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

39077950
10.1093/nar/gkae657
gkae657
AcademicSubjects/SCI00010
Nucleic Acid Enzymes
Structural insights into the assembly of type IIA topoisomerase DNA cleavage-religation center
Liu Ko-Ting Institute of Biochemistry and Molecular Biology, College of Medicine, National Taiwan University, Taipei 100, Taiwan

Chen Shin-Fu Institute of Biochemistry and Molecular Biology, College of Medicine, National Taiwan University, Taipei 100, Taiwan

https://orcid.org/0000-0003-0139-6513
Chan Nei-Li Institute of Biochemistry and Molecular Biology, College of Medicine, National Taiwan University, Taipei 100, Taiwan
Life Science Group, Scientific Research Division, National Synchrotron Radiation Research Center, Hsinchu 30076, Taiwan

To whom correspondence should be addressed. Tel: +886 2 23562214; Fax: +886 2 23915295; Email: nlchan@ntu.edu.tw
09 9 2024
30 7 2024
30 7 2024
52 16 97889802
16 7 2024
9 7 2024
20 12 2023
© 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 reprints@oup.com for reprints and translation rights for reprints. All other permissions can be obtained through our RightsLink service via the Permissions link on the article page on our site-for further information please contact journals.permissions@oup.com.

Abstract

The ability to catalyze reversible DNA cleavage and religation is central to topoisomerases' role in regulating DNA topology. In type IIA topoisomerases (Top2), the formation of its DNA cleavage-religation center is driven by DNA-binding-induced structural rearrangements. These changes optimally position key catalytic modules, such as the active site tyrosine of the WHD domain and metal ion(s) chelated by the TOPRIM domain, around the scissile phosphodiester bond to perform reversible transesterification. To understand this assembly process in detail, we report the catalytic core structures of human Top2α and Top2β in an on-pathway conformational state. This state features an in trans formation of an interface between the Tower and opposing TOPRIM domain, revealing a groove for accommodating incoming G-segment DNA. Structural superimposition further unveils how subsequent DNA-binding-induced disengagement of the TOPRIM and Tower domains allows a firm grasp of the bound DNA for cleavage/religation. Notably, we identified a previously undocumented protein-DNA interaction, formed between an arginine-capped C-terminus of an α-helix in the TOPRIM domain and the DNA backbone, significantly contributing to Top2 function. This work uncovers a previously unrecognized role of the Tower domain, highlighting its involvement in anchoring and releasing the TOPRIM domain, thus priming Top2 for DNA binding and cleavage.

Graphical Abstract

Graphical Abstract

National Science and Technology Council 10.13039/100020595 111-2326-B-002-021 112-2326-B-002-010 111-2113-M-002-014-MY3 National Taiwan University 10.13039/501100006477 112L891605 Taiwan Biotechnological Foundation
==== Body
pmcIntroduction

DNA topoisomerases (Tops) are ubiquitous enzymes essential to all three domains of life (1–6). These enzymes resolve topological issues arising from cellular DNA transactions, such as replication, transcription, repair, and chromosome segregation (1–8). A shared feature across all types of Tops is their capability to reversibly cleave and religate the DNA backbone, thus enabling the manipulation of DNA topology without compromising the structural integrity of the genome (9–11). Depending on whether they cleave single-stranded DNA (ssDNA) or double-stranded DNA (dsDNA), Tops are divided into type I and type II, respectively (11). Each type is further categorized into subtypes (IA, IB, and IC for type I; IIA and IIB for type II), based on sequence divergence, structural characteristics, and mechanistic distinctions (12–15). Members of the type IB and IC subfamilies effectively release DNA superhelical stress by introducing a nick in one DNA strand, thereby allowing the nicked strand to undergo controlled rotation around the uncut strand (16–18). In contrast, type IA and type II enzymes alter DNA topology through a so-called ‘strand passage mechanism’ (10,14,19–21), where they transiently break and introduce an opening (or gate) in ssDNA or dsDNA (termed the gate or G-segment DNA), respectively, and subsequently transporting another DNA strand or duplex (termed the transfer or T-segment DNA) through the gate (20,22,23). Since reversible DNA cleavage is central to all Tops, a thorough understanding of their catalytic mechanisms necessitates detailed knowledge of this process. The main focus of this study is to provide a more complete picture on how the assembly of the DNA cleavage-religation center is achieved in type IIA Tops (Top2s).

Eukaryotic Top2 primarily functions as a homodimer, assembled from two identical multidomain subunits, with each subunit typically ranging from 1400 to 1600 residues, mainly due to species and isoform-specific variations in the C-terminal domain (CTD) (24–27). Starting from the N-terminus, a single subunit comprises the following domains: the GHKL ATPase domain (28), transducer domain (29), topoisomerase-primase (TOPRIM) domain (30,31), winged-helix (WHD) domain (31,32), Tower domain (31,33), coiled-coil region and primary dimerization domain (31), and ends with an intrinsically disordered and less conserved CTD (25,34,35). With a 2-fold symmetric overall architecture, Top2 incorporates a DNA cleavage-religation center constituted of two TOPRIM domains, two WHD domains, and two Tower domains, suitable for introducing a double-stranded DNA break (DSB) (24,25,33,36–45). The TOPRIM domain harbors a metal-binding DxD motif for catalytic Mg2+ binding (24,25,33,36–48) and two conserved DNA-binding elements, the EGDSA and PLRGK motifs (49,50). The WHD domain, possessing a winged helix-turn-helix motif and various conserved and surface-exposed residues, cooperates with TOPRIM and Tower domains to establish a DNA-binding groove that engages the G-segment DNA (24,25,33,36–45). Once bound, the formation of extensive non-covalent interactions induces bending in the G-segment DNA and drives its transition to an A-form-like conformation within the bound region (33,45,51). Furthermore, the WHD domain houses the active site tyrosine residue which, in conjunction with the Mg2+ provided by the TOPRIM domain of the opposite subunit, catalyzes the reversible transesterification reaction (9–11,41,46,47). The formation of a transient DSB is achieved through the attack on a pair of phosphodiester bonds, staggered by 4 base pairs on opposite strands, by the two symmetrically arranged active site tyrosine residues (36,37,39–44). This protein-linked DSB unlocks the DNA backbone and provides a gateway (20,48), termed the DNA-gate of Top2, through which another DNA duplex can be transported to modulate DNA topology. During DNA passage, Top2 undergoes extensive conformational changes, which include the coordinated formation and subsequent disruption of contacts between different domains of the enzyme (20,23,46,48). This process, characterized by a highly ordered sequence of structural rearrangements, enables the directional transport of T-segment DNA through the cleaved G-segment, accomplishing changes in DNA topology.

Over the past two decades, structural studies have offered atomic resolution insights into various key states of Top2’s catalytic cycle. These results have substantially broadened our comprehension of several G-segment DNA-bound states, including the Top2 DNA-gate in pre-cleavage (44), post-cleavage but DNA-gate closed (24,25,33,36–45), and post-cleavage and DNA-gate partially opened states (48). These structures reveal how Top2 interacts with the G-segment DNA, how the active site tyrosine and the catalytic Mg2+ are positioned relative to the scissile phosphodiester bond to perform cleavage, and how the two halves of the cleaved G-segment DNA move apart to permit the passage of T-segment DNA. Further, the N-terminal ATPase domains, recognized as the entrance or N-gate region, have been observed in both nucleotide-free (open) (39) and AMPPNP-bound (closed) (24,25,38,52–56) states, suggesting a role in capturing an incoming T-segment DNA (20,23,57). Moreover, the exit or C-gate region, formed by the primary dimerization domains, can separate to enable the final release of the T-segment DNA (33,40,45). This collection of structural data provides a detailed, step-by-step visualization of how the coordinated manipulation of inter-domain interfaces allows the cleavage of G-segment DNA and opening of DNA-gate for T-segment DNA transit. However, the precise structural state that commits Top2 to G-segment DNA binding and the key inter-domain interfaces associated with this state remain poorly understood.

The DNA binding and cleavage core (DBCC; Figure 1A) of a eukaryotic Top2, in its DNA-free, apo form, has been observed in two distinct conformational states (PDB ID: 1BGW and 1BJT) (31,58). Nevertheless, the configurations of the various DNA-interacting domains in these states differ substantially from the G-segment DNA-bound binary complex (24,25,33,36,43,45). Specifically, the two WHD domains, instead of being in direct contact to form a continuous DNA-binding surface, are widely separated. Furthermore, their potential for association with DNA is impeded due to the presence of TOPRIM domains in the intervening space. This implies that substantial structural rearrangement would be necessary for apo Top2 to engage in G-segment DNA-binding. A conformation seen in the DBCC of prokaryotic Top2, characterized by directly contacting WHD domains and the absence of intervening TOPRIM domains, appears more suitable for G-segment DNA binding. Yet, the prokaryotic DBCC constructs used for structural analyses were generated by introducing artificial linkers to connect the TOPRIM and WHD domains that are located in different subunits (37,40,59), thus the functional relevance of this conformation requires further investigation. Moreover, the structural characteristics of this particular conformational state remain uncharacterized.

