core protein only (pAAV-Dcore) served as controls. Aliquots from cytoplasmic lysates were subjected to native agarose gel electrophoresis; capsids were detected by immunoblotting, and capsidborne viral RNA was detected by molecular hybridization using a 32P-labeled DHBV-specific riboprobe. RNA signals were quantified by phosphorimaging and normalized to the capsid signals from the same sample to derive relative encapsidation efficiencies. Mean values ⫾ SD are based on data from four independent experiments.

representation of interexperiment variation is shown in Fig. S6 in the supplemental material. Accordingly, the Q562A mutant replicated with wt-like efficiency, and the I556A, R557A, and G560A mutants displayed less than 50% reductions; the lowest replication levels were seen for the F558A, L559A, Y561A, and I563A variants. Hence, at the latter four positions (positions ⫹3, ⫹4, ⫹6, and ⫹8 in box E), the replacement of the bulky hydrophobic side chains by the small methyl groups of Ala had the strongest negative impact on P protein functionality. This position-specific

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pattern of relative functional importance in box E strongly resembles that reported previously for Ala-scanning mutations in the HIV-1 RT primer grip (21), with mutations W229A, M230A, and Y232A but not the flanking residues producing the strongest impairments in DNA polymerase activity. We also analyzed four mutants with more conservative exchanges, namely, the F558W and F558L mutants and the L559M and L559I mutants. In all RTs, the position analogous to DHBV P protein F558 (residue ⫹3 in box E) harbors an aromatic residue (F, Y, or W) but invariably F in hepadnaviral P proteins. Residue ⫹4 (L559 in DHBV) is L in all avian hepadnaviruses and M in the mammalian hepadnaviruses and HIV-1 RT; an I residue here is uncommon but may be selected in HBV (rtM250I) in response to entecavir treatment (50). The F558W exchange had little impact (Fig. 2A), whereas the F558L exchange reduced replication strongly, similarly to the F588A mutation. Hence, an aromatic residue at the ⫹3 position of box E in the P protein appears to be as important as it is in the HIV-1 primer grip. Both the L559M and L559I mutations were much better tolerated than the L559A exchange. Hence, the ⫹4 position of box E exerts a strong preference for a large hydrophobic side chain, again in line with the preference for L and M at this position in the primer grip of retroviral RTs (Fig. 1B). Differential effects of box E mutations on pgRNA packaging. Because the formation of capsid-borne RC-DNA is a multistep process, multiple reasons for the pronounced negative impact on viral replication of some of the box E mutations could be envisaged. Reverse transcription is preceded by the coencapsidation of P protein and pgRNA, which depends on P protein binding to ⑀ and the availability of core protein. We did not see major differences in the levels of core protein (Fig. 2A); potential effects on transcript levels (Fig. 3A) or pgRNA functionality were excluded in subsequent experiments by use of a trans-complementation format (see below). Hence, either the amounts or the properties of the mutant P proteins were responsible for poor replication. To address pgRNA encapsidation competence, we used native agarose gel electrophoresis (NAGE) of intact capsids to compare their relative pgRNA contents; in this assay, capsids can be detected by immunoblotting, and encapsidated viral RNA can be detected by molecular hybridization with a 32P-labeled probe. To prevent a loss of pgRNA due to conversion into DNA, we analyzed the box E mutants in pCD16 vectors encoding a P protein lacking the protein-priming Tyr residue in TP (Y96D), which prevents reverse transcription but not pgRNA encapsidation. Another packaging-proficient but catalytically inactivated variant with an altered YMDD motif (YMHA) and a vector encoding DHBV core protein (pAAV-Dcore) as the sole viral component served as additional controls. As shown in Fig. 2B, cytoplasmic lysates of cells transfected with all viral vectors produced similar amounts of capsids (the low capsid levels from the YMHA variant were specific for this particular sample); enhanced capsid production from plasmid pAAVDcore appears to be an intrinsic feature of this vector (M. Nassal, unpublished data). A visual inspection of the RNA blot (Fig. 2B) already indicated clear reductions in encapsidated pgRNA levels for the Y561A and I563A variants. As expected, no signal was produced by the pAAV-Dcore vector. For semiquantitation, all RNA signals (determined by phosphorimaging) were normalized to the capsid signals from the same sample (by densitometry of the immunoblot signals), which confirmed about equal encapsida-

