
==== Front
Nucleic Acids Res
Nucleic Acids Res
nar
Nucleic Acids Research
0305-1048
1362-4962
Oxford University Press

39189451
10.1093/nar/gkae739
gkae739
AcademicSubjects/SCI00010
Structural Biology
Cryo-EM structure of DNA polymerase of African swine fever virus
Kuai Lu CAS Key Laboratory of Pathogen Microbiology and Immunology, Institute of Microbiology, Chinese Academy of Sciences, Beijing 100101, China
Medical School, University of Chinese Academy of Sciences, Beijing 100049, China
Beijing Life Science Academy, Beijing 102209, China

Sun Junqing College of Veterinary Medicine, Shanxi Agricultural University, Jinzhong, 030801, China

Peng Qi CAS Key Laboratory of Pathogen Microbiology and Immunology, Institute of Microbiology, Chinese Academy of Sciences, Beijing 100101, China
Beijing Life Science Academy, Beijing 102209, China

Zhao Xuejin CAS Key Laboratory of Pathogen Microbiology and Immunology, Institute of Microbiology, Chinese Academy of Sciences, Beijing 100101, China
Beijing Life Science Academy, Beijing 102209, China

Yuan Bin CAS Key Laboratory of Pathogen Microbiology and Immunology, Institute of Microbiology, Chinese Academy of Sciences, Beijing 100101, China

Liu Sheng Shenzhen Children's Hospital, Shenzhen 518038, China

Bi Yuhai CAS Key Laboratory of Pathogen Microbiology and Immunology, Institute of Microbiology, Chinese Academy of Sciences, Beijing 100101, China
Medical School, University of Chinese Academy of Sciences, Beijing 100049, China

https://orcid.org/0000-0002-3053-2687
Shi Yi CAS Key Laboratory of Pathogen Microbiology and Immunology, Institute of Microbiology, Chinese Academy of Sciences, Beijing 100101, China
Medical School, University of Chinese Academy of Sciences, Beijing 100049, China
Beijing Life Science Academy, Beijing 102209, China

To whom correspondence should be addressed. Tel: +86 10 64807806; Email: shiyi@im.ac.cn
The first four authors should be regarded as Joint First Authors.

23 9 2024
27 8 2024
27 8 2024
52 17 1071710729
19 8 2024
7 8 2024
25 4 2024
© The Author(s) 2024. Published by Oxford University Press on behalf of Nucleic Acids Research.
2024
https://creativecommons.org/licenses/by-nc/4.0/ This is an Open Access article distributed under the terms of the Creative Commons Attribution-NonCommercial License (https://creativecommons.org/licenses/by-nc/4.0/), which permits non-commercial re-use, distribution, and reproduction in any medium, provided the original work is properly cited. For commercial re-use, please contact journals.permissions@oup.com

Abstract

African swine fever virus (ASFV) is one of the most important causative agents of animal diseases and can cause highly fatal diseases in swine. ASFV DNA polymerase (DNAPol) is responsible for genome replication and highly conserved in all viral genotypes showing an ideal target for drug development. Here, we systematically determined the structures of ASFV DNAPol in apo, replicating and editing states. Structural analysis revealed that ASFV DNAPol had a classical right-handed structure and showed the highest similarity to the structure of human polymerase delta. Intriguingly, ASFV DNAPol has a much longer fingers subdomain, and the thumb and palm subdomain form a unique interaction that has never been seen. Mutagenesis work revealed that the loss of this unique interaction decreased the enzymatic activity. We also found that the β-hairpin of ASFV DNAPol is located below the template strand in the editing state, which is different from the editing structures of other known B family DNAPols with the β-hairpin above the template strand. It suggests that B family DNAPols have evolved two ways to facilitate the dsDNA unwinding during the transition from replicating into editing state. These findings figured out the working mechanism of ASFV DNAPol and will provide a critical structural basis for the development of antiviral drugs.

Graphical Abstract

Graphical Abstract

National Key R&D Program of China 10.13039/501100012166 2021YFC2300200 2021YFC2300700 National Natural Science Foundation of China 10.13039/501100001809 81871658 32192452 32100119 32100129
==== Body
pmcIntroduction

African swine fever (ASF), caused by the African swine fever virus (ASFV), is a severe and highly contagious animal disease, with almost 100% mortality in wild boars and domestic pigs (1). ASF has existed for more than 100 years, and was first reported in Kenya in 1921 (2). Since then, it has spread to many countries in Europe, South America, and Asia, becoming a global threat to the pig industry (2,3). In August 2018, ASF outbreak was firstly reported in China (4), the largest pork producer in the world, and now ASF has spread to other Southeast Asia countries, including the Philippines and Thailand (5,6). Due to the lack of effective vaccines and drugs, the primary countermeasures against ASF rely on culling the pigs in infected farms. Therefore, the disease has caused significant economic loss in many countries. According to an evaluation report, the direct costs of ASF in China and neighboring countries could be as much as $130 billion (http://dx.doi.org/10.22617/WPS200263-2).

Arthropod-borne viruses present a substantial threat to human and animal health worldwide (7,8). ASFV belongs to the family Asfarviridae, genus Asfivirus, and is the only known arthropod-borne DNA virus within the superfamily of the nucleocytoplasmic large DNA viruses (NCLDVs) with a virion diameter of 260–300 nm (9–11). The ASFV genome is a linear double-stranded DNA with covalently closed ends and varies about 170–193 kb, which is mainly due to the loss or gain of open reading frames (ORFs) of the multigene families (MGFs) (12,13). ASFV is an enveloped virus with a multiple-layer structure, consisting of an outer capsule membrane, icosahedral capsid, inner envelope, core shell and nucleoid (9–11). The main target host cells for ASFV infection are swine alveolar macrophages and monocytes (14). ASFV enters the cell through phagocytosis and micropinocytosis, and then the virion is uncoated in the late endosome, and finally the virion core containing the genome is released into the cytoplasm (15,16). The ASFV genome could encode more than 150 proteins, which play critical roles in immune evasion, nucleotide metabolism, genome transcription, replication and repair (15). According to the sequence of gene B646L, which encodes the major capsid protein p72, the ASFV could be classified into 24 genotypes (17). Although many studies demonstrated that the gene-deleted attenuated live vaccines could confer ideal protection against the homologous parental strain, they could cause viremia and side effects in the vaccinated pigs (18–22). Moreover, the same gene deletion strategy used in different ASFV genotypes would not result in identical outcomes, which posed a greater challenge for vaccine development (22–26). Therefore, the development of antiviral molecules is gaining more and more attention to combat the ASF pandemic.

Like other NCLDVs, the ASFV life cycle is autonomous, and ASFV encodes a series of replication-related proteins to duplicate the genome, including DNA polymerase pG1211R, clamp-like processivity factor pE301R, DNA ligase pNP419L, ribonucleotide reductase complex pF334L and pF778R, DNA primase pC962R, etc.(12). Among them, the pG1211R gene is presumed to encode a B family DNA polymerase (DNAPol) which is the primary enzyme for genome replication. G1211R protein is one of the most conserved proteins in all ASFV genotypes, presenting an ideal target for drug development. Recently, the structural and functional features of several replication-related proteins including DNA ligase (27), proliferating cell nuclear antigen-like protein (28–30), helicase (31), topoisomerase (32) and polymerase X (33) have been revealed. However, the structural and functional features of ASFV DNAPol (G1211R) remain elusive, which greatly hampers the understanding of ASFV replication mechanism and the drug development.