Figure 1. A functionally relevant conformational state of hTop2core apo form reveals an exposed DNA-binding groove. (A) Schematic representation of hTop2 isoforms. This diagram illustrates the linear domain organization of hTop2 isoforms. The segment studied, corresponding to the DNA binding and cleavage core (DBCC) of hTop2, spans residues 431–1193 in hTop2α and 445–1201 in hTop2β. The metal-binding motif (DXD) in the TOPRIM domain and the active site tyrosine (Y) in the WHD domain are highlighted. (B) Orthogonal views of hTop2αcore (top) and hTop2βcore (bottom) structures showing that this conformational state is primed for engagement with incoming G-segment DNA. The domains within each structure are color-coded for distinction, with corresponding domains from the second monomer represented in lighter shades. The ‘prime’ (′) suffix denotes domains originating from the second monomer. This conformation is characterized by the interface formation between the TOPRIM and Tower′ domains, along with the symmetrical interface between TOPRIM′ and Tower domains. In both hTop2core apo structures, the DNA-binding groove measures approximately 27 Å in width and 26 Å in depth, dimensions compatible with a B-form DNA duplex. A proportionally scaled model of B-form DNA (PDB ID: 1BNA) is included for reference.

In this study, we report the crystal structures of the DBCC of human Top2α (hTop2αcore) and Top2β (hTop2βcore) in a conformational state competent for DNA binding. This state is characterized by the in trans assembly of two symmetry-related interfaces, each formed between the Tower domain of one subunit and the TOPRIM domain of another subunit. Such an arrangement results in the formation of an extensive DNA-binding groove, optimally configured to accommodate an incoming DNA duplex. Structural superimposition further revealed that subsequent G-segment DNA-induced conformational changes in Top2, including the detachment of the TOPRIM domain from the Tower and its repositioning toward the bound DNA, allow the bound duplex to be tightly grasped and ready for cleavage and religation. Notably, we identified an unprecedented type of protein-DNA interaction, in which the C-terminus of an α-helix in the functionally unrecognized helical bundle region of TOPRIM domain binds the backbone of G-segment through the bridging of a strictly conserved arginine residue, contributing significantly to Top2 function. This work reveals how the Tower-TOPRIM interface formed in trans primes the enzyme for G-segment binding, and how the DNA binding-induced disruption of this interface initiates the formation of Top2’s DNA cleavage/religation center.

Materials and methods

Construction, expression and purification of recombinant proteins

The DNA sequences encoding hTop2βcore (residues 445–1201) and hTop2αcore (residues 429–1188) were cloned into the pET51b vector. Subsequently, these expression plasmids were transformed into Escherichia coli BL21 Star (DE3) pLysS cells for protein expression. The overexpression and purification of these recombinant proteins followed established protocols (36,43). The purified proteins were stored in a buffer comprising 30 mM Tris–HCl (pH 7.5), 70 mM NaCl, and 2 mM β-mercaptoethanol (β-ME) for subsequent crystallization experiments.

For the expression of hTop2βΔCTD (residues 45–1201, a wild-type proxy demonstrating relaxation and decatenation activities similar to the full-length protein (60)) and its mutants (hTop2βΔCTD(R677A) and hTop2βΔCTD(R677K)), the respective YEphTOP2B plasmids were electroporated into Saccharomyces cerevisiae BCY123 (MATα pep3::HIS3 prb1::LEU2 bar1::HISG lys2::GAL1/10-GAL4 can1 ade2 trp1 ura3 his3 leu2-3 112) and purified following previously described methods (43). These purified proteins were adjusted to a concentration of 0.02 mg/ml and stored in a buffer containing 50 mM Tris–HCl (pH 7.5), 200 mM KCl, 5 mM MnCl2 and 2 mM β-ME for gel-shift assay and DNA relaxation/cleavage assays.

For in vivo yeast top2ts complementation assays, full-length hTop2β, hTop2α and yeast Top2 were expressed using plasmids pKN28, pMJ1 and pDED1-yTop2, respectively (61,62). The construction of all Top2 mutants employed in this study was carried out using the Gibson assembly method (NEB), with specific primers detailed in Supplementary Table S1.

Protein crystallization

Crystals of both apo hTop2αcore and hTop2βcore were unexpectedly grown from conditions that included a 20-bp double-stranded DNA (dsDNA) and the anticancer drug doxorubicin. Contrary to our initial aim of obtaining crystals of the Top2-DNA-doxorubicin ternary complex, we instead successfully produced crystals of the apo forms of hTop2αcore and hTop2βcore. The crystallization protocol began with pooling and concentrating fractions from SD200 gel-filtration chromatography to approximately 2 ml. Concentration was achieved at 1455 g, at 4°C, utilizing a 100 kDa ultrafiltration centrifugal filter. This concentrated sample was then mixed with an equal volume of gel-filtration buffer (30 mM Tris–HCl (pH 7.5), 70 mM NaCl, 2 mM β-ME) supplemented with 4 mM MgCl2. Following overnight incubation on ice, the protein sample was further concentrated to 8.5 mg/ml and subsequently combined with a 1.2-fold molar excess of a 20-bp dsDNA (sequence: 5′-AGCCGAGCTGCAGCTCGGCT-3′, prepared in gel-filtration buffer) and 2 mM doxorubicin (dissolved in DMSO). To eliminate any aggregated protein components, the sample mixture was centrifuged at 21500 g for 30 min at 4°C.

Crystallization was carried out using the hanging-drop vapor diffusion method at 4°C. This involved mixing 1 μl of the sample mixture with an equal volume of the reservoir solution and equilibrating against 200 μl of this reservoir solution. The reservoir solutions were composed of 0.1 M ammonium sulfate, 0.1 M citrate buffer (pH 5.5), and 12% PEG4000 for hTop2αcore, and 0.2 M ammonium acetate, 0.01 M calcium chloride dihydrate, 0.05 M sodium cacodylate trihydrate (pH 6.5), and 10% PEG4000 for hTop2βcore. Subsequently, both types of crystals were transferred into their respective cryoprotectant solutions, which consisted of the corresponding reservoir solutions supplemented with 30% 2-methyl-2,4-pentanediol (MPD). These crystals were then flash-frozen in liquid nitrogen to prepare them for data collection.

X-ray diffraction data collection and structure determination

All diffraction data sets were collected at beamline BL15A1 operated by the National Synchrotron Radiation Research Center (NSRRC), Taiwan, and were processed using the HKL2000 software suite (63). The structures of apo hTop2αcore and hTop2βcore were determined by molecular replacement using PHENIX AutoMR (64). For apo hTop2αcore, the TOPRIM domain-deleted structure of hTop2αcore, as observed in the hTop2αcore-DNA-etoposide complex (PDB ID: 5GWK) (43), was used as the search model. For apo hTop2βcore, the TOPRIM domain-deleted structure of hTop2βcore, as seen in the hTop2βcore-DNA-etoposide complex (PDB ID: 3QX3) (36), was used as the search model. In both cases, the presence of unbiased Fo – Fc electron density using MR-derived phases enabled unambiguous incorporation of the TOPRIM domain using the Coot software (65). Subsequent steps involved iterative rounds of manual model rebuilding and refinement employing both Coot and PHENIX. Detailed data collection and refinement parameters are listed in Supplementary Table S2.

Supercoil relaxation assay

For the supercoil relaxation assay (in 20 μl setting), 150 ng of negatively supercoiled pRYG plasmid was used as the substrate and incubated with varying amounts of either wild-type hTop2βΔCTD or its mutant variants. The reactions were carried out in a previously established buffer system, which includes 10 mM Tris–HCl (pH 8.0), 100 mM KCl, 0.1 mM EDTA, 5 mM MgCl2, 1 mM ATP, 5% DMSO and 0.01 mg/ml BSA (43). The reaction containing an additional 0.5 mM etoposide, a poison/inhibitor of human Top2s, was conducted to show the observed relaxation activity is Top2-specific. The reaction was initiated by the addition of Top2 and incubated at 37°C for 30 min. Termination of the reaction was achieved by adding a mixture of 0.125% SDS and 12.5 mM EDTA, incubated for 10 min. To ensure the clear visualization of DNA bands without interference from Top2 binding, proteinase K was added at a concentration of 1 μg/μl, and the mixture was further incubated at 37°C for an additional 30 min. The resulting mixtures were resolved by electrophoresis on a 0.8% TBE agarose gel at 50 V for 75 min, followed by post-staining with ethidium bromide (EtBr). For precise identification of various DNA forms, such as negatively supercoiled, relaxation intermediates, fully relaxed, linearized due to dsDNA breaks, and nicked DNA, the samples were also analyzed on a 0.8% TBE agarose gel containing 0.5 μg/ml EtBr, electrophoresed at 50 V for 65 min. Gels were visualized under UV light, and the images were captured and analyzed using ImageJ software.