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FIG 3 Box E nucleotide exchanges reduce replication at the P protein level but not at the pgRNA level. (A) Lack of a significant impact of box E mutations on steady-state levels of viral RNAs. LMH cells were cotransfected with wt DHBV expression vector pCD16 or its indicated derivatives plus a constant amount of a CMV promoter-based GFP expression vector. Mock indicates cells transfected with the GFP vector only. Cytoplasmic RNAs were analyzed by Northern blotting using probes specific for DHBV (top), glyceraldehyde-3-phosphate dehydrogenase (GAPDH) (middle), or GFP (bottom). Relative viral RNA levels were calculated by the normalization of the pgRNA plus preS/S RNA signals in one lane to the GAPDH (loading control) and GFP (transfection efficiency control) signals in the same lane, relative to the pCD16 wt signals, which were set at 100%. The range of relative signal intensities derived from 2 independent experiments is indicated. (B) Low levels of soluble P protein Y561A and I563A variants are due largely to insolubility rather than proteasomal degradation. LMH cells were transfected with pCD16 or its Y561A and I563A derivatives. At 3 days posttransfection, cells were split and cultured for another 24 h in the absence or presence of the proteasome inhibitor MG132. After cell lysis using NP-40 detergent, the soluble and insoluble fractions were separately analyzed by immunoblotting for DHBV P protein (Dpol), DHBV core protein (Dcore), and cellular tubulin (soluble) and lamin (insoluble), respectively. Note the strong Dpol signals for both variants in the insoluble fraction and their near absence in the soluble fraction; MG132 treatment did not change this distribution. (C and D) Poor replication rescue by box E-mutated P proteins in trans. LMH cells were cotransfected with a DHBV vector deficient for P protein production (pCD16-P-null, providing in each reaction mixture the same pgRNA as the encapsidation substrate) plus the indicated pCDNA-based expression vectors for 3⫻ FLAG-tagged versions of the indicated P proteins or plus the empty pCDNA plasmid. Cells transfected with pCD16 served as the control. (C) Intracellular capsid-borne viral DNAs were analyzed by Southern blotting and quantified by phosphorimaging. Replication efficiencies, relative to those of pCD16-P-null complementation by the wt P protein expression vector, which were set at 100%, were calculated from three independent experiments. (D) Capsids and encapsidated DNA in aliquots from the same samples as those shown in panel C were subjected to NAGE. Capsids were monitored by immunoblotting, and encapsidated viral DNA was monitored by molecular hybridization with a 32P-labeled probe.

tion efficiencies for all other variants; the slightly enhanced pgRNA levels per capsid for some variants were not significant. Hence, none of the mutations, save the mutations Y561A and I563A, affected the mutant P proteins’ ability to interact with ⑀ and core protein. Their reduced replication activities (Fig. 2A) therefore likely resulted from impaired DNA polymerase activity. Notably, low-level pgRNA encapsidation by the Y561A and I563A variants correlated with reduced amounts of detectable P protein (Fig. 2A), suggesting that these mutations affected P protein solubility and/or stability. The immunoblot samples had been obtained from the soluble fraction of cell lysates produced by use of NP-40 detergent, which selectively solubilizes the plasma membrane. To distinguish between reduced production, increased proteolytic turnover, or insolubility, we directly compared the P protein contents in the soluble and the insoluble fractions (containing nuclei and insoluble aggregates) of the NP-40 lysates; in addition, the cells were treated, or not, with the proteasome inhibitor MG132 to reduce potential proteasomal degradation (Fig. 3B). DHBV core protein and tubulin or lamin B (for the soluble and insoluble NP-40 fractions, respectively) were assessed as a reference. Consistent with the data shown in Fig. 2A, much less of the mutant P proteins than of the wt protein was detectable in the soluble NP-40 fraction, whereas the reverse was true for the insoluble fraction. Proteasome inhibition for 24 h, starting on day 3 posttransfection, caused no major change in this distribution, except that the overall core protein levels appeared reduced compared to those of the nontreated cells; similar results were ob-


tained after 7 h of MG132 treatment (not shown). We have not investigated whether this reflects a specific effect on the core protein or is due to the known cytotoxic effects of prolonged MG132 treatment. Clearly, however, the P protein Y561A and I563A variants displayed a reduced solubility regardless of the presence or absence of MG132, in line with a negative impact of these mutations on protein folding. Provision of P proteins in trans excludes that altered properties of box E sequence-modified pgRNAs reduce replication. Although not significantly altering pgRNA steady-state levels (Fig. 3A), the nucleotide exchanges required for the box E mutations could still have affected the quality of the modified pgRNAs as encapsidation and replication templates, thereby contributing to the reduced replication of the F558A, L559A, Y561A, and I563A variants (Fig. 2A). To exclude such effects, we trans-complemented a DHBV plasmid defective for P protein translation (pCD16-P-null) with P protein expression vectors encoding 3⫻ FLAG-tagged versions of the wt and the four mutant P proteins. Hence, regardless of the specific P protein, the pgRNA from the pCD16-P-null vector would always be packaged and reverse transcribed. Cytoplasmic lysates of the cotransfected cells were then analyzed by Southern blotting and by NAGE. The wt P protein vector clearly rescued viral replication (Fig. 3C); the lower level of viral DNA than that of the parental pCD16 vector (about 20%) probably reflects the preferential cis-encapsidation of pgRNA molecules from which the P protein is translated (4). Importantly, all four mutant P proteins generated much weaker signals (5 to