Materials and methods

Protein expression and purification

The ASFV DNAPol was overexpressed using Bac-to-Bac expression system and purified through tandem affinity chromatography and size-exclusion chromatography as preciously described (34,35). The codon-optimized sequence of ASFV DNAPol (GenBank: AXZ95864.1) was synthesized and fused with N-terminal 8 × His and MBP tags and C-terminal 2 × Strep tag and cloned into pFastBacI transfer vector. The TEV cleavage site was inserted between MBP and ASFV DNAPol, as well as between ASFV DNAPol and strep tag. High five cell (B85502; Invitrogen) were collected by centrifugation (2000 rpm, 4°C, 15 min) after 48 h infection and crushed by sonication in buffer containing 20 mM HEPES, pH 7.5, 500 mM NaCl, 10% glycerol, 1 mM Tris (2-carboxyethyl) phosphine (TCEP) and 1 mM Phenylmethanesulfonyl fluoride (PMSF). Cell debris were removed through centrifugation (12 000 rpm, 4°C, 3 h) and 0.22 μm filter to obtain clear lysate. The targeted protein was sequentially purified by HisTrap and Strep columns (GE Healthcare, 5 mL). The eluted protein was incubated with TEV in a ratio of 1:30 (w/w) at 4°C for 16 h to remove affinity tags. The enzyme-digested solution was loaded onto a size-exclusion chromatography (GE Healthcare, Superdex 200) for a further purification. The purified ASFV DNAPol was pooled and concentrated to ∼2 mg/ml using 30 kDa-cutoff Millipore Ultra centrifugal filter. The ASFV DNAPol was flash frozen and stored at -80°C until to use. Protein mutants were expressed and purified using the same process as the wildtype.

In vitro enzymatic activity assay

To measure whether the purified ASFV DNAPol is catalytically active, polymerase and exonuclease activities were determined as previously reported (36). For polymerase activity assay, the template strand (5′-CTGCACGAATTAAGCAATTCGTAATCATGGTCATAGCT-3′) and primer strand (5′-Cy5-AGCTATGACCATGATTACGAATTG-3′) were mixed and denatured at 95°C for 5 min and immediately cooled on ice. ASFV DNAPol (0.015 μM) was incubated with annealed dsDNA (0.05 μM) in a reaction buffer containing 20 mM Tris–HCl, pH 7.2, 10 mM KCl, 40 mM NaCl, 5 mM MgCl2, 1 mM DTT and 0.05 mM dNTP at 37°C for 2 min and 15 min. For exonuclease activity assay, the reaction system is identical to that of the polymerase activity, except that it lacked dNTP. The assays were quenched, supplemented with formamide and boiled at 100°C for 15 min. The reaction products were analyzed and separated by 20% polyacrylamide-8 M urea gels in 0.5 × TBE buffer. Images were taken using the Vilber Fusion system.

Processive DNA synthesis assay

To evaluate the processivity of ASFV DNAPol, reaction system A containing 0.6 μM ASFV DNApol, 0.5 μM annealed DNA (template DNA: 5′-AATCTCTTTTCACCATAACTTTCTACACCAATTATGCAATTCGTAATCATGGTCATAGCT-3′; primer DNA: 5′- Cy5-AGCTATGACCATGATTACGAATTG -3′), 20 mM Tris–HCl (pH 7.2), 10 mM KCl, 40 mM NaCl and 1 mM DTT was incubated for 10 min at 4°C.The assay was started by mixing reaction system A with reaction system B containing 0.1 mM dNTP mix, 20 mM Tris–HCl, 10 mM KCl, 40 mM NaCl, 10 mM MgCl2, 1 mM DTT, 10 mg/ml heparin and incubating at 30°C for 2 min. The reaction products were analyzed by polyacrylamide-8 M urea gels, as described above.

Cryo-EM sample preparation and data collection

To prepare the cryo-EM specimen of ASFV DNAPol in apo state, the protein was diluted to 0.1 mg/ml using glycerol-free buffer. The diluted protein (4 μl) was applied onto the graphene oxide (GO) coated grids (R1.2/1.3, 300 mesh) and blotted for 2 s with a humidity of 100% at 4°C, and then plunged into liquid ethane using an FEI Vitrobot Mark IV. The prepared grids were transferred to a 300 kV Titan Krios transmission electron microscope (TEM) equipped with Gatan K3 detector and GIF Quantum energy filter. Movies were collected at 105 000× magnification with a calibrated pixel size of 0.69 Å over a defocus range of -1.0 μm to −2.0 μm in super-resolution counting mode with a total dose of 60 e−/Å2 using EPU (ThermoFisher Scientific) automated acquisition software.

For ASFV DNAPol in replicating state, the protein was diluted to 0.1 mg/ml using reaction buffer and mixed with a template strand (5′-CTGCACGAATTAAGCAATTCGTAATCATGGTCATAGCT-3′) and a modified primer containing 3′ di-deoxycytosine chain terminator (5′-AGCTATGACCATGATTACGAATTG-ddC-3′) supplemented with dTTP at ice for 1h. The procedures of cryo-EM sample preparation and data collection were identical to those described above.

For ASFV DNAPol in editing state, the protein was diluted to 0.4 mg/ml with glycerol-free and low salt buffer in absence of Mg2+ and incubated with a template (5′-AATGGTAGGGGAAGGATCGTATGGCCT-3′) and a primer (5′-AGGCCATACGATCCTTCCCCTAC-3′) strand at ice for 30 min. The complex (3 μl) was applied to glow-discharged Nitai grid (R1.2/1.3, 300 mesh) which was blotted for 3 seconds and then plunged into liquid ethane. The micrographs were automatically collected on a 300 kV Titan Krios TEM equipped with K2 detector and GIF Quantum energy filter using SerialEM software (http://bio3d.colorado.edu/SerialEM/). Images were recorded in super-resolution counting mode with a calibrated pixel size of 1.04 Å and a total dose of 60 e−/Å2. The defocus range of this dataset was roughly −1.0 to −2.5 μm.

Image processing

The movie stacks were 2 × binned, aligned, dose-weighted and summed using MotionCor2 (37). All imaging processes including contrast transfer function (CTF) estimation, particle picking and extraction, 2D classification, ab initio model generation and 3D refinements were performed in cryoSPARC v.3.3.1 (38).