Electrophoretic mobility shift assay (EMSA)

For the gel-shift assay conducted in a 20 μl reaction volume, 100 ng of 30bp dsDNA labeled with FAM (5′-GAAGATGACGAGCTGCAGTTAGATAGAGAA-3′-FAM; 5′-TTCTCTATCTAACTGCAGCTCGTCATCTTC-3′) was employed as the substrate. This DNA was incubated with varying concentrations of either wild-type hTop2βΔCTD or its mutant variants. The reaction was initiated upon addition of Top2 and proceeded at 37°C for 30 min under the same buffer conditions as those used in the supercoil relaxation assay. The resultant mixtures, in a 5% glycerol solution, were separated via electrophoresis on an 8% acrylamide gel utilizing 0.5× TBE buffer at 100 V for 60 min. Gel visualization was achieved through a FITC filter, and subsequent image acquisition and analysis were performed using ImageJ software.

In vivo functional complementation using a yeast top2ts strain

Complementation assays were performed using the S. cerevisiae strain JN362at2-4 (MATa ura3-52 leu2 trp1 his7 ade1-2 ISE2 top2-4), which possesses a temperature-sensitive t2-4 top2 allele exhibiting a loss-of-function phenotype at 34°C (61). Plasmids for vector-alone negative controls, namely pDED1-vec and pMJ1-vec, were generated by HindIII digestion of pDED1-yTop2WT (containing wild-type yTop2) and pMJ1-hTop2αWT (containing wild-type hTop2α), respectively (66). Additionally, for the construction of the pKN28-vec, PacI digestion was employed on pKN28-hTop2βWT (containing wild-type hTop2β), followed by re-ligation to excise the respective Top2 coding sequences. Electro-competent JN362at2-4 yeast cells were prepared using the lithium acetate method and cultured at 25°C. A range of plasmids, including pKN28-vec, pKN28-hTop2β(WT), pKN28-hTop2β(K450A/R494A), pKN28-hTop2β(R677K), pKN28-hTop2β(R677A), pMJ1-vec, pMJ1-hTop2α(WT), pMJ1-hTop2α(R434A/R478A), pMJ1-hTop2α(R661K), pMJ1-hTop2α(R661A), pDED1-vec, pDED1-yTop2(WT), pDED1-yTop2(R422A/R466A), pDED1-yTop2(R650K), and pDED1-yTop2(R650A) were individually electroporated into JN362at2-4. Selection was performed on solid SC-Ura media, with incubation at 25°C for 3–4 days. Single colonies were then inoculated into SC-Ura liquid medium and allowed to grow overnight at 25°C. Post centrifugation and PBS wash, the cell pellets were resuspended in PBS and adjusted to an OD600 of 1.0. This was followed by the preparation of 10-fold serial dilutions, which were spotted onto both YPD and SC-Ura plates (66). After air-drying, the plates were incubated at room temperature and the non-permissive temperature of 34°C. Colony viability, growth, and morphology were assessed using images captured after 3–4 days of incubation.

Results and discussion

An on-pathway conformational state of eukaryotic Top2 competent for engaging G-segment DNA

In this study, we produced crystals of apo hTop2αcore and hTop2βcore in new crystal forms (Supplementary Table S2). The emergence of these apo hTop2 crystals was unexpected, as our crystallization trials were originally aimed at exploring G-segment DNA binding by Top2. Nevertheless, the serendipitous production of these apo Top2 crystals provides significant structural insights into the conformational landscape of Top2, the state of Top2 poised for G-segment DNA engagement, and the structural transformations leading to the assembly of Top2’s DNA cleavage-religation center. The corresponding crystal structures, designated herein as hTop2αcore(R3) and hTop2βcore(P21212), were determined at resolutions of 2.67 and 2.8 Å, respectively. The asymmetric unit of the rhombohedral (R3) crystal comprises four monomers that assemble into two functional hTop2αcore dimers; the orthorhombic (P21212) crystal has an asymmetric unit with two monomers, forming a single hTop2βcore dimer. Together, we obtained three crystallographically independent observations of eukaryotic Top2. Structural superimposition revealed that these three Top2 dimer structures resemble each other closely and thus correspond to the same quaternary conformational state (Supplementary Figure S1; Figure 1). Remarkably, this state approximates the conformation observed in the artificially constructed prokaryotic Top2s (37,40,59), for which the functional relevance and structural characteristics have yet to be addressed. Given that the identical quaternary structure can be obtained from the two distinct eukaryotic Top2 isoforms, and even from prokaryotic Top2 DBCC when artificial linkers are present, this structural state most likely represents a functionally relevant and energetically accessible conformation of Top2 that warrants thorough analysis.

Compared to previously published apo structures of eukaryotic Top2s (31,58), the quaternary conformation observed in hTop2αcore(R3) and hTop2βcore(P21212) is distinctive (Figure 1; Supplementary Figure S2). A defining feature of this conformational state is the structural rearrangement of the two TOPRIM domains relative to the WHD and Tower domains (Supplementary Figure S2). In apo Top2 structures of yeast (31,58), the TOPRIM domains juxtapose the dimer's two-fold axis and are positioned between the two WHD domains. This configuration keeps the WHD domains, which constitute a considerable portion of the G-segment DNA binding surface, apart and physically impedes their association with incoming DNA. Consequently, a relocation of the TOPRIM domains is expected to be essential for Top2 to engage with the G-segment DNA. In contrast, in hTop2αcore(R3) and hTop2βcore(P21212), the TOPRIM domains have shifted away from the structural dyad and are stabilized in new positions through extensive interactions with the opposing Tower domains, forming a pair of hitherto unobserved dimer interfaces, which we have designated as the TOPRIM-Tower′ interface (Figure 2A). (The prime symbol next to Tower indicates that the interacting domains belong to different monomers.) The formation of these two symmetry-related TOPRIM-Tower′ interfaces results in the burial of approximately 2000 Å² of accessible surface area, nearly doubling that of the interface formed by the C-gate dimerization domain. Superimposition analysis (Figure 2A, enlarged) and structure-based sequence alignment (Figure 2B) reveal that the structural features and the majority of residues at the TOPRIM-Tower′ interface are conserved among Top2s. Furthermore, residues participating in interface formation adopt identical side-chain rotamer conformations (Supplementary Figure S3). Together, these observations strongly implicate the functional importance of this interface. This conformational state is likely to represent an initial, on-pathway state in the catalytic cycle of Top2 (Supplementary Figure S4). As the TOPRIM domains move aside, the two WHD domains can come together, creating a continuous and extensive positively-charged groove poised to accommodate the incoming G-segment DNA. We propose that the establishment of the TOPRIM-Tower′ interfaces unveils the G-segment DNA binding site and thus primes the enzyme for subsequent catalytic actions.

Figure 2. Structural-sequence conservation and functional significance of the TOPRIM-Tower′ interface in Top2. (A) Conservation of the TOPRIM-Tower′ interface structure. This panel illustrates the alignment of apo hTop2α and hTop2β structures from our study with those of bacterial Top2 DBCCs, including E. coli gyrase (PDB ID: 3NUH (59)), A. baumannii Top-IV (PDB ID: 2XKJ (40)), and S. aureus gyrase (PDB ID: 2XCO (37)). Note that the bacterial structures were artificially constructed by linking the TOPRIM and WHD domains together. The structural conservation of the TOPRIM-Tower′ interface across Top2s is highlighted, with interface residues displayed in spherical representations and are labeled in Supplementary Figure S8. (B) Residue conservation at the TOPRIM-Tower′ Interface. An ‘open book’ view shows the conservation of interface-forming residues. The residues are color-coded based on their conservation level, as determined by the ConSurf server (79). Residue numbering is indicated in blue for hTop2α and in red for hTop2β. (C) Impact of interface mutations on hTop2β relaxation activity. This panel depicts the supercoil relaxation activity of hTop2β wild-type proxy (hTop2βΔCTD) and its double mutant hTop2βΔCTD(K450A/R494A) in an ATP-dependent relaxation assay, the locations of these residues are highlighted by green lines in panel (B). Reactions with VP-16 (etoposide), a potent poison/inhibitor of human Top2s, were included to demonstrate that the formation of fully relaxed and partially relaxed DNA intermediates is dependent on hTop2β activity. The right panel shows the quantification of gel images from three independent experiments. Data are presented as mean ± SEM. ****Adjusted P value < 0.0001, ***P = 0.0002. ‘SC’ denotes negatively supercoiled DNA, while ‘L’ indicates linearized DNA. (D) In vivo functional analysis of interface mutations on hTop2β, hTop2α and yTop2. Plasmids encoding intact wild-type Top2 along with their double mutant (hTop2β(K450A/R494A), hTop2α(R434A/R478A) and yTop2(R422A/R466A), were introduced into the yeast JN362at2-4 strain, which carries a temperature-sensitive yTop2 allele (61). Following 10-fold serial dilutions, yeast growth was monitored at both permissive room temperature and the non-permissive temperature of 34°C.