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10% of those seen with the wt P protein vector); these data were confirmed by the equally small amounts of viral DNA per capsid seen in the NAGE analysis (Fig. 3D). Hence, the reduced replication levels were attributable to the mutant P proteins and not to altered properties of the packaged pgRNA. Reduced replication efficiency of box E F558A and L559A mutations is due to decreased polymerase activity. To directly confirm an impact of the F558A and L559A mutations on polymerase activity, we monitored the kinetics of recovery of viral DNA formation after treatment with the pyrophosphate analog phosphonoformic acid (PFA). PFA traps HIV-1 RT in the pretranslocated state (39), where the dNTP binding pocket is still occupied by the primer 3= end and thus prevents the access of the next incoming dNTP. PFA also inhibits the P protein’s polymerase activity but not pgRNA encapsidation and protein priming (26). Cells were transfected with pCD16 vectors encoding the two variants or wt DHBV; at 5 h posttransfection, the cells were split and subjected to four different treatments (Fig. 4A). Group 1 was cultured without PFA for another 96 h to serve as untreated controls. The treatment groups received 0.4 mM PFA for 72 h; for group 2, treatment was continued until harvest at 96 h posttransfection; group 3 was cultured for another 7 h; and group 4 was cultured for another 24 h in the absence of PFA. The viral DNA in cytoplasmic nucleocapsids was then analyzed by Southern blotting (Fig. 4B). Untreated cells showed similar relative amounts of viral DNA as before (Fig. 2), with the F558A variant reaching about 20% and the L559A variant reaching about 40% of the level of the wt virus. Permanent PFA treatment reduced DNA levels to nearly undetectable amounts; notably, the signals for the two mutants were slightly stronger than those for wt DHBV, suggesting a low level of resistance to the drug, as confirmed below. In cells allowed to recover for 7 h, signals for wt DHBV already exceeded those for both mutants (group 3) (Fig. 4B), and after 24 h, a similar ratio of the DNA signals of wt DHBV versus the F558A variant versus the L559A variant was restored as in the untreated cells. A graphic representation of the results is shown in Fig. 4C. As both variants encapsidated wt-like amounts of pgRNA (Fig. 2B), the reduced levels of viral DNA must result from reduced polymerase activity. This conclusion was corroborated by using the endogenous polymerase reaction, in which the P protein present in isolated nucleocapsids extends immature replicative intermediates in vitro when provided with dNTPs; by employing at least one ␣-32Plabeled dNTP, the extended products can be visualized directly by autoradiography. Here we subjected nucleocapsids from cells kept for 96 h in the presence of 0.4 mM PFA to this procedure and analyzed the levels of newly formed radiolabeled DNA 1 h, 3 h, 5 h, and 16 h after the addition of dNTP (see Fig. S2 in the supplemental material). As in cells (Fig. 4), the de novo synthesis of 32Plabeled viral DNA in the mutant nucleocapsids proceeded 5 to 10 times slower and less efficiently than in wt nucleocapsids, corroborating a direct impact on polymerase activity. To detect potentially strand-specific impairments in DNA synthesis activity, we analyzed viral DNAs from cells transfected with pCD16 vectors for the F558A and L559A variants by Southern blotting using strand-specific probes (data not shown). The levels of immature minus strands were affected similarly to the corresponding mature strands, and no accumulation of single-stranded minus strands (as expected from a defect in plus-strand synthesis)

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FIG 4 Box E mutations F558A and L559A affect the P protein’s DNA polymerase activity. (A) Treatment schemes for groups 1 to 4. Cells were transfected with pCD16 or its F558A and L559A variants. At 5 h posttransfection, cells received fresh media without PFA (group 1) or with 0.4 mM PFA (groups 2 to 4); 72 h later, PFA was removed for groups 3 and 4, and the cells were grown for another 7 h (group 3) or 24 h (group 4) in the absence of PFA. (B) Viral DNA production. Viral DNAs in cytoplasmic nucleocapsids were analyzed by Southern blotting and quantitated by phosphorimaging. Values were normalized to the amount of DHBV core protein present in the same sample. Permanent PFA treatment reduced the DNA signals by 50- to 200-fold (group 2). A release of the PFA block caused a partial recovery of DNA levels at 7 h (group 3) and a more complete recovery at 24 h post-PFA removal (group 4). (C) Graphic evaluation of DNA levels upon different treatments relative to those of untreated group 1, which were set at 100%. Comparable results were obtained upon the monitoring of de novo DNA synthesis in isolated nucleocapsids by using the endogenous polymerase reaction (see Fig. S2 in the supplemental material).

was observed. Although not excluding minor impacts, these data gave no hints for strand-specific differences. Box E in the DHBV P protein is important for protein priming and ⑀ RNA binding. For in vitro priming, the DHBV P protein is provided with D⑀ RNA as the template plus 32P-labeled dNTPs; the successful formation of the tyrosyl-DNA phosphodiester bond results in the 32P labeling of the protein. The simplest, chaperoneindependent system exploits severely truncated miniP proteins that lack, inter alia, the C-terminal region, which in HIV-1 RT forms the thumb (see Fig. S1 in the supplemental material), such that the region encompassing box E is not stably folded (7). We therefore used the full-length P protein in vitro translated in RRL to assess the impact of box E mutations on protein priming, using [␣-32P]dGTP as the only exogenously added dNTP; dGMP, specified by the C residue at the last position of the D⑀ bulge, constitutes the natural first nucleotide that is added to the protein.

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FIG 5 Differential impact of box E mutations on in vitro priming and D⑀ RNA binding of the DHBV P protein. (A) In vitro priming. The wt P protein, the protein-priming-defective but D⑀ RNA binding-proficient Y96D mutant, and the indicated box E variants were in vitro translated in rabbit reticulocyte lysates in the presence of [35S]Met; a reaction programmed for luciferase (Luc) translation and nonprogrammed lysate (No P) served as the controls. For priming, the reaction mixtures were supplemented with D⑀ RNA and [␣32 P]dGTP in priming buffer, as detailed in Materials and Methods. Primed, 32 P-labeled P proteins were detected by autoradiography after SDS-PAGE. The amounts of soluble P protein present in each reaction mixture were determined by SDS-PAGE and 35S autoradiography of aliquots withdrawn from each translation reaction mixture immediately before adjustment to priming conditions. Relative priming activities were assessed by normalizing the 32P signals to the 35S-labeled P protein signals, with those for the wt P protein set at 100%. Mean values and SD are derived from data from three independent experiments. (B) D⑀ RNA binding. His6-tagged P proteins were in vitro translated in the presence of [35S]Met and then mixed with 32P-labeled D⑀ RNA; one aliquot each was analyzed directly by SDS-PAGE and autoradiography for protein and RNA (panel input). From the rest of each mixture, P protein and bound RNA were captured on Ni-NTA beads. The protein and RNA remaining bound after extensive washing were analyzed by SDS-PAGE/autoradiography as described above for panel A. D⑀ RNA binding activity was assessed by normalizing the 32P-labeled RNA signals to the 35S-labeled P protein signal in the same lane. Mean values relative to those produced by the wt P protein, set at 100%, and SD were derived from data from three independent experiments. Note the significantly enhanced RNA binding by the L559A variant. The complete autoradiograms are shown in Fig. S3 in the supplemental material.