For ASFV DNAPol in apo state, a total of 8 423 micrographs were collected. About 1 000 000 particles were picked out using blob-pick procedure of cryoSPARC from 1000 micrographs, and then these particles were extracted and subjected to 2D classification. After three rounds of 2D classification, we selected good particles in different views for Topaz training and then generated the Topaz model (39). Then we applied the Topaz procedure (39) to select particles against entire micrographs, a total of 1 252 665 particles were picked out and extracted from 8 423 micrographs. After two rounds of 2D classification, approximately 758 371 good particles were selected for heterogeneous refinement. Among six classes, two of them showed obvious structural features were selected for homogeneous refinement. We further performed a 3D classification without alignment and three classes were selected for the non-uniform refinement which yielded a density map at 3.32 Å resolution. To polish the resolution, the global CTF-refinement and non-uniform refinement were performed which generated a better density map at 3.28 Å resolution.

For ASFV DNAPol in replicating state, imaging process was similar to that described above. A total of 2 651 305 particles were picked out and extracted from 7013 micrographs using Topaz. After the extensive 2D classification, approximately 1 035 527 particles were selected for heterogeneous refinement and 3D classification which reduced to 290 488 good particles. After global CTF-refinement and non-uniform refinement, we obtained a density map at 2.7 Å resolution.

For ASFV DNAPol in editing state, imaging process was much straightforward. A total of ∼3 200 000 particles were picked out and extracted from 3778 micrographs. After two rounds of 2D classification and heterogeneous refinement, a clean dataset containing 356 638 particles was used for non-uniform refinement which yield a density map at 3.2 Å resolution. The density for the dsDNA was much weak, and we performed another 3D classification without alignment and selected four classes with well-ordered dsDNA for the final round of non-uniform refinement, generating a density map at a resolution of 3.1 Å. All density maps were sharpened using DeepEMhancer (40).

Model building and refinement

The full-length structure of ASFV DNAPol was predicted by AlphaFold2 (41) to act as a starting model. The predicted structure of ASFV DNApol was rigidly docked into the density maps using Chimera and the DNA strands were built manually. The structures were refined against the corresponding density maps using PHENIX (42) with secondary structure restraints and Ramachandran restraints applied. And then, we performed manual adjustment to improve local fit using COOT (43). The stereochemical quality of each coordinate was evaluated using MolProbity (44). Structural figures were prepared by Pymol (https://pymol.org/) and CHIMERAX (45).

Molecular docking

The structure of ASFV DNAPol in replicating state was modified by changing the A13 to G13 in COOT and used as the template structure for molecular docking. The sdf files of CDVpp and Ara-CTP were obtained from Protein Data Bank. The ddTTP was selected to generate the grid box. The protein and ligands were prepared and processed with standard routines. The molecular docking analysis was performed using Schrodinger suites-Maestro 11.5.

Results

The enzymatic feature and apo structure of ASFV DNAPol

To investigate the functional and structural characteristics of ASFV DNAPol, we expressed the protein using bac-to-bac baculovirus expression system. After purification by affinity chromatography, the MBP tag was removed by the TEV enzyme and then the digested solution was further separated using size-exclusion chromatography. The majority of the protein formed aggregates, and a minority was eluted as a monomer (Figure 1A). To test whether the purified ASFV DNAPol is catalytically active, we performed the primer extension assay using a Cy5-labeled 24 nt primer and a 38 nt template, and demonstrated that the ASFV DNAPol could synthesize full-length product in reaction system supplemented with dNTP (Figure 1B). We also showed that the ASFV DNAPol could exert exonuclease activity in dNTP independent manner to digest the labeled primer strand (Figure 1B). To measure the processivity of ASFV DNAPol, we used a 60 nt DNA as a template strand and added heparin into the reaction buffer to perform a single-round replication assay. It showed that the processivity of ASFV DNAPol was poor and the DNAPol could only extend less than 14 nt (Figure 1C). These results suggested that the purified ASFV DNAPol was fully catalytically active and then we used this protein for structural analysis.

Figure 1. Overall structure of ASFV DNAPol. (A) Size-exclusion chromatography and SDS-PAGE profiles of ASFV DNAPol. (B) In vitro enzymatic activity of ASFV DNAPol. The ASFV DNAPol could perform polymerization activity in primer-dependent manner and exonuclease activity in dNTP independent manner. (C) The processivity of ASFV DNAPol. The processivity of ASFV DNAPol is poor and it could synthesize less than 14 nt products in a single-round reaction. (D) Schematic diagrams of domains of ASFV DNAPol. (E, F) Cryo-EM density map (E) and atomic model (F) of ASFV DNAPol in apo state. The active sites of polymerase and exonuclease were indicated by dashed ovals. NTD, blue; Exo, magenta; Palm, cyan; Fingers, yellow; Thumb, green.

Using cryo-electron microscopy (cryo-EM) method, we determined the structure of ASFV DNAPol in apo state at a global resolution of 3.28 Å (Supplementary Figure S1 and Supplementary Table S1). The 3D reconstruction map enabled us to build atomic models for residues 3–993, while the EM density of the C-terminal region was invisible due to its flexibility (Figures 1D–F). By structural comparison with human polymerase delta (46), the ASFV DNAPol could also be divided into five regions including the N-terminal domain (NTD, residues 1–205 and 436–484), exonuclease (Exo, residues 206–435), palm (residues 485–579 and 648–791), fingers (580–647) and thumb (residues 792–1202) subdomains. The polymerase active site is located at the tip of an anti-parallel β-sheet in the palm subdomain, harboring the conserved ‘GDTD’ motif. The exonuclease active site is opposite to the polymerase active site, with a distance of about 40 Å (Figures 1E, F). Without binding of DNA and dNTP substrate, the fingers subdomain adopts an open conformation. We then performed 3D variability analysis to probe the flexibility of ASFV DNAPol. The result showed that the thumb and exonuclease domains are very flexible, moving close or away from each other (Supplementary Movie S1).

Structural basis of ASFV DNAPol in replicating state

To further study the replication mechanism of ASFV DNAPol, we determined the structure of the ASFV DNAPol in replicating state, by incubating dTTP, a template strand and a primer strand with 3′-dideoxycytosine chain terminator and DNAPol in the reaction buffer. The replicating structure of ASFV DNAPol was resolved to 2.7 Å resolution (Supplementary Figure S2 and Supplementary Table S1). In the EM map, we could clearly observe the densities of 14 nt DNA of primer strand, 20 nt DNA of template strand and an incoming dTTP (Figures 2A–B). The NTD and exonuclease domains form an open template entry channel to accommodate the template strand which is similar to other B family DNAPols, except that the poxvirus DNAPol would interact with its processivity factor to form a closed template entry channel (36). Compared with the apo state, the fingers domain and thumb domain make great conformational changes to bind with the dsDNA and the incoming dTTP (Figure 2C). The fingers domain rotates by ∼30 degrees, switching from the open state to the closed state to stabilize the dTTP (Figure 2D). The thumb domain changes its conformation to clamp and hold up the dsDNA (Figure 2E). The global conformation of polymerase became more stable, while the distal dsDNA was flexible, possibly due to lack of processivity factor (Supplementary Movie S2).