Mutation-induced destabilization of the TOPRIM-Tower′ interface impairs Top2 function

To validate the functional significance of this conformational state, characterized by the formation of the TOPRIM-Tower′ interfaces (Figure 2A), we performed site-directed mutagenesis to destabilize this interface. Examination of the interface's constituent residues (Figure 2B) identified K450 and R494 on hTop2β (R434 and R478 of hTop2ɑ; R422A and R466A of yTop2), among several others, in the TOPRIM domain as key stabilizers of this interface, forming an interacting network with residues D458, G490, and H977′ (L441, D442, E941′, Y957′, H961′ of hTop2ɑ) (Supplementary Figure S5). Three lines of evidence suggest that mutating these residues will specifically perturb the TOPRIM-Tower′ interface without compromising the structural integrity or other known functions of the TOPRIM domain. Firstly, K450 and R494 are surface-located on the TOPRIM domain. Secondly, they are not implicated in DNA binding or metal chelation. Consequently, substituting these residues with alanine is anticipated to selectively weaken the TOPRIM-Tower′ interface while preserving other TOPRIM domain properties. In contrast to the robust relaxation activity of wild-type Top2, the double mutant hTop2βΔCTD(K450A/R494A) displayed greatly reduced activity both in vitro andin vivo (Figure 2C, D). To further assess the impact of these mutations on Top2 function, we conducted an electrophoretic mobility shift assay using a 30 bp double-stranded DNA segment labeled with a FAM fluorescent group (Supplementary Figure S6). The result demonstrated that while the wild-type Top2β exhibited a dose-dependent accumulation of the Top2-DNA complex, the double mutant displayed no such complex formation. This finding supports our hypothesis that the TOPRIM-Tower′ interface is crucial in preparing Top2 for subsequent catalytic events (Supplementary Figure S4). Before this interface is formed, the TOPRIM domain likely exhibits excessive flexibility and adopts multiple orientations, as observed in yeast Top2 structures (31,58), which could hinder G-segment DNA binding.

Our structural and functional analysis of the Tower domain reveals its multifaceted role in Top2’s function. Previous studies have shown that the Tower domain acts as an auxiliary DNA-binding module, engaging with the bent G-segment DNA (33,45). Additionally, a narrow surface groove on the Tower domain of hTop2α guides a loop from the intrinsically disordered C-terminal domain to interact with the G-segment DNA, influencing catalytic efficiency (25). Moreover, the recently published cryo-EM structure of full-length E. coli gyrase has revealed an additional function of the Tower domain in contributing to T-segment DNA binding (67). In this study, we further propose a crucial role for the Tower domain in anchoring the TOPRIM domain, thereby enabling Top2 to effectively engage G-segment DNA, which in turn facilitates the onset of the ensuing catalytic cycle. It should be emphasized that a large number of Tower domain residues are involved in the formation of the TOPRIM-Tower′ interface, contrasting sharply with the fewer residues that engage in interactions with G-segment DNA or the Top2 CTD (in the case of hTop2α) (Supplementary Figure S7) (25,43,45). Such disparity underscores the functional importance of the formation of the TOPRIM-Tower′ interface. Consistent with this notion, mutations at residues located within the TOPRIM-Tower′ interface, or those that potentially destabilize this interface, have been linked to various diseases (Supplementary Figure S8) (68–70). Specifically, the S483L and A485P mutations are associated with Hoffman syndrome and BILU syndrome, respectively. S483 directly participates in forming the TOPRIM-Tower′ interface (Figure 2B; Supplementary Figure S3). While A485, also on the Bα2 helix alongside S483, does not directly engage in the interface interactions, its mutation to a helix-disrupting proline could undermine the structural integrity of the TOPRIM-Tower′ interface, thereby compromising the DNA-binding capacity and catalytic activity of hTop2β. Moreover, several drug-resistant mutations (G474A/V475M and G474C/S in hTop2α, along with E430K, G465D, R466K/T, G919D, M933I, E934K and H937Y in yTop2) were identified at the TOPRIM-Tower′ interface (66). This suggests that destabilization of this interface may diminish Top2’s catalytic activity, thereby conferring drug resistance by reducing the formation of the cleavage complex induced by Top2 poisons. In conclusion, our findings characterize a functionally relevant structural state of Top2, indicative of a G-segment DNA binding-competent conformation. By analyzing this conformation alongside the G-segment DNA-bound states (36,37,43,44,48), we were able to elucidate the conformational transitions leading to the assembly of Top2’s DNA cleavage-religation center.

The assembly of Top2’s DNA cleavage-religation center requires extensive rearrangement of molecular interfaces

The conformational state adopted by human Top2s, as characterized in this study, sheds light on the structural features marking the initial phase of the assembly of its DNA cleavage-religation center. When integrated with various published structures of DNA-bound human Top2s (24,25,33,36–45), corresponding to the end point of this process, we are in position to delineate/simulate a comprehensive sequence of conformational transitions that Top2 undergoes during the assembly process. To this end, we utilized structural superimposition and morphing between the starting and end states to unveil these changes (Figure 3; Supplementary Movies). The structural transformations triggered by G-segment DNA binding comprise two sequential events. Firstly, the disruption of the TOPRIM-Tower′ interface, initiated by G-segment DNA binding, leads to the detachment of the TOPRIM domain from the Tower′ domain. Mechanistically, this separation is driven by the interaction of the G-segment DNA with helix Aɑ4 of the WHD domain (and its counterpart, helix Aɑ4′ from the opposing subunit), a primary DNA-binding element of Top2 (Figure 4A). Specifically, the helix Aɑ4 adapts into the major groove of the DNA through an induced-fit mechanism. As a result, the symmetrically arranged WHD domains undergo lateral sliding, moving away from each other in a direction orthogonal to the structural dyad (Figure 4B,C). This movement drives the two monomers apart, consequently weakening the TOPRIM-Tower′ interface formed in trans and leading to the detachment of the TOPRIM from the Tower domain (Figure 4D). This detachment frees the TOPRIM domain, allowing it to undergo a rigid body-like movement to form extensive interactions with the bound G-segment DNA (Figure 3). These interactions not only secure the DNA firmly within the binding groove but also position the metal-binding DxD motif near the active site tyrosine residue and scissile phosphodiester bond. Collectively, these structural rearrangements facilitate the recruitment of Mg2+, bringing all catalytic components into alignment and culminating in the formation of the Top2’s DNA cleavage-religation center, ready for the subsequent metal-assisted DNA cleavage and religation.

Figure 3. DNA-binding-induced structural rearrangement facilitates the formation of Top2’s DNA cleavage-religation center. (A) Superimposition of hTop2αcore structures. This panel compares the apo form of hTop2αcore (left, PDB ID: 8W50, from this study, colored orange) with its G-segment DNA-bound form (middle, PDB ID: 5GWK (43), colored blue). Structural superimposition reveals an extensive rearrangement of the TOPRIM domain upon G-segment DNA binding, culminating in the formation of the DNA cleavage-religation center. (B) Superimposition of hTop2βcore structures. A similar comparison between the apo form of hTop2βcore (left, PDB ID: 8KE7, from this study, colored pink) and its G-segment DNA-bound form (middle, PDB ID: 3QX3 (36), colored blue) underscores an analogous conformational change, as observed in panel A, critical for assembling the Top2’s DNA cleavage-religation center. The TOPRIM domain's position is accentuated with a surface representation to clearly depict its movement. Superimpositions of the two hTop2core conformational states were performed using PyMol by aligning regions that span the WHD and Tower domains. Triangular markers highlight the interface in the apo structures (left) and the gap between the TOPRIM and Tower' domains in the DNA-bound structures (middle).

Figure 4. G-segment DNA binding triggers lateral sliding in WHD domains, leading to TOPRIM detachment from the Tower' domain. (A) Structural changes of the G-segment DNA-binding groove. This panel presents top views of the G-segment DNA-binding groove in its apo form (left, PDB ID: 8KE7, this study) and subsequent DNA-bound states along Top2’s catalytic cycle (second left to right, referenced by PDB accession codes (36,37,44,48)). The van der Waals surfaces are rendered with 60% transparency, and WHD domains are depicted in cartoon representation for each structure. The interfaces formed by the TOPRIM, WHD, and Tower domains, highlighted in pink/purple, are present except for the DNA-gate open structure (rightmost; PDB ID: 5ZEN (48)). These structures provide a sequential visualization of the structural transitions that occur upon G-segment DNA binding and cleavage. (B) Enlarged view of the relative WHD domain movement. Focusing on the encircled region from panel (A), we enlarge each state to detail the DNA binding-induced lateral sliding of the two WHD domains relative to one another. Arrowed vectors, created in PyMOL, mark helix Aα4 and its symmetry-related Aα4′, facilitating recognition of the relative movements. The distances between the N-termini of Aα4 and Aα4′ are annotated. A gray backdrop serves as a reference for comparing the relative positions of Aα3/Aα4 and Aα3′/Aα4′ in each state. (C) DNA binding-induced separation of WHD domains. By aligning the 2-fold axes of regions shown in panel (B), the trajectory of the WHD domains’ separation induced by DNA binding is illustrated. (D) DNA binding-induced TOPRIM detachment. Structural superimposition demonstrates the detachment of TOPRIM from the opposing Tower' domain upon DNA binding. Different catalytic states are color-coded as indicated.