The 35S-labeled in vitro translation products of the F558A, L559A, Y561A, and I563A variants plus the wt P protein as a positive control and the Y96D variant as a negative control were first subjected to centrifugation (at 16,000 ⫻ g for 20 min) to remove insoluble molecules. SDS-PAGE followed by autoradiography showed similar amounts of P protein in all supernatants, including for the Y561A and I563A variants (Fig. 5A). When subjected to priming conditions, all reaction mixtures except those containing no P protein or the priming-deficient Y96D variant generated specific signals but of markedly differing intensities (Fig. 5A). The signal from the L559A variant was about 70% as intense as that from the wt protein, followed by the Y561A and F558A variants (30% to 40% of the wt signal); the signal for the I563A variant was barely above background levels. These results correlated largely with the results of the replication assays shown in Fig. 2A, except that the well-priming L559A variant poorly supported replication (see below for a likely explanation). For the F558W,


F558L, L559M, and L559I variants (Fig. 5A, right), high-level priming activity was seen when F558 was replaced by an aromatic but not an aliphatic residue, and L559 was tolerant to substitutions with other large hydrophobic (I and M) residues, as in the replication assays. This indicated that many of the box E mutations exert their effects already during the initiation of reverse transcription. Finally, we directly assessed the D⑀ RNA binding capacity of the F558A, L559A, Y561A, and I563A variants; the wt P protein and the Y96D variant served as positive controls, and a reaction mixture containing no P served as a negative control. The in vitrotranslated, 35S-labeled, His-tagged proteins were incubated with 32 P-labeled D⑀ RNA, the proteins were immobilized onto Ni-NTA beads (6), and the protein and RNA remaining on the beads after extensive washings were analyzed by SDS-PAGE and autoradiography (Fig. 5B; the complete autoradiograms are shown in Fig. S3 in the supplemental material). For quantitation, the RNA signals were normalized to the full-length P protein 35S signals in the same lane. Expectedly, the wt P protein and the Y96D variant bound similar amounts of D⑀ RNA, as did the F558A variant. Clearly, less RNA was bound by the Y561A and I563A variants, which also had shown low (Y561A) and very low (I563A) levels of priming activity (Fig. 5A). Most notably, the L559A variant reproducibly retained about twice as much RNA as the wt protein, indicating an enhanced affinity for D⑀ RNA. Nonetheless, it produced a weaker priming signal (Fig. 5A), indicating a reduced capacity to use the bound RNA as the template. Box E, and even box C to box E, from HIV-1 RT can functionally replace the corresponding sequences in the DHBV P protein. Together, the results described above suggested that box E in the DHBV P protein is of a similar importance for functionality as the primer grip in HIV-1 RT, implying a similar structure. To substantiate this interpretation, we replaced the box E motif in the P protein (residues I556 to I563) with the primer grip sequence from HIV-1 RT (F227 to L234) (chimera 1), in which the GY dipeptide is the only strictly identical feature (Fig. 1B). More daringly, in chimera 2, the entire ⬃60-aa sequence, encompassing the active-site YMDD motif in box C to box E, is derived from the HIV-1 enzyme (P protein N505 to I563 replaced by HIV-1 RT D177 to L237) (Fig. 1A). Functional activity would be expected only if both sequences, despite their divergent primary sequences (15 identical plus 11 similar residues within 59 residues total), could adopt similar 3D structures. To avoid impacts on pgRNA by the numerous nucleotide exchanges, functionality was assessed in the trans-complementation format. The cotransfection of pCD16-P-null with the wt P protein expression vector and the empty pCDNA plasmid served as the control. As shown in Fig. 6, both chimeric P proteins supported the formation of the same pattern of RC-DNA plus DL-DNA as the trans-complemented wt P protein albeit with a lower efficiency (around 30% of the wt). This was partly due to reduced levels of soluble chimeric P proteins (Fig. 6), but additional contributions of the numerous mutations to reduced polymerase activity are likely. Most importantly, however, replacement by the short and even the long HIV-1 RT sequences still rendered the chimeric proteins capable of fulfilling all P protein functions, including pgRNA encapsidation and proper protein-primed reverse transcription. Compared to the strong negative effects that even some single box E mutations exerted on DHBV P protein function, this high level of tolerance toward multiple mutations derived from the homologous

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FIG 6 Replacement of box E, or box C to E, by the corresponding HIV-1 RT elements yields functional P proteins. Cells were cotransfected with pCD16P-null plus pCDNA vectors encoding the 3⫻ FLAG-tagged wt DHBV P protein or its chimera 1 and 2 derivatives (Fig. 1A). Cotransfection with an empty pCDNA vector and transfection with the wt DHBV expression plasmid pCD16 served as controls. In chimera 1, box E residues 556 to 563 were replaced by HIV-1 RT residues 227 to 234; in chimera 2, residues 505 to 563 (box C to E) were replaced by HIV-1 RT residues 177 to 234. Viral DNA from cytoplasmic nucleocapsids was analyzed by Southern blotting, and levels of DHBV core protein and tubulin were monitored by Western blotting. The presence of soluble P proteins was addressed by using anti-FLAG and anti-DHBV P protein antibodies. Replication capacities relative to those of the trans-complemented wt P protein were calculated from the DNA signal intensities after normalization to tubulin and core protein levels in the same sample; means ⫾ SD are from three independent experiments.