Figure 2. The structure of ASFV DNAPol in replicating state. (A, B) Cryo-EM density map (A) and atomic model (B) of ASFV DNAPol in replicating state. The polymerase is colored by domains as in Figure 1. Template, grey; primer, red. The template entry channel is indicated by black arrowhead. (C) Comparison of structures of ASFV DNAPol in replicating and apo states. (D, E) Conformational changes of the fingers (D) and thumb (E) domains once the ASFV DNAPol transforming from apo into replicating state. The structures of ASFV DNAPol in replicating state and apo state are colored in domains as in Figure 1 and grey, respectively.

Although the sequence identity between ASFV DNAPol and other B family DNA polymerases is low, their three-dimensional structures are comparable (Supplementary Figures S3A-F). Interestingly, the ASFV DNAPol exhibited the highest similarities to the human DNA polymerase delta, giving a root-mean-square deviation (RMSD) of 2.1 Å, instead of the DNA polymerase of Mpox virus which belongs to the NCLDVs (Supplementary Figures S3A−F). The ASFV DNAPol and human polymerase delta structures could be overlaid very well, except two regions which show obvious differences (Supplementary Figures S3G-H). The loop of a helix-turn-helix structure (residues 802–841) of thumb domain in ASFV DNAPol is much longer than the counterpart in the human polymerase delta, leading to a more extended conformation (Supplementary Figures S3G, H). In the palm subdomain of human polymerase delta, four β strands make up the polymerase active site, and the last two β strands are connected by a loop and a helix. While in ASFV DNAPol, these two β strands are connected by two helices that are more complex and much longer than the counterpart of the human polymerase delta (Supplementary Figures S3G, H). Intriguingly, in ASFV DNAPol, residues Y707 and K712 of palm subdomain form hydrogen bonds with the E821 and R819 of thumb domain which was not seen in other B family DNAPols (Supplementary Figures S3G, H). The fingers domain consists of two helices and is perpendicular to the axis of dsDNA which is critical to the formation of replicating complex. Although the topological structure of the fingers domain in ASFV DNAPol and human polymerase delta is identical, the length of the fingers domain varies in these two structures (40 Å in human polymerase delta vs. 50 Å in ASFV DNAPol) (Supplementary Figure S3I).

The interactions between ASFV DNAPol and DNA are mainly formed by hydrogen bonds between positively charged amino acids and phosphodiester backbone (Figure 3). For template strand, residues R920, K921 and Y947 of the thumb subdomain form four hydrogen bonds with the sugar-phosphate backbone of the G21-A23 (Figure 3C). A few weak van der Waals contacts are formed between C20 and residues of the thumb subdomain. The sidechains of R954 and K770 form three hydrogen bonds with the phosphate group of T19. Meanwhile, the K770 also interacts with the backbone of T18 (Figure 3D). There are no hydrogen bonds formed between polymerase and G14-A17. Residue K344 of the β-hairpin of exonuclease subdomain formed two hydrogen bonds with the backbone of A12 and A13 (Figure 3E). For primer strand, residues R857, D861 and R897 of thumb subdomain form three hydrogen bonds with the sugar-phosphate backbone of A20-T22 (Figure 3F). The residues K376 of exonuclease subdomain and Y771 and K791 of palm subdomain make three hydrogen bonds with the last three bases of the primer strand (Figure 3G).

Figure 3. The interactions between polymerase and dsDNA and incoming dTTP in replicating state. (A, B) Overall structure of ASFV DNAPol in replicating state in different views. The polymerase is shown as cartoon and the dsDNA is shown as surface and colored as in Figure 2. (C–E) Major interactions between polymerase and template strand. (F, G) Major interactions between polymerase and primer strand. (H) Major interactions between polymerase and incoming dTTP. The critical residues are shown as sticks, hydrogen bonds and coordinate bonds are shown as yellow and green dashed lines, respectively.

The incoming dTTP substrate is clearly visible in the ASFV DNAPol active site with its triphosphate moiety sandwiched between the palm and fingers subdomains (Figure 3). The γ-phosphate forms two hydrogen bonds with the side chains of R593 and K635 of the fingers subdomain, and the β-phosphate makes two hydrogen bonds with the main chain of S520 and L521. There exists a metal ion coordinated by the side chains of highly conserved aspartate residues (D517 and D689), main chain of F518 and triphosphate tail of dTTP (Figure 3H). The incoming dTTP is bound in a complex hydrogen-bonding network which is similar to that was seen in other B family polymerase-dsDNA-dNTP complexes (28,36,46). Of note, the main chain of Y522 makes a hydrogen bond with the 3′-OH of the deoxyribose and the side chain of Y522 stacks with the deoxyribose, acting as a ‘steric gating’ to exclude the ribonucleotides binding by generating a repulsion with the 2′-OH of the ribose (36,46,47).

Structural basis of ASFV DNAPol in editing state

Although the ASFV DNAPol is capable of selecting and incorporate the correct substrates into the nascent strand, the mismatched incorporation is still able to occur in a low rate as other B family DNA polymerases (48–50). The proofreading process executed by exonuclease subdomain could increase the replication fidelity by 10-fold (49,50). To clarify the proofreading mechanism of ASFV DNAPol, we determined the structure of ASFV DNAPol-dsDNA complex in editing state using cryo-EM at a resolution of 3.1 Å (Supplementary Figure S4 and Supplementary Table S1). In the editing structure, the dsDNA was unwound with three base pairs and the 3′-terminal of primer strand was bound in the exonuclease active site (Figures 4A, B). The unwound three bases insert into a hydrophobic pocket and form extensive interactions with surrounding residues (Figures 4C, D). Residue T214 of the exonuclease subdomain makes a hydrogen bond with the 3′-OH of the deoxyribose of the last deoxynucleotide residue C23 of primer strand, and side chain of residue Y399 not only forms a hydrogen bond with the backbone of the C23 but also stacks with the base of the C23. The side chain of Y420 also forms a hydrogen bond with the backbone of the C23. The penultimate deoxynucleotide residue A22 forms a hydrogen bond with the mainchain of L382, and makes a π–π interaction with the side chain of Y299. In addition, residue T380 and R367 form three hydrogen bonds with the backbone of T21 (Figure 4D). Through these abundant interactions, the unwound bases are fixed at the exonuclease subdomain, in which the last phosphodiester bond is placed in a position close to catalytic residues D211 and E213 in the exonuclease active site with a distance of ∼4 Å. The last phosphodiester bond would be cleaved by a two-metal mechanism to remove the mismatched base (51). In our structure, we failed to observe any density for metal ions mainly due to no addition of Mg2+ into the cryo-EM sample preparation solution.

Figure 4. Overall structure of ASFV DNAPol in editing state. (A, B) The density map (A) and atomic model (B) of ASFV DNAPol in editing state. The polymerase and dsDNA are colored as in Figure 2. The exonuclease active site is indicated by black dashed oval. (C) The hydrophobic binding pocket for unwound primer in exonuclease domain. The exonuclease is shown as surface and colored by hydrophobic properties. (D) Major interactions between polymerase and unwound primer. The critical residues are shown as sticks, hydrogen bonds and π–π interactions are shown as yellow and black dashed lines, respectively.

