1. Biochemistry and Chemical Biology
  2. Structural Biology and Molecular Biophysics
Download icon

Topoisomerase VI senses and exploits both DNA crossings and bends to facilitate strand passage

  1. Timothy J Wendorff
  2. James M Berger  Is a corresponding author
  1. University of California, Berkeley, United States
  2. Johns Hopkins University School of Medicine, United States
Research Article
  • Cited 0
  • Views 1,174
  • Annotations
Cite this article as: eLife 2018;7:e31724 doi: 10.7554/eLife.31724

Abstract

Type II topoisomerases manage DNA supercoiling and aid chromosome segregation using a complex, ATP-dependent duplex strand passage mechanism. Type IIB topoisomerases and their homologs support both archaeal/plant viability and meiotic recombination. Topo VI, a prototypical type IIB topoisomerase, comprises two Top6A and two Top6B protomers; how these subunits cooperate to engage two DNA segments and link ATP turnover to DNA transport is poorly understood. Using multiple biochemical approaches, we show that Top6B, which harbors the ATPase activity of topo VI, recognizes and exploits the DNA crossings present in supercoiled DNA to stimulate subunit dimerization by ATP. Top6B self-association in turn induces extensive DNA bending, which is needed to support duplex cleavage by Top6A. Our observations explain how topo VI tightly coordinates DNA crossover recognition and ATP binding with strand scission, providing useful insights into the operation of type IIB topoisomerases and related meiotic recombination and GHKL ATPase machineries.

https://doi.org/10.7554/eLife.31724.001

eLife digest

Each human cell contains genetic information stored on approximately two meters of DNA. Like holiday lights in a storage box, packing so much DNA into such a small space leads to its entanglement. This snarled DNA prevents the cell from properly accessing and copying its genes.

Type II topoisomerases are a group of enzymes that remove DNA tangles. They attach to one segment of a DNA tangle, cut it in half, remove the knot, and then repair the broken DNA strand. The process requires the proteins to ‘burn’ chemical energy. If topoisomerases make mistakes when they cut and reseal DNA, they could damage genetic information and harm cells. It is still unclear how these proteins recognize DNA tangles and use energy to remove knots instead of adding them.

Here, Wendorff and Berger use biochemical approaches to look into topo VI, a type II topoisomerase found in plants and certain single-celled organisms. When DNA is tangled, it forms sharp bends and crossings. Their experiments reveal that topo VI has certain ‘sensors’ that detect where DNA bends, and others that recognize the crossings. Only when both features are present does the enzyme start working and using energy. These sensors act as fail-safes to ensure that topo VI only breaks DNA when it encounters a proper knot, and is not ‘set loose’ on untangled DNA.

Future work will look at topo VI at an atom-by-atom level to reveal how exactly the enzymes ‘see’ DNA bends and crossings, and how interactions with the correct type of DNA triggers energy use and DNA untangling. Knowing more about topo VI can help researchers to understand how human and bacterial topoisomerases work. These results could also be generalized to other enzymes, for example those that help the genetic processes at play when sperm and egg cells form.

https://doi.org/10.7554/eLife.31724.002

Introduction

The appropriate control of transcription, DNA replication, and chromosome segregation are essential to cell proliferation. These three processes are antagonized, however, by the double helical structure of DNA, which supercoils in response to helicase and polymerase activity and which promotes chromosome interlinkage during replicative synthesis (reviewed in [Vos et al., 2011; Wang, 2002]). In all cells, enzymes known as topoisomerases are used to overcome the natural topological impediments arising from these physical transactions that impinge on DNA. Many different topoisomerase families exist to relieve super-helical tension and DNA entanglements, all of which transiently form either single strand or double strand breaks to manipulate DNA topology (reviewed in [Chen et al., 2013; Forterre and Gadelle, 2009; Schoeffler and Berger, 2008]).