HIV-1 RT sequence strongly supports the formation of similar 3D structures by both protein sequences. Molecular modeling is consistent with a primer grip-like structure of box E in the DHBV P protein. The modeled structures of the RT domain of the HBV P protein have been derived by using HIV-1 RT as the template (5, 16, 17). Here we took a similar approach for the DHBV P protein and its two chimeric derivatives, as described in Materials and Methods. In brief, P protein residues 382 to 657, and the corresponding regions of chimeras 1 and 2, were modeled on residues 53 to 314 from chain A of the crystal structure of HIV-1 RT complexed with DNA and dTTP (PDB accession number 1RTD [28]); that structure displays the typical right-hand architecture with palm, fingers, and thumb (Fig. 7A). DNA and dTTP were then superimposed onto the energy-minimized model structures. The DHBV sequence adopted an architecture that was similar overall to that of HIV-1 RT (Fig. 7B), and the box E residues formed a hairpin structure (Fig. 7F), as in the HIV-1 RT primer grip (Fig. 7E), around which the subdomains, including the YMDD active-site motif, and the 3=-proximal primer strand residues organize. Residues L559 and G560 were situated at the tip of the primer grip hairpin (Fig. 7F), like HIV-1 RT residues M230 and G231 (residues ⫹4 and ⫹5) (Fig. 7E) and, thus, were similarly able to contact the penultimate primer residue and properly position the terminal 3=-OH relative to the incoming dNTP. These interactions would be perturbed by the replacement of either residue with Ala. F558, at the ⫹3 position, contacted Y511 in the YMDD motif (highlighted in purple in Fig. 7E to H), an arrangement that was overall similar although not identical to that of HIV-1 RT residues W229/M230 (in the primer grip) and Y183 (in the YMDD motif); there, M230, rather than W229, is closest to

FIG 7 Molecular modeling supports the formation of a primer grip-like structure by box E in the DHBV P protein. (A to D) Overall structures of the RT domains plus the bound template/primer duplex (in blue) and dTTP (in red) from HIV-1 RT (PDB accession number 1RTD) and modeled structures of the wt DHBV P protein (Dpol) and its chimeric derivatives in which box E (chimera 1) or box C to box E (chimera 2) were replaced by the corresponding HIV-1 RT elements. The primer grip hairpin is shown in yellow, template and primer strands are shown in blue, and bound dTTP is shown in red. The red squares indicate the parts of the structures that are enlarged in panels E to H. (E to H) Close-up views of the interactions between the active-site YMDD motif (in magenta, with Y183 in HIV-1 RT and Y511 in the P protein shown as stick models) and the primer grip loop. The residue numbering in the primer grip is described in the legend of Fig. 1B. As in the HIV-1 enzyme, primer grip residues ⫹3, ⫹4, and ⫹5 are in close proximity to the penultimate primer residue and the Y residue in the YMDD motif; however, the major interaction with the latter is with the position ⫹3 residue F558 rather than with the position ⫹4 residue as in HIV-RT (M230); in the chimeric proteins, a more HIV-1 RT-like arrangement appears to be restored.

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Y183. The replacement of F558 by smaller residues would weaken the interaction with the YMDD motif, consistent with the reduced replication capacity of the F558A variant, and would possibly affect resistance to NAs (a modest hypersensitivity toward LAM was indeed observed [see below]). The side chains of Y561 and I563 displayed extensive hydrophobic interactions with residues of the palm and thumb subdomains (L432, I570, and F609, and L432, I435, I556, and I570) (see Fig. S4 in the supplemental material), which would not be maintained by the smaller Ala side chain; the implied impact on overall folding is in line with partial misfolding and the observed insolubility. The ⫹7 position (Q562 in DHBV P) shows substantial variability between different RTs (Fig. 1B), and thus, the negligible impact of its replacement by Ala in the DHBV P protein is consistent with a nonessential role in folding and activity. The modeling of the two chimeric DHBV P protein RT domains resulted in a similar overall architecture, with some alterations in the thumb domain (Fig. 7C and D). The primer grip hairpin was also formed, yet the contact to Y511 in the YMDD motif was mediated largely by M559 (position ⫹4) rather than W558 (Fig. 7G and H), very similar to HIV-1 RT. Together, these and other (7) modeling studies make a common structure of the box E motifs in the DHBV P protein and HIV-1 RT highly plausible. F558A and L559A in the DHBV P protein inversely affect sensitivity to lamivudine and PFA. As a final, independent line of evidence supporting such a similarity, we analyzed the impact of the box E mutations F558A and L559A in the DHBV P protein on sensitivity to lamivudine and PFA. In HIV-1 RT, the primer grip does not directly contact the incoming dNTP, but due to its scaffolding function, mutations can indirectly affect susceptibility to nucleoside analogs. For instance, the box E mutation F227A rendered the enzyme 8-fold hypersensitive to azidothymidine (AZT), and various HIV-1 RT mutants selected for resistance to PFA increased sensitivity to AZT (23, 44). Both DHBV P protein mutations induced an about 3-fold, statistically significant resistance to PFA (mean IC50s of 189 ␮M and 143 ␮M, versus 55 ␮M for wt DHBV), whereas sensitivity to lamivudine was about 2-fold enhanced (IC50s of 3.4 and 2.5 ␮M, versus 6.8 ␮M for wt DHBV) (see Fig. S5 in the supplemental material for representative titration experiments). For HBV, a similar cross talk between box E and the dNTP binding pocket in the HBV P protein was suggested by the changes in sensitivity to lamivudine and entecavir reported previously for mutations of the box E ⫹4 residue rtM250 (3, 50); for instance, rtM250V (in the absence of other mutations) rendered the HBV enzyme 7-fold hypersensitive to LAM (50). Hence, these data are again consistent with a common structural role for the box E motif in hepadnaviral P proteins and the DNA primer grip in HIV-1 RT. Similar impacts on replication of box E mutations in human HBV and DHBV P proteins. DHBV and HBV are more closely related to each other than is either of them to HIV-1. Hence, if box E in the DHBV P protein can adopt a primer grip-like structure, this should also hold for the HBV protein, as also implied by previously reported HBV RT models (5, 16, 17, 50). Mutations of box E in HBV P protein should then have functional consequences similar to those for DHBV. We therefore replaced residues F584, M585 (rtM250), Y587, and I589 in the HBV P protein (box E positions ⫹3, ⫹4, ⫹6, and ⫹8, corresponding to F558, L559, Y561, and L563 in DHBV, respectively) with Ala and compared