Superimposed with the replicating structure, we found that the 3′-terminal of primer strand was translocated by a long distance of 40 Å from polymerase active site to exonuclease active site, accompanied by the rotation of the dsDNA axis and thumb subdomain (Supplementary Figure S5A). Because of no incoming dNTP substrate binding in the polymerase active site, the fingers domain retracts back to adopt an open conformation like the apo structure (Supplementary Figure S5B) and the conformational flexibility increases (Supplementary Movie S3). We also noticed that the conformations of the novel palm-thumb interaction site in the ASFV DNAPol have changed slightly for different functional states (Figure 5A). Compared with the replicating structure, in the apo structure, R819 and E821 fail to form hydrogen bonds with K712 and Y707 due to the longer distance. While in the editing structure, residue R819 fails to form a hydrogen bond to the main chain of K712, and instead E821 forms a hydrogen bond with the side chain of K712 (Figure 5). It suggests that these interactions might be involved in the DNA synthesis process of the ASFV DNAPol, which is accompanied by conformational change. To test our hypothesis, we substituted the key residues K712, R819 and E821 with alanine and demonstrated that both polymerase and exonuclease activities decreased for the mutant DNAPol (Figure 5E–F). We then compared the structures of ASFV DNAPol and other B family DNA polymerases in editing state, the exonuclease subdomain and last three unwound bases could be well overlaid (Supplementary Figure S6A). However, the interaction details are different. For ASFV DNAPol, the last two bases simultaneously form two π–π stacking interactions with Y299 and Y399 located in two helices flanking at the exonuclease active site (Supplementary Figure S6B). For P.abyssi (52) and RB69 (53) DNAPols in editing state, only the last base or the penultimate base forms an π-π stacking interaction with an aromatic amino acid located in a loop connecting the β-sheet of exonuclease active site (Supplementary Figures S6C, D).

Figure 5. Comparison of the novel palm-thumb interaction site in the ASFV DNAPol under different functional states. (A) Structural comparisons of the novel palm-thumb interaction regions in the ASFV DNAPol for apo, editing and replicating states. (B–D) The interactions between palm and thumb domain in replicating (B), editing (C) and apo (D) structures. (E) The influence of palm-thumb interaction on the ASFV DNAPol enzymatic activity. (F) Quantification of the polymerase enzymatic activity shown in the (E). The hydrogen bonds were indicated by yellow dashed lines.

Previous studies have reported that the β-hairpin, located at the interface between NTD and exonuclease, plays an important role in the separation of DNA duplex strands when the polymerase switches from replicating to editing state (54,55). The β-hairpin would hold the template in place preventing it from translocating with primer strand (54,56). In the replicating structure of ASFV DNAPol, the β-hairpin (residues 337–357) directly inserts into the major groove at the junction between duplex and single-stranded DNA, and makes interactions with the unpaired template strand (Figure 3). In the editing structure of ASFV DNAPol, the density of unpaired template is much weaker, we could just trace the main chain of dA1–dA7 using the unsharpened EM density map (Supplementary Figure S7). No matter what working state ASFV DNAPol adopts, the β-hairpin always resides below the unpaired template strand, to stabilize the template strand during duplex DNA unwinding (Figure 6A). While in other known editing structures of B family DNA polymerase, the position of β-hairpin is located above the unpaired template strand (Figures 6B–C). Structural comparison reveals that the β-hairpin in ASFV DNAPol has a different orientation compared with those in P.abyssi (52) and RB69 (53) DNAPols (Figure 6D). Furthermore, we found that the β-hairpins of ASFV, human and yeast DNAPols have similar orientations (Figure 6E), suggesting that the β-hairpin in these DNAPols would have a similar working mechanism for the duplex DNA unwinding. We propose that B family polymerases might have evolved two ways to facilitate the duplex strand separation during polymerase switching from replicating state to editing state.

Figure 6. Distinctive roles of β-hairpin in separating template and primer strands. (A-C) The structures of ASFV (A), P.abyssi (PDB ID:4FLT) (52) (B) and RB69 (PDB ID:1CLQ) (53) (C) DNA polymerase in editing state. In different B-family DNA polymerase structures, the β-hairpins are located in different positions related to unwound template strand. For ASFV DNAPol, the β-hairpin is underneath the template strand resembling that it drags the template to facilitate the separation of primer and template strands. While for P.abyssi and RB69 DNAPol, the β-hairpin is above the template strand resembling that it pushes the template to help the separation of primer and template strands. (D) Overlay of structures of exonuclease domains of ASFV, P.abyssi and RB69 DNA polymerase. The relative position of β-hairpins is varied, in which the ASFV β-hairpin is located in the lowest position. (E) Overlay of structures of exonuclease domains of ASFV, human and yeast DNA polymerase, in which all β-hairpins are located in similar position. There exist two modes to unwind the dsDNA during DNAPol switching from replicating to editing state by β-hairpin.

Discussion

ASF is one of the most severe swine viral diseases, which has caused significant economic losses, and it is still ongoing in the world. Nowadays, ASFV has spread into many non-endemic countries including Sweden, Bosnia and Herzegovina, Korea and Germany (https://wahis.woah.org/). Due to the complexity of virion and viral life cycle, currently no effective vaccines and antiviral drugs are available. The ASFV DNAPol, encoded by G1211R, is response for genome replication and an ideal target for drug development. In the past years, we only knew that ASFV DNAPol belongs to the B family DNA polymerase, but the structural and functional features are yet elusive. In this study, we demonstrated that ASFV DNAPol possessed polymerase and exonuclease activities with poor processivity and resolved the structures of ASFV DNAPol in different working states including apo, replicating and editing states. Based on these structures, we could build up the working model of ASFV DNAPol during genome replication.

Once the virus entering into the cytoplasm, the DNAPol was expressed immediately from partially uncoated viral core (12). The newly generated DNAPol was in apo state and would bind to a single strand nick in the genome, in which the exposed 3′-OH group acting as a primer (12). The thumb domain would make a conformation change to accommodate the dsDNA. The substrates dNTPs enter the polymerase active site to pair with the template and form a complicated hydrogen-bonding network with the metal ions and residues of the palm and fingers domains. Due to interactions between the incoming dNTP and positive amino acids (R593 and K635) of the fingers domain, the fingers domain was dragged up to be close to the polymerase active site and formed a closed conformation. In the active site, the activated 3′-OH of primer would perform nucleophilic attack on the α-phosphate of the incoming dNTP to form a phosphodiester bond. Subsequently, the byproduct pyrophosphate group was released along with the translocation of dsDNA and retraction of fingers domain to empty the polymerase active site. If the incorporated dNTP was correct, the polymerase would continue to catalyze the next round of polymerization for DNA elongation. If the polymerase incorporated a mismatched base, the dsDNA would be unwound, and 3′-terminal of primer strand would transfer ∼40 Å from polymerase active site into the exonuclease active site with the help of the β-hairpin in the exonuclease subdomain. The β-hairpin is located below the template strand and stabilizes the unpaired template strand to facilitate the dsDNA separation. The last three bases of primer strand were fixed in the exonuclease active site via extensive interactions, with the last phosphodiester bond exposed to the exonuclease active residues. After the last mismatched base was incised, the primer strand was translocated back into the polymerase active site for a new incorporation of dNTP (Supplementary Figure S8).