Type II topoisomerases introduce transient double strand breaks into DNA, and play a key role in unlinking catenated DNA molecules (Holm et al., 1985; Spell and Holm, 1994; Zechiedrich and Cozzarelli, 1995). The type IIA subfamily of topoisomerases, which are principally used in bacteria and eukaryotes, utilize a so-called ‘two-gate’ mechanism (Roca et al., 1996; Roca and Wang, 1994; Roca and Wang, 1992), in which one DNA duplex (termed the transport- or ‘T’-segment) is captured by one half of the enzyme, actively passed through a second, protein-bound DNA duplex (the gate- or ‘G’-segment), and expelled through the other end of the enzyme. T-segment capture is regulated by the ATP-dependent closure of one subunit dimer interface (Roca and Wang, 1992; Wigley et al., 1991), referred to as the ‘ATP-gate’, while a pair of catalytic tyrosine residues responsible for G-segment cleavage and opening reside in a second, separable subunit-subunit contact point termed the ‘DNA-gate’ (Berger et al., 1996; Morais Cabral et al., 1997; Morrison and Cozzarelli, 1979; Tse et al., 1980). In most instances, repeated cycles of ATP binding and hydrolysis allow for the processive removal of multiple DNA crossings. The use of ATP by type II topoisomerases in general has been proposed to serve as a mechanism for preventing the inappropriate formation of potentially cytotoxic DNA breaks (Bates et al., 2011); however, the molecular basis for the coupling between ATP turnover and DNA cleavage has remained enigmatic for the superfamily as a whole.

The type IIB topoisomerases, which are exemplified by DNA topoisomerase VI (topo VI) (Bergerat et al., 1997; Bergerat et al., 1994; Forterre et al., 2007), share evolutionarily conserved catalytic elements with their type IIA counterparts but are structurally distinct (Corbett and Berger, 2003; Nichols et al., 1999). Topo VI comprises an A2B2 heterotetramer formed by two Top6A and two Top6B subunits: Top6A forms a ‘U’-shaped dimer that serves as the DNA-gate for G-segment cleavage and opening (Bergerat et al., 1997; Nichols et al., 1999), while Top6B constitutes the ATP-gate and dimerizes in response to nucleotide binding (Corbett and Berger, 2003). Topo VI is thought to serve as the primary topoisomerase for DNA decatenation and supercoil relaxation in archaea and is required for endoreduplication and cell growth in plants (Bergerat et al., 1997; Bergerat et al., 1994; Hartung et al., 2002; Sugimoto-Shirasu et al., 2002; Yin et al., 2002). Topo VI is also found sporadically throughout the bacterial domain, and a single chain variant, topo VIII, is found in certain plasmid-based mobile elements as well (Forterre et al., 2007; Gadelle et al., 2014). Interestingly, the type IIB topoisomerase scaffold has been co-opted to serve as the machinery responsible for introducing double-strand DNA breaks to initiate meiotic recombination in eukaryotes (Bergerat et al., 1997; Keeney et al., 1997; Robert et al., 2016; Vrielynck et al., 2016). How topo VI and its cousins engage DNA segments has yet to be determined.

The ATPase region is generally well-preserved between type IIA and type IIB topoisomerases, with the exception of an additional helix-two-turn-helix (H2TH) domain of unknown function found in topo VI and topo VIII (Bergerat et al., 1997; Corbett and Berger, 2003; Gadelle et al., 2014; Wigley et al., 1991). By contrast, the catalytic domains that comprise the DNA breakage-reunion region of type IIA and IIB enzymes have been extensively shuffled. One consequence of this rearrangement is that Top6A lacks a third subunit-subunit interface present in the type IIA enzymes, the ‘C-gate’ dimerization domain, which is thought to help mitigate the risk of aberrant double-strand break formation (Bates et al., 2011; Berger et al., 1996; Nichols et al., 1999; Roca, 2004). To compensate for the loss of this element, type IIB topoisomerases appear to have evolved a stringent mechanism for controlling strand scission by Top6A that represses transesterase activity until ATP productively binds to Top6B (Buhler et al., 1998; Buhler et al., 2001). How Top6B activates Top6A is unknown; however, given that Spo11, a paralog of Top6A used in meiotic recombination, is also thought to require activation for DNA cleavage, aspects of this control mechanism may be broadly conserved (Lam and Keeney, 2014).