FIG 8 Box E mutations in the HBV P protein generate replication phenotypes similar to those of the DHBV P protein. (A) Southern blot of HBV DNA from cytoplasmic nucleocapsids. Human Huh7 cells were transfected with the wildtype HBV expression vector pCH-9/3091 (pCH-wt) or derivatives carrying the indicated P protein mutations. For easier comparison, the box E positions of the mutated residues are indicated on the top; homologous residues in HBV versus DHBV are F584/F558, M585 (rtM250)/L559, Y587/Y561, and I589/ I563. Relative replication efficiencies were calculated from the DNA signals relative to the amounts of HBV core protein (Hcore) and tubulin present in the same samples; means ⫾ SD are from three independent experiments. ss, single-stranded DNA. (B) Relative viral DNA contents of cytoplasmic nucleocapsids. Equal aliquots from the cytoplasmic lysates were analyzed by NAGE, followed by the immunodetection of HBV capsids and molecular hybridization of packaged DNA with a 32P-labeled HBV probe. The relative DNA content per capsid was calculated from the intensity of the DNA signals normalized to the capsid signals from the same sample. (C) Relative viral RNA contents of cytoplasmic nucleocapsids. Huh7 cells transfected with the same vectors as those described above (A) were cultured in the presence of 250 ␮M LAM to prevent the conversion of packaged pgRNA into DNA. Aliquots from cytoplasmic lysates were subjected to NAGE and processed as described above (B). RNA contents per capsid relative to those of wild-type vector-transfected cells (100%) are derived from three experiments.

the replication competence and pgRNA encapsidation capacity of the corresponding virus variants to those of wt HBV upon transfection into the human hepatoma cell line Huh7 (Fig. 8A). Indeed, all mutations reduced replication and RNA encapsidation efficiency without detectably influencing capsid levels (Fig. 8B and C). Relative reductions were similar to those of DHBV for the ⫹6 position and were more pronounced for the ⫹8 position and less pronounced for the ⫹3 and ⫹4 positions than in DHBV (F558 and L559); a similarly minor effect of the rtM250A mutation at position ⫹3 was also seen previously in the context of a different HBV genotype, where the corresponding residue is M598 (3). Apart from the limitations in the accurate quantitation of Southern blots and immunoblots (see Fig. S6 in the supplemental material for the DHBV mutants), the gradual differences in the mutational impact on HBV and DHBV likely reflect the substantial sequence divergence between the two proteins. However, compared with the results reported for mutations at position ⫹3 other than M598A (3), the trend for DHBV L559 mutations was very similar in that larger aliphatic amino acids were better tolerated than smaller ones. Hence, together, the data are fully compatible with a primer grip-like structure of the box E residues in HBV and DHBV P proteins. DISCUSSION

Despite the importance of HBV as a pathogen, structural information on hepadnaviral replication is limited to homology-based