We have noticed that the C-terminal region (CTD) in our determined structures was invisible due to its flexibility. To analyze the function of the C-terminal region on ASFV DNApol activity, we constructed the CTD-deletion mutant and purified the protein. Compared with the wild-type DNAPol, the CTD-deletion mutant showed little aggregates and a higher protein expression level. We also tested the polymerase and exonuclease activity of the CTD-deletion mutant, and found that it displayed a higher activity than the wild-type DNAPol (Supplementary Figure S9). We deduced that the deletion of the CTD allowed the ASFV DNAPol to become more stable. The structure of ASFV DNAPol shows the highest similarity to the human DNAPol, suggesting that they would take similar mechanisms to synthesize the product. The CTD of human DNAPol is responsible for binding with the processivity factor (46), and thus we supposed the CTD of ASFV DNAPol might also interact with the processivity factor.

Previous studies have reported that brincidofovir (57) and cytarabine (58) could inhibit ASFV infection. Both drugs are nucleoside analogs that would be transformed into active metabolites in vivo and served as competitive inhibitors of dCTP. To estimate the binding mode between drugs and ASFV DNAPol, we performed molecular docking analysis based on our determined structures. From our docking results, the CDVpp (cidofovir diphosphate, the active form of brincidofovir) entered into the active site of ASFV DNAPol through forming hydrogen bonds with the main chain of S520 and L521 and the side chain of R593 and K635, which were also critical to stabilize the incoming dNTP (Supplementary Figure S10A). However, it lacked the hydrogen bond with Y522, implying that the CDVpp might be not an ideal competitive substrate, as validated in poxvirus DNAPol (59). For Ara-CTP (cytarabine triphosphate), it not only forms hydrogen bonds with the residues S520, L521, Y522, R593 and K635, but also forms extra hydrogen bonds with the residue N639, making it is more potent in binding with ASFV DNAPol than the native dNTP (Supplementary Figure S10B).

During the genome replication, the DNAPol is required to interact with other replicative proteins to form a replication machinery. Currently, several critical proteins have been identified, such as pE301R (28–30), pC962R (31) and pP1192R (32). Notably, the pE301R was presumed to bind with DNAPol and act as processivity factor which is indispensable for genome replication. Recently, the structure of pE301R was determined in trimeric (28,30) or tetrameric (29) form and assembled into a ring architecture like previously reported structures of other proliferating cell nuclear antigens (PCNA). In eukaryotes, other replication factors are required to load the PCNA onto the DNAPol-dsDNA complex. However, there are no homolog proteins encoded by ASFV genome. Therefore, the molecular mechanism of how pE301R is loaded onto DNAPol–dsDNA complex is required to be studied in the future.

In summary, we determined the structures of ASFV DNAPol in apo, replicating and editing states, and greatly improved the molecular understanding of the replication mechanism of ASFV. Furthermore, the interaction details of ASFV DNAPol in different work states would provide an important molecular basis for the development of antiviral drugs.

Supplementary Material

gkae739_Supplemental_Files

Acknowledgements

We thank all the staff members at the center for Biological Imaging (CBI), Institute of Biophysics (IBP), Chinese Academy of Science (CAS) and the cryo-EM Center, Shanxi Academy of Advanced Research and Innovation for assistance with data collection.

Author contributions: Y.S. conceived and supervised the study. L.K. and X.Z. purified the protein samples and conducted biochemical experiments. L.K., J.S. and Q.P. prepared the cryo-EM specimens and collected data. J.S. and Q.P. conducted the image processing and reconstruction. Q.P. built the atomic models. Y.S. and Q.P. analyzed the structures. Q.P. and Y.S. wrote the manuscript. All authors participated in the discussion and manuscript editing.

Data availability

The cryo-EM density maps and atomic coordinates have been deposited to the Electron Microscopy Data Bank (EMDB) and the Protein Data Bank (PDB), under accession codes 8YWG (EMDB-39632), 8YWG (EMDB-39634) and 8YWM (EMDB-39638)for the ASFV DNApol in apo, replicating and editing states, respectively.

Supplementary data

Supplementary Data are available at NAR Online.

Funding

This work was funded by grants from the National Key R&D Program of China (2021YFC2300200 to Q.P. and 2021YFC2300700 to Y.S.), and National Natural Science Foundation of China (NSFC) (81871658 and 32192452 to Y.S., 32100119 to Q.P. and 32100129 to S.L.). Funding for open access charge: National Key R&D Program of China [2021YFC2300200].