To better understand how type IIB topoisomerases coordinate DNA cleavage, we performed a comprehensive biochemical investigation of Methanosarcina mazei topo VI, a model mesophilic type IIB topoisomerase. We find that topo VI discriminates between linear and supercoiled DNA using an extensive and unanticipated DNA binding interface that specifically recognizes DNA crossings. Both gate closure and ATP hydrolysis by Top6B as well as transesterase activity by Top6A require engagement along this entire interface. Site-directed mutagenesis studies show that three conserved, positively charged regions on Top6B sense both the DNA bends and crossings present in supercoiled substrates and further serve to couple the binding of DNA crossings to B-subunit dimerization, nucleotide turnover, and DNA strand scission. Our results explain why type IIB topoisomerases absolutely depend upon the ATPase activity of the B-subunit to generate double strand breaks. These observations in turn reinforce the functional importance for DNA bending and potential T-segment-sensing elements in the related type IIA topoisomerases, and also provide insights as to how recently discovered meiotic Top6B homologs might promote Spo11 mediated strand scission during meiotic recombination.

Results

Topo VI is a distributive DNA relaxase that preferentially recognizes DNA crossings

We began our investigations of type IIB topoisomerase mechanism by measuring the affinity of M. mazei topo VI for DNAs of varying length or topological status. The relative affinity of the holoenzyme for fluorescein-labeled duplex DNAs ranging from 20 bp to 70 bp in length was assessed using a fluorescence anisotropy-based approach (the predicted G-segment binding channel of a Top6A dimer is ~16–20 bp in length [Nichols et al., 1999]). The DNA sequence used for these oligomers was based on a previously determined cleavage hotspot for Sulfolobus shibatae topo VI (Buhler et al., 2001) (Figure 1—source data 1). These experiments showed that whereas a 20 bp duplex binds relatively weakly to topo VI, apparent affinity increases with length, plateauing between 40 and 70 bp (Figure 1A, Figure 1—source datas 23). As the binding isotherms did not show any sign of complex interactions (such as cooperativity) and could be fit well by a single-site binding model (Heyduk and Lee, 1990), this result provided the first clue that topo VI might have more extensive interactions with DNA than previously hypothesized.

Figure 1 with 1 supplement see all
Topo VI binds longer duplexes and preferentially engages features of supercoiled DNA.

(A) Binding of a 20, 30, 40, 60 or 70 bp fluorescein-labeled duplex (20 nM, sequences in Figure 1—source data 1) to topo VI, observed as a change in fluorescence anisotropy (ΔFA) measured in milli-anisotropy units (mA) as a function of enzyme concentration. Points and error bars correspond to the mean and standard deviation of three independent experiments. Curves represent fits to a single-site ligand depletion binding model. Apparent dissociation constants are reported in Figure 1—source data 2. (B) Fluorescence anisotropy experiment assessing the ability of supercoiled DNA and sheared salmon-sperm DNA to compete a fluorescein-labeled 70 bp duplex (20 nM duplex, 1.4 μM bp) from topo VI (100 nM). Non-labeled DNA was titrated from 0.1 μM bp to 106.5 μM bp and competition was observed as a change in fluorescence anisotropy (ΔFA) as measured in milli-anisotropy units (mA). Data are plotted as a function of the base-pair concentration (μM) of competing DNA. Points and error bars correspond to the mean and standard deviation of three independent experiments. Curves represent a fit to a competitive displacement model. Numerical data for (A–B) are reported in Figure 1—source data 3. (C–D) Test of processive supercoil relaxation by topo VI on negatively supercoiled plasmid DNA. Topo VI was pre-incubated in a 1:1.4 ratio to a 2.9 kb negatively supercoiled plasmid (6.7 ng/μL in assay). Reactions were started by addition of either (C) ATP or (D) ATP and a 6.5 kb ‘chase’ plasmid (6.7 ng/μL in assay) to compete for unbound enzyme. Samples were quenched at 0, 1, 2, 4, 6, 8, 10, 15, and 20 min. Each condition was also incubated without ATP for 20 min as a negative control. Plasmid size and topoisomer species are indicated to the left of each gel. For an example of processive supercoil relaxation by a type II topoisomerase, see Figure 1—figure supplement 1.