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models of the DNA polymerase (RT) domain of the HBV P protein (5, 16, 17, 50). While consistent with NRTI resistance data, HBV provides few experimental opportunities to directly validate the structural predictions. Using the related DHBV P protein as a model, here we extended our previous functional analysis (11) to the conserved box E, which forms the DNA primer grip in HIV-1 RT. All our experimental data plus molecular modeling strongly suggest that box E in the DHBV P protein as well as the HBV P protein adopts a structure similar to that of the primer grip in HIV-1 RT, with a similarly central role in coordinating the interactions between RT subdomains and the primer 3= end. Most compellingly, this is indicated by the replication competence of chimeric DHBV P proteins carrying box E, or the entire sequence from box C to box E, from HIV-1 RT. Our data provide new insights into the replication mechanism of hepadnaviruses that should also be useful for the design of new nonnucleosidic antiHBV compounds. Specific versus global effects of box E mutations of the DHBV P protein: evidence for a primer grip. Most box E mutations in the DHBV P protein, unless in chemically similar residues, reduced the overall replication efficiency (Fig. 2A); such negative effects could be specific or may simply reflect global misfolding. For distinction, we tested the impacts of the mutations on P protein solubility and stability, and we subjected the variants to multiple functional assays, reflecting the consecutive steps of hepadnaviral reverse transcription. The soluble fraction of the NP-40 lysates should contain P protein encapsidated in cytoplasmic nucleocapsids plus free, nonaggregated molecules; molecules that were aggregated and/or associated with nuclei, large cellular complexes, or debris should accumulate in the insoluble fraction. In transfected cells, the fraction of nonencapsidated P protein exceeded that of encapsidated molecules, and most of those apparently precipitated (59). We did not differentiate between capsid-borne and free soluble P protein, yet clearly, the Y561A and I563A variants produced much less soluble protein than did the wt P protein and the other variants (Fig. 2A and 3B), suggesting a folding defect. In all viral RTs, the corresponding primer grip positions ⫹6 and ⫹8 are occupied by an aromatic residue and a large hydrophobic residue (Fig. 1B), which undergo extensive hydrophobic interactions with the palm and thumb subdomains, and the same was predicted in our modeled DHBV RT domain (see Fig. S4 in the supplemental material). The lack of these interactions in the Y561A and I563A mutants could therefore easily promote misfolding and insolubility; notably, an L234A exchange (equivalent to I563A) in HIV-1 RT also affected proper folding (29). In turn, the lack of soluble P protein largely explains the reduced replication capacities of these two variants. Insolubility in the RRL in vitro translation system was less pronounced, perhaps because only freshly translated protein was analyzed. However, the low levels of D⑀ RNA binding and in vitro priming displayed by the two variants (Fig. 5) suggest that even the RRL-soluble molecules are functionally impaired. In sum, an impact of the Y561A and I563A mutations on folding is compatible with the predicted primer grip structure, but more specific conclusions can be drawn from the other variants, which did not display obvious folding problems. Implications for the hepadnaviral replication mechanism. While protein priming is unique to hepadnaviruses, the subsequent chain elongation reactions must proceed as with other reverse transcriptases. Mutations in the primer grip of HIV-1 RT,

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depending on their nature and relative position, can severely impair polymerase activity (14, 21, 22, 29, 34, 56, 57). Due to the central position of the primer grip, such an impairment can occur by various mechanisms; these may be summarized as misalignments of the components involved in catalysis and/or a hindrance of the dynamic structure changes (“ratcheting”) that accompany the cycling of the enzyme through successive polymerization steps (43). A comprehensive mechanistic analysis was beyond the scope of this study, but the data still allow for several conclusions of relevance for the hepadnaviral replication mechanism. Box E variants other than the Y561A and I563A variants displayed low-replication phenotypes despite the absence of obvious solubility or pgRNA packaging deficits (Fig. 2). We first confirmed a direct effect on the P protein’s polymerase activity by monitoring the kinetics of DNA synthesis after the release of a block by PFA. In both intact cells (Fig. 4) as well as isolated nucleocapsids (see Fig. S2 in the supplemental material), the F558A and L559A variants generated less DNA from the encapsidated pgRNA than the did the wt protein, and they did so with slower kinetics. In HIV-1 RT, the corresponding box E positions ⫹3 and ⫹4, W229 and M230, and the universally conserved G at position ⫹5 form the loop of the ␤12-␤13 primer grip hairpin and mediate the contacts to the Y residue in the YMDD motif and the ⫺2 residue in the primer (Fig. 1C). In our modeled DHBV P protein RT domain, F558 and L559 are located at equivalent positions (Fig. 7F). The tolerance of the polymerase activity to replacement by the HIV-1-specific W and M residues (F558W and L559M) but not by Ala with its small side chain (F558A and L559A) is fully consistent with this model. Similarly, the replacement of the universally conserved glycine at position ⫹5 by a more bulky amino acid would affect the interaction with the second-last primer residue and the proper positioning of the primer 3= end. Indeed, the G560A mutant displayed a modestly reduced replication competence despite the maintained pgRNA encapsidation efficiency (Fig. 2). Thus, all these data are in line with box E of the P protein forming a primer grip structure with a function during polymerization homologous to that in other RTs. Perhaps the most compelling evidence for a common catalytic core structure comes from the ability of the two chimeric P proteins to support DHBV replication, with the formation of the same replication products as those formed by the genuine DHBV P protein (Fig. 6). Hence, despite the replacement of box E, or even the entire sequence from box C containing the catalytic YMDD motif to box E, by the divergent sequence from the HIV-1 enzyme, the chimeric proteins were able to specifically package pgRNA and to initiate and complete reverse transcription, including RNase H-mediated pgRNA degradation. Thus, the unique ⑀ RNA and TP-dependent steps of hepadnaviral replication also proceeded normally although with decreased efficiency. Notably, the reduction in the overall replication level was not more pronounced than that caused by the single F558A and L559A mutations in the genuine P protein context. Together with the importance of the YMDD motif in P proteins and our previous demonstration that DHBV P protein F451 in box A fulfills a role in dNTP versus NTP discrimination analogous to that of Y115 in HIV-1 RT, our new data firmly establish a common catalytic core structure in the RT domains of P proteins and retroviral reverse transcriptases; in turn, this strongly supports the plausibility of the modeled P protein RT domain structures.