Conflict of interest statement. None declared.
==== Refs
References

1. Simoes M. , FreitasF.B., LeitaoA., MartinsC., FerreiraF. African swine fever virus replication events and cell nucleus: new insights and perspectives. Virus Res. 2019; 270 :197667.31319112
2. Arzt J. , WhiteW.R., ThomsenB.V., BrownC.C. Agricultural diseases on the move early in the third millennium. Vet. Pathol. 2010; 47 :15–27.20080480
3. Sánchez-Cordón P.J. , MontoyaM., ReisA.L., DixonL.K. African swine fever: a re-emerging viral disease threatening the global pig industry. Vet. J. 2018; 233 :41–48.29486878
4. Zhou X.T. , LiN., LuoY.Z., LiuY., MiaoF.M., ChenT., ZhangS.F., CaoP.L., LiX.D., TianK.G.et al . Emergence of African swine fever in China, 2018. Transbound. Emerg. Dis. 2018; 65 :1482–1484.30102848
5. Mighell E. , WardM.P. African Swine Fever spread across Asia, 2018-2019. Transbound. Emerg. Dis. 2021; 68 :2722–2732.33599077
6. Cooper T.L. , SmithD., GonzalesM.J.C., MaghanayM.T., SandersonS., CornejoM.R.J.C., PinedaL.L., SagunR.A.A., SalvacionO.P. Beyond numbers: determining the socioeconomic and livelihood impacts of African Swine Fever and its control in the Philippines. Front. Vet. Sci. 2022; 8 :734236.35224068
7. Mayer S.V. , TeshR.B., VasilakisN. The emergence of arthropod-borne viral diseases: a global prospective on dengue, chikungunya and zika fevers. Acta Trop. 2017; 166 :155–163.27876643
8. Liu Z. , PengQ., HanP., KuaiL., QiJ., ShiY. Crystal structures of RNA-dependent RNA polymerases from Jingmen tick virus and Alongshan virus. Hlife. 2024; 2 :18–31.
9. Liu S. , LuoY.Z., WangY.J., LiS.H., ZhaoZ.N., BiY.H., SunJ.Q., PengR.C., SongH., ZhuD.J.et al . Cryo-EM structure of the African Swine Fever Virus. Cell Host Microbe. 2019; 26 :836–843.31787524
10. Andrés G. , CharroD., MatamorosT., DillardR.S., AbresciaN.G.A. The cryo-EM structure of African swine fever virus unravels a unique architecture comprising two icosahedral protein capsids and two lipoprotein membranes. J. Biol. Chem. 2020; 295 :1–12.31649031
11. Wang N. , ZhaoD.M., WangJ.L., ZhangY.L., WangM., GaoY., LiF., WangJ.F., BuZ.G., RaoZ.H.et al . Architecture of African swine fever virus and implications for viral assembly. Science. 2019; 366 :640–644.31624094
12. Dixon L.K. , ChapmanD.A.G., NethertonC.L., UptonC. African swine fever virus replication and genomics. Virus Res. 2013; 173 :3–14.23142553
13. Chapman D.A.G. , TcherepanovV., UptonC., DixonL.K. Comparison of the genome sequences of nonpathogenic and pathogenic African swine fever virus isolates. J. Gen. Virol. 2008; 89 :397–408.18198370
14. Gómez-Villamandos J.C. , BautistaM.J., Sánchez-CordónP.J., CarrascoL. Pathology of African swine fever: the role of monocyte-macrophage. Virus Res. 2013; 173 :140–149.23376310
15. Duan X.H. , RuY., YangW.P., RenJ.J., HaoR.Z., QinX.D., LiD., ZhengH.X. Research progress on the proteins involved in African swine fever virus infection and replication. Front. Immunol. 2022; 13 :947180.35935977
16. Wang Y. , KangW.F., YangW.P., ZhangJ., LiD., ZhengH.X. Structure of African Swine Fever virus and associated molecular mechanisms underlying infection and immunosuppression: a review. Front. Immunol. 2021; 12 :715582.34552586
17. Malogolovkin A. , BurmakinaG., TitovI., SeredaA., GoginA., BaryshnikovaE., KolbasovD. Comparative analysis of African Swine Fever Virus genotypes and serogroups. Emerg. Infect. Dis. 2015; 21 :312–315.25625574
18. Reis A.L. , GoatleyL.C., JabbarT., Sanchez-CordonP.J., NethertonC.L., ChapmanD.A.G., DixonL.K. Deletion of the African Swine fever virus gene DP148R does not reduce virus replication in culture but reduces virus virulence in pigs and induces high levels of protection against challenge. J. Virol. 2017; 91 :e01428-17.
19. Gallardo C. , SánchezE.G., Pérez-NúñezD., NogalM., de LeónP., CarrascosaA.L., NietoR., SolerA., AriasM.L., RevillaY. African swine fever virus (ASFV) protection mediated by NH/P68 and NH/P68 recombinant live-attenuated viruses. Vaccine. 2018; 36 :2694–2704.29609966
20. O’Donnell V. , HolinkaL.G., KrugP.W., GladueD.P., CarlsonJ., SanfordB., AlfanoM., KramerE., LuZ.Q., ArztJ.et al . African Swine Fever Virus Georgia 2007 with a deletion of virulence-associated gene (B119L), when administered at low doses, leads to virus attenuation in swine and induces an effective protection against homologous challenge. J. Virol. 2015; 89 :8556–8566.26063424
21. O’Donnell V. , RisattiG.R., HolinkaL.G., KrugP.W., CarlsonJ., Velazquez-SalinasL., AzzinaroP.A., GladueD.P., BorcaM.V. Simultaneous deletion of the and genes from the African Swine fever virus Georgia 2007 isolate offers increased safety and protection against homologous challenge. J. Virol. 2017; 91 :e01760-16.27795430
22. O’Donnell V. , HolinkaL.G., GladueD.P., SanfordB., KrugP.W., LuX.Q., ArztJ., ReeseB., CarrilloC., RisattiG.R.et al . African Swine Fever virus Georgia isolate harboring deletions of MGF360 and MGF505 genes is attenuated in swine and confers protection against challenge with virulent parental virus. J. Virol. 2015; 89 :6048–6056.25810553
23. Ramirez-Medina E. , VuonoE., O’DonnellV., HolinkaL.G., SilvaE., RaiA., PruittS., CarrilloC., GladueD.P., BorcaM.V. Differential effect of the deletion of African Swine Fever virus virulence-associated genes in the induction of attenuation of the highly virulent Georgia strain. Viruses-Basel. 2019; 11 :599.
24. Borca M.V. , O’DonnellV., HolinkaL.G., RisattiG.R., Ramirez-MedinaE., VuonoE.A., ShiJ.S., PruittS., RaiA., SilvaE.et al . Deletion of CD2-like gene from the genome of African swine fever virus strain Georgia does not attenuate virulence in swine. Sci. Rep. 2020; 10 :494.31949276
25. Monteagudo P.L. , LacastaA., LópezE., BoschL., ColladoJ., Pina-PedreroS., Correa-FizF., AccensiF., NavasM.J., VidalE.et al . BA71ΔCD2: a new recombinant live attenuated African Swine fever virus with cross-protective capabilities. J. Virol. 2017; 91 :e01058-17.28814514
26. O’Donnell V. , TiolinkaL.G., SanfordB., KrugP.W., CarlsonJ., PachecoJ.M., ReeseB., RisattiG.R., GladueD.P., BorcaM.V. African swine fever virus Georgia isolate harboring deletions of 9GL and MGF360/505 genes is highly attenuated in swine but does not confer protection against parental virus challenge. Virus Res. 2016; 221 :8–14.27182007
27. Chen Y.Q. , LiuH.H., YangC., GaoY.Q., YuX., ChenX., CuiR.X., ZhengL.N., LiS.H., LiX.H.et al . Structure of the error-prone DNA ligase of African swine fever virus identifies critical active site residues. Nat. Commun. 2019; 10 :387.30674878