The conformational changes and domain movement trajectories discussed above are consistently observed not only in our analyses of both hTop2αcore and hTop2βcore (Figure 3; Supplementary Movies) but also in prokaryotic Top2s. Specifically, cryo-EM structures of full-length E. coli gyrase, resolved without artificial linkers, display various conformational states in the presence of DNA (38,67). These states correlate with the binary complex, pre-cleavage complex, or cleavage complex (Figure 4A). Comparing these cryo-EM structures to prokaryotic Top2 DBCC apo structures, which were obtained with artificial linkers (37,40,59), reveals that the TOPRIM, WHD, and Tower domains follow a similar trajectory during the assembly of the DNA cleavage-religation center (Figure 4; Supplementary Movies). This observation underscores a unified structural framework that is applicable across all Top2 isoforms, highlighting the conserved nature of these mechanisms.

Unveiling a previously unknown type of protein-DNA contact mediated by a conserved arginine in the TOPRIM-DNA interface

The DNA binding-induced repositioning of the TOPRIM domain, as previously described, allows it to interact extensively with the G-segment DNA (Figure 3; Supplementary Movies). Some TOPRIM-DNA contacts have been documented in earlier studies (24,25,33,36–45). For instance, the EGDSA motif interacts with the + 1/+4 base pair, the PLRGK motif binds to the -1/+5 base pair, and the metal-chelating DxD motif strengthens the Top2-DNA interaction through a water-mediated hydrogen bond with the -1 nucleotide. Unexpectedly, our analysis revealed a hitherto unrecognized interaction between the TOPRIM domain and G-segment DNA, established between the C-terminus of helix Bα9 and the phosphodiester linking the +8 and +9 nucleotides (Figure 5A, B). Bα9-DNA interaction is mediated by a bridging arginine residue from the N-terminal portion of helix Bɑ10 (R661 in hTop2α and R677 in hTop2β). Remarkably, this arginine is strictly conserved across Top2s, ranging from bacteria to humans (Figure 5C). It should be noted that a conserved isoleucine residue intercalates and bends the bound G-segment DNA at this precise interaction site, a prerequisite for DNA cleavage (33,71). It is tempting to speculate that this arginine anchors the + 8 and + 9 nucleotides and potentially contributes to DNA bending (Figure 5A, D). Moreover, this arginine-mediated interaction is consistently present in all DNA-bound states of Top2, including binary complexes and cleavage complexes in both closed and partially opened states (Supplementary Figure S9) (36,37,44,48). On the contrary, some TOPRIM-DNA contacts, like those mediated by the PLRGK motif, manifest only post-DNA cleavage and are absent in the initial Top2-DNA binary complex. These findings suggest that this specific arginine-mediated protein-DNA interaction is likely functionally significant.

Figure 5. A previously undocumented type of protein-DNA interaction mediated by an arginine-capped helix C-terminus. (A) Involvement of an arginine-capped helix C-terminus in DNA-binding. In both hTop2α and hTop2β, the C-terminus of helix Bα9 is capped by a guanidino group from a strictly conserved arginine residue (R661 in hTop2α and R677 in hTop2β), located within the N-terminal region of Bα10. This capping appears to serve dual functions: it maintains the tertiary structure stability of TOPRIM during DNA-induced conformational changes, and critically, it facilitates engagement with the DNA backbone by counteracting the charge repulsion between the negatively charged C-terminus of Bα9 and the DNA. (B) Impact of arginine capping on the electrostatic surface potential of the C-terminus of Bɑ9. The electrostatic surface potential of the Bα9 C-terminus, when devoid of the capping arginine, is predominantly negative (top), a consequence of the helix dipole. The introduction of the arginine cap reverses the negative charge at the C-terminus of Bα9, enabling it to participate actively in DNA binding. (C) Conservation of the capping arginine across species. A multiple sequence alignment of Top2 sequences, from a range of species from bacteria to humans, demonstrates the strict conservation of this capping arginine residue (indicated by an asterisk). (D) The arginine-capped C-terminal helix engages the +8/+9 site of the DNA, coinciding with the intercalation site of I872 from the WHD domain, which plays a crucial role in DNA bending prior to cleavage. The interaction between the arginine-capped helix and the G-segment DNA is present throughout the entire catalytic cycle until the G-segment DNA is released (Supplementary Figure S9).

While the exploitation of helical dipoles for DNA binding is common in protein-DNA complexes (72,73), interactions of this type typically involve the positively charged N-terminus of an α-helix. In contrast, this arginine-bridged protein-DNA interaction uniquely features the C-terminus of helix Bα9 approaching and engaging with the acidic DNA backbone. This interaction is facilitated by an apparent charge reversal (Figure 5B), where the negative charge at the C-terminus of helix Bα9 is shielded and inverted by the positively charged guanidino group of the bridging arginine, enabling the capped C-terminus of an α-helix to function as a DNA-binding module. To the best of our knowledge, this is the first instance of the C-terminus of an α-helix being directly participated in DNA binding.

The arginine residue that caps the C-terminus of helix Bα9 and bridges the Bα9-DNA interaction is crucial for the catalytic and cellular function of Top2

The hypothesized functional significance of the strictly conserved arginine residue in bridging the interaction between the TOPRIM domain and G-segment DNA led us to further investigate its role. We introduced mutations at residue R677 of hTop2β, replacing it with alanine and lysine to generate hTop2βΔCTD(R677A) and hTop2βΔCTD(R677K) mutants, respectively. These mutants were then assessed for their ability to relax negatively supercoiled plasmid DNA (Figure 6A). Compared to hTop2βΔCTD, our ‘wild-type’ proxy that manifests relaxation and decatenation activities comparable to those of full-length hTop2β, the hTop2βΔCTD(R677A) mutant displayed almost negligible relaxation activity. The hTop2βΔCTD(R677K) mutant also exhibited a pronounced reduction in both activity and processivity, as evidenced by the diminished accumulation of relaxed plasmid DNA and the increased presence of relaxation intermediates. The marginally higher activity of the hTop2βΔCTD(R677K) mutant, compared to the hTop2βΔCTD(R677A), may be attributed to the positively charged lysine side-chain partially compensating for the arginine side-chain's helix capping function. Nevertheless, the marked decrease in the relaxation activity of the hTop2βΔCTD(R677K) mutant is consistent with the perception that lysine is a less effective helix capping residue (74). This result emphasizes the pivotal role of the arginine-mediated interaction between the C-terminus of helix Bα9 and G-segment DNA; even replacing the arginine with lysine profoundly disrupts Top2 function.

Figure 6. Crucial role of the conserved capping arginine in Top2 Function. (A) hTop2β capping arginine mutants' relaxation activities. This panel assesses the ATP-dependent relaxation activities of hTop2β wild-type proxy (hTop2βΔCTD) and the corresponding capping arginine mutants (hTop2βΔCTD(R677A) and hTop2βΔCTD(R677K)). ‘SC’ denotes negatively supercoiled, and ‘L’ represents linearized pRYG plasmid DNA. The R677A mutation completely abolishes hTop2β’s relaxation activity, while the R677K variant greatly impairs function. Lower panel shows the quantification of gel images from three independent experiments. Data are presented as mean ± SEM. ****Adjusted P value < 0.0001, ***P = 0.0007. (B) In vivo functional analysis of yTop2, hTop2α and hTop2β capping arginine mutants. Plasmids encoding intact wild-type yTop2, hTop2α and hTop2β, along with their capping arginine mutants (hTop2β(R677A), hTop2β(R677K), hTop2α(R661A), hTop2α(R661K), yTop2(R650A), yTop2(R650K)) were introduced into the yeast JN362at2-4 strain, which carries a temperature-sensitive yTop2 allele (61). Following 10-fold serial dilutions, yeast growth was monitored at both permissive room temperature and the non-permissive temperature of 34°C. Mutations in hTop2α (R661A and R661K) inhibited yeast growth at non-permissive temperatures. In the case of yTop2, only the R650K mutant displayed marginal growth at high cell density (10−2 dilution).