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Role for the primer grip in protein priming. Regarding the periphery of the RT domain, the reliability of the predicted P protein structures is limited by the low sequence similarity to the HIV-1 RT template outside the conserved boxes. For the lack of known homologues, homology modeling is not at all applicable to the hepadnavirus TP domain; direct structural analyses of P proteins, including TP, therefore remain a highly desirable goal. Still, our in vitro priming data with the box E variants also provide new insights into hepadnaviral replication initiation. Priming as well as pgRNA encapsidation depend on P protein binding to the ⑀ RNA stem-loop. Many of the box E variants supported efficient pgRNA encapsidation similarly to the wt enzyme, suggesting the absence of global misfolding, yet they converted the RNA into DNA less efficiently (Fig. 2). This can be attributed in part to less efficient DNA elongation (Fig. 4, and see Fig. S2 in the supplemental material), but the in vitro priming data suggest that box E mutations can also affect TP-dependent protein priming. Most evident was the strict correlation between the in vitro priming activity and the overall replication capacity of most variants. An interpretation of the data for the Y561A and I563A variants is not straightforward due to the solubility issues related to these mutations (see above). However, replacements of F558 by Ala and Leu but not Trp reduced both activities; priming, like replication, was more tolerant to the mutations L559I and L559M than to the mutation L559A, and the F558A variant performed more poorly than did the L559A variant in either assay (Fig. 5 versus Fig. 2A). Hence, the addition of the first nucleotide to the protein-priming Tyr residue in TP apparently has requirements very similar to those of the subsequent strand elongations. This is in accord with previous data: active-site mutations such as YMHD or YMHA prevent polymerization as well as priming, and, as in elongation, the addition of the first nucleotide to TP occurs in a templated fashion (7). Hence, the templating nucleotide in the ⑀ RNA bulge must be properly positioned so as to base pair with the bound dNTP, and the priming tyrosyl-OH group must occupy the same position in the catalytic center as the 3= end of a nucleic acid primer (see Fig. 9 in reference 7). By analogy, the primer grip might be involved in Tyr positioning by contacting nearby TP residues. Alternatively, the primer grip mutations could more indirectly distort the geometry of the initiation complex because, just as during elongation, all components must be properly aligned. Information on how the ⑀ RNA interacts with the P protein during priming is limited. The regions around the bulge (which serves as the template) and the apical loop are particularly important for P protein binding (7, 40, 41), and once the RNA is bound, the unfolding of the upper stem is critical for its template function (9). On the P protein, one RNA binding site appears to comprise the conserved T3 motif in the C-terminal TP region (2, 45) and the so-called RT-1 motif (2) in the N-terminal RT domain part (residues 381 to 416), far apart in primary sequence from box E. Most notable in this light is the enhanced D⑀ RNA binding yet reduced priming activity observed for the L559A variant (Fig. 5), a phenotype previously observed for mutations in the RNA rather than in the protein (27). Although the effects were only 2-fold statistically significant, they indicate that the primer grip contributes to ⑀ RNA binding as well as its utilization as the template. Two, not mutually exclusive, options are that structural changes in the mutant protein favor nonproductive interactions with the RNA, e.g., by im-


peding its rearrangement into the template-active conformation, or that the RNA can rearrange but that the distortions in the catalytic core of the protein reduce its efficient utilization as the template. Nuclease accessibility experiments comparing the D⑀ RNA bound to the mutant versus the wt P protein (9) may help to distinguish between these possibilities. The primer grip in the P protein as an antiviral target? As for HIV-1 RT, we found that box E mutations in the DHBV P protein had modest, differential impacts on sensitivity to the nucleoside analog lamivudine and the pyrophosphate analog PFA (see Fig. S5 in the supplemental material), supporting cross talk between box E and the dNTP binding pocket, as in the HIV-1 enzyme. Such cross talk is also implied for the HBV P protein, where mutations of residue rtM250 (equivalent to box E residue L559) can differentially affect resistance to entecavir and lamivudine (3, 50). While independently corroborating an architecture similar to that of HIV-1 RT, a practically more important implication of the presence of a primer grip in the DHBV P protein and most likely also in the HBV P protein is that it may provide a new target for nonnucleosidic RT inhibitors. NNRTIs are important components of highly active antiretroviral therapy (HAART) against HIV-1, which distort the geometry of the catalytic core, including by dislodging the primer grip (43). In HIV-1 RT, the primer grip forms one side wall of the NNRTI binding pocket, together with residues immediately upstream of box A, the box C ␤9␤10 hairpin presenting the YMDD motif, and some others. Our data predict that hepadnaviral P proteins contain a similar pocket that may provide a valuable new target for HBV antiviral agents. Given that most NNRTIs against HIV-1 RT are not active against HIV-2 RT (37), it is unlikely that the already established anti-HIV-1 drugs will be active against the more distantly related HBV P protein. However, by the screening of compound libraries that maintain the principal HIV-1 NNRTI butterfly-like shape (13, 35) and by advanced modeling, NNRTIs with activity against HBV should be identifiable. Because all current anti-HBV therapies beyond type I interferon rely exclusively on nucleos(t)ide analogs, such NNRTIs, with their different mechanism of action, would be a most desired alternative for, or complement to, the limited drug repertoire available against chronic hepatitis B. ACKNOWLEDGMENTS This work was supported in part by the Shanghai Education and Development Foundation (grant KBF101038), the National Natural Science Foundation of China (grant 30800048), and the Deutsche Forschungsgemeinschaft (grant DFG Na154/7-3). Y.X.W. is indebted to the Alexander von Humboldt Foundation for a postdoctoral fellowship. We thank John E. Tavis for providing the anti-DHBV P protein antibody mAb9.

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