28. Wu J. , ZhengH., GongP. Crystal structure of African swine fever virus pE301R reveals a ring-shaped trimeric DNA sliding clamp. J. Biol. Chem. 2023; 299 :104872.37257822
29. Li S. , GeH.L., LiY.H., ZhangK.H., YuS.X., CaoH.W., WangY.J., DengH., LiJ.Q., DaiJ.W.et al . The E301R protein of African swine fever virus functions as a sliding clamp involved in viral genome replication. mBio. 2023; 15 :e0164523.
30. Shao Z.W. , YangJ., GaoY.Q., ZhangY.X., ZhaoX., ShaoQ.Y., ZhangW.Z., CaoC.L., LiuH.H., GanJ.H. Structural and functional studies of PCNA from African swine fever virus. J. Virol. 2023; 97 :e0074823.37534905
31. Shao Z.W. , SuS.C., YangJ., ZhangW.Z., GaoY.Q., ZhaoX., ZhangY.X., ShaoQ.Y., CaoC.L., LiH.L.et al . Structures and implications of the C962R protein of African swine fever virus. Nucleic Acids Res. 2023; 51 :9475–9490.37587714
32. Zhao Y. , KuangW., AnQ., LiJ., WangY., DengZ. Cryo-EM structures of African swine fever virus topoisomerase. mBio. 2023; 14 :e01228-23.37610250
33. Chen Y.Q. , ZhangJ., LiuH.H., GaoY.Q., LiX.H., ZhengL.N., CuiR.X., YaoQ.Q., RongL., LiJ.X.et al . Unique 5′-P recognition and basis for dG: dGTP misincorporation of ASFV DNA polymerase X. PLoS Biol. 2017; 15 :e1002599.28245220
34. Peng Q. , YuanB., ChengJ.L., WangM., GaoS.W., BaiS.R., ZhaoX.J., QiJ.X., GaoG.F., ShiY. Molecular mechanism of de novo replication by the Ebola virus polymerase. Nature. 2023; 622 :603–610.37699521
35. Peng Q. , PengR., YuanB., WangM., ZhaoJ., FuL., QiJ., ShiY. Structural basis of SARS-CoV-2 polymerase inhibition by Favipiravir. Innovation (Camb). 2021; 2 :100080.33521757
36. Peng Q. , XieY.F., KuaiL., WangH., QiJ.X., GaoG.F., ShiY. Structure of monkeypox virus DNA polymerase holoenzyme. Science. 2023; 379 :100–105.36520947
37. Zheng S.Q. , PalovcakE., ArmacheJ.P., VerbaK.A., ChengY., AgardD.A. MotionCor2: anisotropic correction of beam-induced motion for improved cryo-electron microscopy. Nat. Methods. 2017; 14 :331–332.28250466
38. Punjani A. , RubinsteinJ.L., FleetD.J., BrubakerM.A. cryoSPARC: algorithms for rapid unsupervised cryo-EM structure determination. Nat. Methods. 2017; 14 :290–296.28165473
39. Bepler T. , MorinA., RappM., BraschJ., ShapiroL., NobleA.J., BergerB. Positive-unlabeled convolutional neural networks for particle picking in cryo-electron micrographs. Nat. Methods. 2019; 16 :1153–1160.31591578
40. Sanchez-Garcia R. , Gomez-BlancoJ., CuervoA., CarazoJ.M., SorzanoC.O.S., VargasJ. DeepEMhancer: a deep learning solution for cryo-EM volume post-processing. Commun. Biol. 2021; 4 :874.34267316
41. Jumper J. , EvansR., PritzelA., GreenT., FigurnovM., RonnebergerO., TunyasuvunakoolK., BatesR., ZídekA., PotapenkoA.et al . Highly accurate protein structure prediction with AlphaFold. Nature. 2021; 596 :583–589.34265844
42. Adams P.D. , AfonineP.V., BunkocziG., ChenV.B., DavisI.W., EcholsN., HeaddJ.J., HungL.W., KapralG.J., Grosse-KunstleveR.W.et al . PHENIX: a comprehensive Python-based system for macromolecular structure solution. Acta. Crystallogr. D Biol. Crystallogr. 2010; 66 :213–221.20124702
43. Emsley P. , CowtanK. Coot: model-building tools for molecular graphics. Acta. Crystallogr. D Biol. Crystallogr. 2004; 60 :2126–2132.15572765
44. Chen V.B. , ArendallW.B.3rd, HeaddJ.J., KeedyD.A., ImmorminoR.M., KapralG.J., MurrayL.W., RichardsonJ.S., RichardsonD.C. MolProbity: all-atom structure validation for macromolecular crystallography. Acta. Crystallogr. D Biol. Crystallogr. 2010; 66 :12–21.20057044
45. Goddard T.D. , HuangC.C., MengE.C., PettersenE.F., CouchG.S., MorrisJ.H., FerrinT.E. UCSF ChimeraX: meeting modern challenges in visualization and analysis. Protein Sci. 2018; 27 :14–25.28710774
46. Lancey C. , TehseenM., RaducanuV.S., RashidF., MerinoN., RaganT.J., SavvaC.G., ZaherM.S., ShirbiniA., BlancoF.J.et al . Structure of the processive human pol δ holoenzyme. Nat. Commun. 2020; 11 :1109.32111820
47. Swan M.K. , JohnsonR.E., PrakashL., PrakashS., AggarwalA.K. Structural basis of high-fidelity DNA synthesis by yeast DNA polymerase δ. Nat. Struct. Mol. Biol. 2009; 16 :979–986.19718023
48. Arana M.E. , KunkelT.A. Mutator phenotypes due to DNA replication infidelity. Semin. Cancer Biol. 2010; 20 :304–311.20934516
49. Shcherbakova P.V. , PavlovY.I., ChilkovaO., RogozinI.B., JohanssonE., KunkelT.A. Unique error signature of the four-subunit yeast DNA polymerase ϵ. J. Biol. Chem. 2003; 278 :43770–43780.12882968
50. Korona D.A. , LeCompteK.G., PursellZ.F. The high fidelity and unique error signature of human DNA polymerase ϵ. Nucleic Acids Res. 2011; 39 :1763–1773.21036870
51. Beese L.S. , SteitzT.A. Structural basis for the 3′-5′ Exonuclease activity of Escherichia-coli DNA-polymerase-I - a 2 metal-ion mechanism. EMBO J. 1991; 10 :25–33.1989886
52. Gouge J. , RalecC., HennekeG., DelarueM. Molecular recognition of canonical and deaminated bases by P. abyssi family B DNA polymerase. J. Mol. Biol. 2012; 423 :315–336.22902479
53. Shamoo Y. , SteitzT.A. Building a replisome from interacting pieces: sliding clamp complexed to a peptide from DNA polymerase and a polymerase editing complex. Cell. 1999; 99 :155–166.10535734
54. Hogg M. , AllerP., KonigsbergW., WallaceS.S., DoubliéS. Structural and biochemical investigation of the role in proofreading of a β hairpin loop found in the exonuclease domain of a replicative DNA polymerase of the B family. J. Biol. Chem. 2007; 282 :1432–1444.17098747
55. Subuddhi U. , HoggM., Reha-KrantzL.J. Use of 2-aminopurine fluorescence to study the role of the β hairpin in the proofreading pathway catalyzed by the phage T4 and RB69 DNA polymerases. Biochemistry. 2008; 47 :6130–6137.18481871
56. Reha-Krantz L.J. , MarquezL.A., ElisseevaE., BakerR.P., BloomL.B., DunfordH.B., GoodmanM.F. The proofreading pathway of bacteriophage T4 DNA polymerase. J. Biol. Chem. 1998; 273 :22969–22976.9722519
57. Guo S.B. , ZhangY.B., LiuZ.K., WangD.Z., LiuH., LiL., ChenQ.C., YangD., LiuQ.Y., GuoH.H.et al . Brincidofovir is a robust replication inhibitor against African swine fever virus in vivo and in vitro. Emerg. Microbes Infect. 2023; 12 :2220572.37272334
58. Li T.T. , ZhengJ., HuangT., WangX., LiJ.N., JinF., WeiW.J., ChenX., LiuC.X., BaoM.F.et al . Identification of several African swine fever virus replication inhibitors by screening of a library of FDA-approved drugs. Virology. 2024; 593 :110014.38401340
59. Magee W.C. , HostetlerK.Y., EvansD.H. Mechanism of inhibition of vaccinia virus DNA polymerase by cidofovir diphosphate. Antimicrob. Agents Chemother. 2005; 49 :3153–3162.16048917