To further demonstrate the functional importance of this capping arginine, similar mutations were introduced into yeast Top2 (yTop2) and both human Top2 isoforms, resulting in the creation of six additional mutants: yTop2(R650A), yTop2(R650K), hTop2α(R661A), hTop2α(R661K), hTop2β(R677A) and hTop2β(R677K). The impact of these mutations on Top2 function was assessed in vivo using the yeast JN362at2-4 strain, which carries a temperature-sensitive yTop2 allele (61). The activity of the wild-type and mutant Top2 variants was evaluated by shifting the yeast to a non-permissive temperature (Figure 6B). While wild-type yTop2, hTop2α and hTop2β supported yeast growth under non-permissive conditions, all six mutants with the altered arginine residue failed to do so. This finding reiterates the indispensable nature of this capping arginine residue that is strictly conserved across all Top2s.

It is worth noting that in hTop2α, the lysine residue (K662) adjacent to the capping arginine (R661) can be SUMOylated (45,75). The attachment of a bulky SUMO group at K662 is expected to physically block interactions of both R661 and K662 with the G-segment DNA, thus inhibiting hTop2α function. Accordingly, our proposed functional significance of R661 aligns with observations that SUMOylation at K662 results in a loss of decatenation activity in hTop2α, and that de-SUMOylation is required to restore its function.

A detailed examination of the TOPRIM domain's structural features sheds light on why this capping arginine is critical for Top2 function. The TOPRIM domain of Top2 is composed of a Rossmann-like metal-binding fold, a Greek key extension, and a helix bundle region (Supplementary Figure S10). The Rossmann fold and helix bundle regions are interconnected by a stable core of interdigitated hydrophobic side-chains from conserved residues. As discussed earlier, structural comparison reveals that the TOPRIM domain undergoes a rigid body-like movement during G-segment DNA-induced structural rearrangement (Figure 3). These characteristics indicate that the TOPRIM domain functions as an integrated unit, with interactions at structurally rigid spots influencing the spatial positioning of the entire domain. We hypothesize that the arginine-capped Bα9, situated in the rigid helix bundle region, binds to the G-segment DNA, thus bringing the TOPRIM domain into proximity with the DNA. This interaction allows the remaining, more flexible metal- and DNA-binding elements to engage with the DNA, culminating in the assembly of Top2’s DNA cleavage-religation center. Disturbance of this arginine-mediated interaction between Bα9 and DNA likely prevents the TOPRIM domain from achieving the optimal position for DNA interaction and coordination with the catalytic tyrosine, ultimately impairing Top2 function. Consistent with the proposed functional significance of this region, the C-terminal tail of the GyrB subunit in bacterial Top2, residing near the helix bundle region of the TOPRIM domain, is also known to be essential for catalytic function (76,77).

Conclusion

Top2s are essential for resolving DNA supercoiling and entanglements, thereby playing pivotal roles in all aspects of cellular DNA transactions. These enzymes operate through a sophisticated mechanism, wherein a transient DSB is produced to allow the passage of another DNA segment. Previous structural studies have revealed that Top2 undergoes extensive conformational changes during its catalytic cycle; however, the nature of these catalytically essential transitions remains to be further explored. This work aims to elucidate how Top2 initially interacts with DNA and maneuvers through various conformational states, culminating in the assembly of Top2’s DNA cleavage-religation center. It particularly highlights functionally significant inter-domain interactions, DNA-binding-induced conformational changes, and the discovery of a unique type of protein-DNA interaction crucial for the enzyme's catalytic mechanism. Our study begins with identifying an on-pathway conformational state of Top2, pivotal for engaging G-segment DNA. This state, essential for initiating Top2’s catalytic cycle, is characterized by the formation of an interface between the Tower and TOPRIM domains. The establishment of this TOPRIM-Tower' interface anchors the otherwise flexible TOPRIM domains, thereby exposing a DNA-binding groove and priming Top2 for engagement with incoming G-segment DNA. This revelation underscores a previously unrecognized role of the Tower domain in anchoring the TOPRIM domain and preparing Top2 for DNA binding. Moreover, our observation that Top2 function can be impaired by mutation-induced destabilization of the TOPRIM-Tower' interface, along with its likely involvement in the catalytic cycles of bacterial Top2s (DNA gyrase and Top IV), suggests that targeting this interface with novel interfacial inhibitors (78) could bear medical implications. Structural superimposition and morphing analyses between this state and DNA-bound human Top2 structures have unveiled a series of essential conformational transitions for Top2’s function. These include the detachment and repositioning of the TOPRIM domain upon G-segment DNA binding, achieved through the disengagement of the Tower and TOPRIM domains followed by a subsequent rigid body-like movement of the TOPRIM domain. This leads to extensive interaction with the G-segment DNA, securing it within the binding groove. A critical discovery in our research is the identification of a previously undocumented type of protein–DNA contact, mediated by a strictly conserved arginine residue in the helical bundle region of the TOPRIM domain. This residue uniquely caps and bridges the C-terminus of helix Bα9 to the bound DNA. Our mutational analysis of this residue in hTop2β, as well as its yeast and human counterparts, underscores its indispensable role in Top2’s catalytic and cellular functions. This arginine's unique positioning and function represent a divergence from the common helical dipole mechanism typically observed in protein–DNA interactions, thus enriching our insight into the mechanisms of DNA binding. In summary, our research offers crucial insights into the structural basis for the assembly of Top2’s DNA cleavage-religation center. These findings not only deepen our comprehension of Top2’s structural and functional nuances but also have medical relevance, implicating the molecular basis of certain disease-related Top2 mutations and suggesting new avenues for developing Top2 inhibitors by targeting the TOPRIM-Tower′ interface of human and bacterial Top2s.

Supplementary Material

gkae657_Supplemental_Files

Acknowledgements

Portions of this research were carried out at beamlines 15A1 of the National Synchrotron Radiation Research Center (Taiwan). We are grateful to the technical supports from Prof. Shu-Chun Teng and colleagues, Graduate Institute of Microbiology, National Taiwan University. Plasmids pDED1-yTop2, pMJ1, and pKN28 and yeast strain JN362at2-4 were kindly provided by Prof. John Nitiss and Dr. Karen Nitiss (University of Illinois, Chicago).

Data availability

Atomic coordinates and structure factors have been deposited in the PDB with accession codes 8KE7 (hTop2βcore apo structure) and 8W50 (hTop2ɑcore apo structure).

Supplementary data

Supplementary Data are available at NAR Online.

Funding

National Science and Technology Council [111-2326-B-002-021, 112-2326-B-002-010, 111-2113-M-002-014-MY3]; National Taiwan University [112L891605]; Taiwan Biotechnological Foundation. Funding for open access charge: National Taiwan University.

Conflict of interest statement. None declared.

Notes

Present address: Shin-Fu Chen, Department of Biochemistry and Molecular Biology, McGovern Medical School, University of Texas Health Science Center at Houston, Houston, TX 77030, USA.
==== Refs
References

1. Wang J.C. Cellular roles of DNA topoisomerases: a molecular perspective. Nat. Rev. Mol. Cell Biol. 2002; 3 :430–440.12042765
2. Chen S.H. , ChanN.L., HsiehT.S. New mechanistic and functional insights into DNA topoisomerases. Annu. Rev. Biochem. 2013; 82 :139–170.23495937
3. Schoeffler A.J. , BergerJ.M. DNA topoisomerases: harnessing and constraining energy to govern chromosome topology. Q. Rev. Biophys. 2008; 41 :41–101.18755053
4. Vidmar V. , VayssieresM., LamourV. What's on the other side of the gate: a structural perspective on DNA gate opening of type IA and IIA DNA topoisomerases. Int. J. Mol. Sci. 2023; 24 :3986.36835394
5. Pommier Y. , SunY., HuangS.N., NitissJ.L. Roles of eukaryotic topoisomerases in transcription, replication and genomic stability. Nat. Rev. Mol. Cell Biol. 2016; 17 :703–721.27649880
6. McKie S.J. , NeumanK.C., MaxwellA. DNA topoisomerases: advances in understanding of cellular roles and multi-protein complexes via structure-function analysis. Bioessays. 2021; 43 :e2000286.33480441
7. Lee J.H. , BergerJ.M. Cell cycle-dependent control and roles of DNA topoisomerase II. Genes (Basel). 2019; 10 :859.31671531
8. Pommier Y. , NussenzweigA., TakedaS., AustinC. Human topoisomerases and their roles in genome stability and organization. Nat. Rev. Mol. Cell Biol. 2022; 23 :407–427.35228717
9. Deweese J.E. , OsheroffN. The DNA cleavage reaction of topoisomerase II: wolf in sheep's clothing. Nucleic Acids Res. 2009; 37 :738–748.19042970
10. Brown P.O. , CozzarelliN.R. Catenation and knotting of duplex DNA by type 1 topoisomerases: a mechanistic parallel with type 2 topoisomerases. Proc. Natl. Acad. Sci. U.S.A. 1981; 78 :843–847.6262776
11. Liu L.F. , LiuC.C., AlbertsB.M. Type II DNA topoisomerases: enzymes that can unknot a topologically knotted DNA molecule via a reversible double-strand break. Cell. 1980; 19 :697–707.6244895
12. Taneja B. , PatelA., SlesarevA., MondragonA. Structure of the N-terminal fragment of topoisomerase V reveals a new family of topoisomerases. EMBO J. 2006; 25 :398–408.16395333
13. Buhler C. , GadelleD., ForterreP., WangJ.C., BergeratA. Reconstitution of DNA topoisomerase VI of the thermophilic archaeon sulfolobus shibatae from subunits separately overexpressed in Escherichia coli. Nucleic Acids Res. 1998; 26 :5157–5162.9801313
14. Bergerat A. , de MassyB., GadelleD., VaroutasP.C., NicolasA., ForterreP. An atypical topoisomerase II from Archaea with implications for meiotic recombination. Nature. 1997; 386 :414–417.9121560
15. Wang J.C. DNA topoisomerases. Annu. Rev. Biochem. 1996; 65 :635–692.8811192
16. Taneja B. , SchnurrB., SlesarevA., MarkoJ.F., MondragonA. Topoisomerase V relaxes supercoiled DNA by a constrained swiveling mechanism. Proc. Natl. Acad. Sci. U.S.A. 2007; 104 :14670–14675.17804808
17. Koster D.A. , CroquetteV., DekkerC., ShumanS., DekkerN.H. Friction and torque govern the relaxation of DNA supercoils by eukaryotic topoisomerase IB. Nature. 2005; 434 :671–674.15800630
18. Stewart L. , RedinboM.R., QiuX., HolW.G., ChampouxJ.J. A model for the mechanism of human topoisomerase I. Science. 1998; 279 :1534–1541.9488652
19. Kampranis S.C. , BatesA.D., MaxwellA. A model for the mechanism of strand passage by DNA gyrase. Proc. Natl. Acad. Sci. U.S.A. 1999; 96 :8414–8419.10411889
20. Roca J. , BergerJ.M., HarrisonS.C., WangJ.C. DNA transport by a type II topoisomerase: direct evidence for a two-gate mechanism. Proc. Natl. Acad. Sci. U.S.A. 1996; 93 :4057–4062.8633016
21. Lima C.D. , WangJ.C., MondragonA. Three-dimensional structure of the 67K N-terminal fragment of E. coli DNA topoisomerase I. Nature. 1994; 367 :138–146.8114910
22. Mizuuchi K. , FisherL.M., O’DeaM.H., GellertM. DNA gyrase action involves the introduction of transient double-strand breaks into DNA. Proc. Natl. Acad. Sci. U.S.A. 1980; 77 :1847–1851.6246508
23. Roca J. , WangJ.C. The capture of a DNA double helix by an ATP-dependent protein clamp: a key step in DNA transport by type II DNA topoisomerases. Cell. 1992; 71 :833–840.1330327
24. Schmidt B.H. , OsheroffN., BergerJ.M. Structure of a topoisomerase II-DNA-nucleotide complex reveals a new control mechanism for ATPase activity. Nat. Struct. Mol. Biol. 2012; 19 :1147–1154.23022727
25. Vanden Broeck A. , LotzC., DrillienR., HaasL., BedezC., LamourV. Structural basis for allosteric regulation of Human topoisomerase IIalpha. Nat. Commun. 2021; 12 :2962.34016969
26. Tsai-Pflugfelder M. , LiuL.F., LiuA.A., TeweyK.M., Whang-PengJ., KnutsenT., HuebnerK., CroceC.M., WangJ.C. Cloning and sequencing of cDNA encoding human DNA topoisomerase II and localization of the gene to chromosome region 17q21-22. Proc. Natl. Acad. Sci. U.S.A. 1988; 85 :7177–7181.2845399
27. Chung T.D. , DrakeF.H., TanK.B., PerS.R., CrookeS.T., MirabelliC.K. Characterization and immunological identification of cDNA clones encoding two human DNA topoisomerase II isozymes. Proc. Natl. Acad. Sci. U.S.A. 1989; 86 :9431–9435.2556712
28. Dutta R. , InouyeM. GHKL, an emergent ATPase/kinase superfamily. Trends Biochem. Sci. 2000; 25 :24–28.10637609
29. Murzin A.G. A ribosomal protein module in EF-G and DNA gyrase. Nat. Struct. Biol. 1995; 2 :25–26.7719848
30. Aravind L. , LeipeD.D., KooninE.V. Toprim–a conserved catalytic domain in type IA and II topoisomerases, DnaG-type primases, OLD family nucleases and RecR proteins. Nucleic Acids Res. 1998; 26 :4205–4213.9722641
31. Berger J.M. , GamblinS.J., HarrisonS.C., WangJ.C. Structure and mechanism of DNA topoisomerase II. Nature. 1996; 379 :225–232.8538787
32. Berger J.M. , FassD., WangJ.C., HarrisonS.C. Structural similarities between topoisomerases that cleave one or both DNA strands. Proc. Natl. Acad. Sci. U.S.A. 1998; 95 :7876–7881.9653108
33. Dong K.C. , BergerJ.M. Structural basis for gate-DNA recognition and bending by type IIA topoisomerases. Nature. 2007; 450 :1201–1205.18097402
34. McClendon A.K. , GentryA.C., DickeyJ.S., BrinchM., BendsenS., AndersenA.H., OsheroffN. Bimodal recognition of DNA geometry by human topoisomerase II alpha: preferential relaxation of positively supercoiled DNA requires elements in the C-terminal domain. Biochemistry. 2008; 47 :13169–13178.19053267
35. Delint-Ramirez I. , KonadaL., HeadyL., RuedaR., JacomeA.S.V., MarlinE., MarchioniC., SegevA., KritskiyO., YamakawaS.et al . Calcineurin dephosphorylates topoisomerase IIbeta and regulates the formation of neuronal-activity-induced DNA breaks. Mol. Cell. 2022; 82 :3794–3809.36206766
36. Wu C.C. , LiT.K., FarhL., LinL.Y., LinT.S., YuY.J., YenT.J., ChiangC.W., ChanN.L. Structural basis of type II topoisomerase inhibition by the anticancer drug etoposide. Science. 2011; 333 :459–462.21778401
37. Bax B.D. , ChanP.F., EgglestonD.S., FosberryA., GentryD.R., GorrecF., GiordanoI., HannM.M., HennessyA., HibbsM.et al . Type IIA topoisomerase inhibition by a new class of antibacterial agents. Nature. 2010; 466 :935–940.20686482
38. Vanden Broeck A. , LotzC., OrtizJ., LamourV. Cryo-EM structure of the complete E. coli DNA gyrase nucleoprotein complex. Nat. Commun. 2019; 10 :4935.31666516
39. Laponogov I. , VeselkovD.A., CrevelI.M., PanX.S., FisherL.M., SandersonM.R. Structure of an ‘open’ clamp type II topoisomerase-DNA complex provides a mechanism for DNA capture and transport. Nucleic Acids Res. 2013; 41 :9911–9923.23965305
40. Wohlkonig A. , ChanP.F., FosberryA.P., HomesP., HuangJ., KranzM., LeydonV.R., MilesT.J., PearsonN.D., PereraR.L.et al . Structural basis of quinolone inhibition of type IIA topoisomerases and target-mediated resistance. Nat. Struct. Mol. Biol. 2010; 17 :1152–1153.20802486
41. Laponogov I. , PanX.S., VeselkovD.A., McAuleyK.E., FisherL.M., SandersonM.R. Structural basis of gate-DNA breakage and resealing by type II topoisomerases. PLoS One. 2010; 5 :e11338.20596531
42. Laponogov I. , SohiM.K., VeselkovD.A., PanX.S., SawhneyR., ThompsonA.W., McAuleyK.E., FisherL.M., SandersonM.R. Structural insight into the quinolone-DNA cleavage complex of type IIA topoisomerases. Nat. Struct. Mol. Biol. 2009; 16 :667–669.19448616
43. Wang Y.R. , ChenS.F., WuC.C., LiaoY.W., LinT.S., LiuK.T., ChenY.S., LiT.K., ChienT.C., ChanN.L. Producing irreversible topoisomerase II-mediated DNA breaks by site-specific Pt(II)-methionine coordination chemistry. Nucleic Acids Res. 2017; 45 :10861–10871.28977631
44. Germe T. , VorosJ., JeannotF., TaillierT., StavengerR.A., BacqueE., MaxwellA., BaxB.D. A new class of antibacterials, the imidazopyrazinones, reveal structural transitions involved in DNA gyrase poisoning and mechanisms of resistance. Nucleic Acids Res. 2018; 46 :4114–4128.29538767
45. Wendorff T.J. , SchmidtB.H., HeslopP., AustinC.A., BergerJ.M. The structure of DNA-bound human topoisomerase II alpha: conformational mechanisms for coordinating inter-subunit interactions with DNA cleavage. J. Mol. Biol. 2012; 424 :109–124.22841979
46. Bax B.D. , MurshudovG., MaxwellA., GermeT. DNA topoisomerase inhibitors: trapping a DNA-cleaving machine in motion. J. Mol. Biol. 2019; 431 :3427–3449.31301408
47. Deweese J.E. , OsheroffN. The use of divalent metal ions by type II topoisomerases. Metallomics. 2010; 2 :450–459.20703329
48. Chen S.F. , HuangN.L., LinJ.H., WuC.C., WangY.R., YuY.J., GilsonM.K., ChanN.L. Structural insights into the gating of DNA passage by the topoisomerase II DNA-gate. Nat. Commun. 2018; 9 :3085.30082834
49. Liu Q. , WangJ.C. Similarity in the catalysis of DNA breakage and rejoining by type IA and IIA DNA topoisomerases. Proc. Natl. Acad. Sci. U.S.A. 1999; 96 :881–886.9927662
50. West K.L. , MeczesE.L., ThornR., TurnbullR.M., MarshallR., AustinC.A. Mutagenesis of E477 or K505 in the B' domain of human topoisomerase II beta increases the requirement for magnesium ions during strand passage. Biochemistry. 2000; 39 :1223–1233.10684600
51. Hardin A.H. , SarkarS.K., SeolY., LiouG.F., OsheroffN., NeumanK.C. Direct measurement of DNA bending by type IIA topoisomerases: implications for non-equilibrium topology simplification. Nucleic Acids Res. 2011; 39 :5729–5743.21421557
52. Wigley D.B. , DaviesG.J., DodsonE.J., MaxwellA., DodsonG. Crystal structure of an N-terminal fragment of the DNA gyrase B protein. Nature. 1991; 351 :624–629.1646964
53. Classen S. , OllandS., BergerJ.M. Structure of the topoisomerase II ATPase region and its mechanism of inhibition by the chemotherapeutic agent ICRF-187. Proc. Natl. Acad. Sci. U.S.A. 2003; 100 :10629–10634.12963818
54. Ling E.M. , BasleA., CowellI.G., van den BergB., BlowerT.R., AustinC.A. A comprehensive structural analysis of the ATPase domain of human DNA topoisomerase II beta bound to AMPPNP, ADP, and the bisdioxopiperazine, ICRF193. Structure. 2022; 30 :1129–1145.35660158
55. Wei H. , RuthenburgA.J., BechisS.K., VerdineG.L. Nucleotide-dependent domain movement in the ATPase domain of a human type IIA DNA topoisomerase. J. Biol. Chem. 2005; 280 :37041–37047.16100112
56. Brino L. , UrzhumtsevA., MousliM., BronnerC., MitschlerA., OudetP., MorasD. Dimerization of Escherichia coli DNA-gyrase B provides a structural mechanism for activating the ATPase catalytic center. J. Biol. Chem. 2000; 275 :9468–9475.10734094
57. Smiley R.D. , CollinsT.R., HammesG.G., HsiehT.S. Single-molecule measurements of the opening and closing of the DNA gate by eukaryotic topoisomerase II. Proc. Natl. Acad. Sci. U.S.A. 2007; 104 :4840–4845.17360343
58. Fass D. , BogdenC.E., BergerJ.M. Quaternary changes in topoisomerase II may direct orthogonal movement of two DNA strands. Nat. Struct. Biol. 1999; 6 :322–326.10201398
59. Schoeffler A.J. , MayA.P., BergerJ.M. A domain insertion in Escherichia coli GyrB adopts a novel fold that plays a critical role in gyrase function. Nucleic Acids Res. 2010; 38 :7830–7844.20675723
60. Austin C.A. , MarshK.L., WassermanR.A., WillmoreE., SayerP.J., WangJ.C., FisherL.M. Expression, domain structure, and enzymatic properties of an active recombinant human DNA topoisomerase II beta. J. Biol. Chem. 1995; 270 :15739–15746.7797575
61. Nitiss J.L. , LiuY.X., HarburyP., JannatipourM., WassermanR., WangJ.C. Amsacrine and etoposide hypersensitivity of yeast cells overexpressing DNA topoisomerase II. Cancer Res. 1992; 52 :4467–4472.1322791
62. Hsiung Y. , JannatipourM., RoseA., McMahonJ., DuncanD., NitissJ.L. Functional expression of human topoisomerase II alpha in yeast: mutations at amino acids 450 or 803 of topoisomerase II alpha result in enzymes that can confer resistance to anti-topoisomerase II agents. Cancer Res. 1996; 56 :91–99.8548781
63. Otwinowski Z. , MinorW. Processing of X-ray diffraction data collected in oscillation mode. Methods Enzymol. 1997; 276 :307–326.27754618
64. Liebschner D. , AfonineP.V., BakerM.L., BunkocziG., ChenV.B., CrollT.I., HintzeB., HungL.W., JainS., McCoyA.J.et al . Macromolecular structure determination using X-rays, neutrons and electrons: recent developments in Phenix. Acta Crystallogr D Struct. Biol. 2019; 75 :861–877.31588918
65. Emsley P. , LohkampB., ScottW.G., CowtanK. Features and development of Coot. Acta. Crystallogr. D Biol. Crystallogr. 2010; 66 :486–501.20383002
66. Blower T.R. , BandakA., LeeA.S.Y., AustinC.A., NitissJ.L., BergerJ.M. A complex suite of loci and elements in eukaryotic type II topoisomerases determine selective sensitivity to distinct poisoning agents. Nucleic Acids Res. 2019; 47 :8163–8179.31287876
67. Vayssieres M. , MarechalN., YunL., Lopez DuranB., MurugasamyN.K., FoggJ.M., ZechiedrichL., NadalM., LamourV. Structural basis of DNA crossover capture by Escherichia coli DNA gyrase. Science. 2024; 384 :227–232.38603484
68. Broderick L. , YostS., LiD., McGeoughM.D., BooshehriL.M., GuaderramaM., BrydgesS.D., KucharovaK., PatelN.C., HarrM.et al . Mutations in topoisomerase IIbeta result in a B cell immunodeficiency. Nat. Commun. 2019; 10 :3644.31409799
69. Broderick L. , ClayG.M., BlumR.H., LiuY., McVicarR., PapesF., BooshehriL.M., CowellI.G., AustinC.A., PutnamC.D.et al . Disease-associated mutations in topoisomerase IIbeta result in defective NK cells. J. Allergy Clin. Immunol. 2022; 149 :2171–2176.35063500
70. Cepni E. , BorkluE., AvciS., KalayciT., EraslanS., KayseriliH. Revisiting TOP2B-related phenotypes: three new cases and literature review. Clin. Genet. 2023; 104 :251–258.37068767
71. Lee S. , JungS.R., HeoK., BylJ.A., DeweeseJ.E., OsheroffN., HohngS. DNA cleavage and opening reactions of human topoisomerase IIalpha are regulated via Mg2+-mediated dynamic bending of gate-DNA. Proc. Natl. Acad. Sci. U.S.A. 2012; 109 :2925–2930.22323612
72. Hol W.G. The role of the alpha-helix dipole in protein function and structure. Prog. Biophys. Mol. Biol. 1985; 45 :149–195.3892583
73. Hol W.G. , van DuijnenP.T., BerendsenH.J. The alpha-helix dipole and the properties of proteins. Nature. 1978; 273 :443–446.661956
74. Forood B. , FelicianoE.J., NambiarK.P. Stabilization of alpha-helical structures in short peptides via end capping. Proc. Natl. Acad. Sci. U.S.A. 1993; 90 :838–842.8430094
75. Ryu H. , FurutaM., KirkpatrickD., GygiS.P., AzumaY. PIASy-dependent SUMOylation regulates DNA topoisomerase IIalpha activity. J. Cell Biol. 2010; 191 :783–794.21079245
76. Funatsuki K. , TanakaR., InagakiS., KonnoH., KatohK., NakamuraH. acrB mutation located at carboxyl-terminal region of gyrase B subunit reduces DNA binding of DNA gyrase. J. Biol. Chem. 1997; 272 :13302–13308.9148951
77. Fu G. , WuJ., LiuW., ZhuD., HuY., DengJ., ZhangX.E., BiL., WangD.C. Crystal structure of DNA gyrase B' domain sheds lights on the mechanism for T-segment navigation. Nucleic Acids Res. 2009; 37 :5908–5916.19596812
78. Pommier Y. , MarchandC. Interfacial inhibitors: targeting macromolecular complexes. Nat. Rev. Drug Discov. 2011; 11 :25–36.22173432
79. Ben Chorin A. , MasratiG., KesselA., NarunskyA., SprinzakJ., LahavS., AshkenazyH., Ben-TalN. ConSurf-DB: an accessible repository for the evolutionary conservation patterns of the majority of PDB proteins. Protein Sci. 2020; 29 :258–267.31702846