https://doi.org/10.7554/eLife.31724.003

To determine whether topo VI displayed any preference for the topological status of DNA, the relative binding affinities of the holoenzyme were next assessed for supercoiled plasmid vs. sheared, linear salmon-sperm DNA using a competitive binding assay. Topo VI was incubated with the fluorescein-labeled 70 bp duplex DNA and varying amounts of unlabeled, supercoiled plasmid or sheared salmon-sperm DNA. The relative affinity of topo VI for each substrate was determined by monitoring how well the competitor DNAs interfered with binding of the labeled probe. The response to the titration of supercoiled DNA or sheared salmon-sperm DNA was fit to an explicit competitive binding model (Wang, 1995) to indirectly estimate affinities for the unlabeled substrates. Based on these measurements, topo VI showed a ~60-fold preference for supercoiled DNA (KI,app = 0.6 ± 0.3 nM) compared to sheared salmon-sperm DNA (KI,app = 39.3 ± 2.6 nM) (Figure 1B).

The difference in affinity between supercoiled and linear DNA suggested that topo VI might preferentially engage supercoiled substrates by binding to DNA crossings, DNA bends, or both. To distinguish between these modes, we examined the time-dependent processivity of topo VI in relaxing negatively supercoiled DNA. For type II topoisomerases in general, processivity describes the ability of a single enzyme to remain bound to a G-segment DNA during multiple strand passage events. For topo VI, the progress of ATP-dependent supercoil relaxation was followed by native agarose-gel electrophoresis, using a slight molar excess of plasmid over enzyme to disfavor the binding of two topo VI molecules to a single DNA substrate (Figure 1C). A highly processive topoisomerase, such as Saccharomyces cerevisiae topoisomerase II (ScTop2), removes the majority of supercoils on a closed circular DNA in a single enzyme-DNA encounter (Figure 1—figure supplement 1) as evidenced by a paucity of intermediate DNA topoisomers between the supercoiled substrate and fully relaxed plasmid product. In contrast, topo VI produced a broad distribution of intermediate topoisomers that were gradually converted to the fully relaxed distribution, a behavior more consistent with low processivity.

To more thoroughly investigate supercoil processing by topo VI, we followed plasmid relaxation using two differently sized plasmids in a chase experiment. Following pre-incubation of a defined amount of topo VI with a slight molar excess of a primary 2.9 kb plasmid, a second, larger plasmid (6.5 kb) was added along with ATP to serve as a competing substrate for any dissociated enzymes (Figure 1D). In the case of a processive enzyme such as ScTop2, the competing plasmid does not alter the initial rate at which a fully relaxed topoisomer distribution of the primary plasmid is generated (Figure 1—figure supplement 1). By contrast, topo VI again displayed clearly distributive behavior, relaxing both plasmids more slowly and simultaneously. Although assay conditions can modulate whether a topoisomerase acts processively or distributively (salt concentration in particular), both timecourse experiments were run under low-salt conditions where type IIA topoisomerases are primarily processive. Collectively, these findings demonstrate that topo VI operates by a principally distributive mechanism, whereby once a DNA crossing is resolved by strand passage, the enzyme will tend to dissociate from the substrate before acting on a new crossing and/or bent DNA segment.

Topo VI actively uses DNA crossings to couple ATP hydrolysis with DNA strand passage

A defining characteristic of type II topoisomerases is the coupling of ATP turnover with efficient and rapid strand passage. In type IIA topoisomerases, DNA binding strongly stimulates ATPase activity (Lee et al., 2013; Lindsley and Wang, 1993; Liu et al., 1979; Mizuuchi et al., 1978; Osheroff et al., 1983; Sugino and Cozzarelli, 1980). However, the coupling of DNA topological state to the magnitude of the ATP hydrolysis stimulation varies between different type IIA homologs (Anderson et al., 1998; Gubaev and Klostermeier, 2011; Harkins and Lindsley, 1998; McClendon et al., 2005; Osheroff et al., 1983; Sugino and Cozzarelli, 1980; Vaughn et al., 2005). To determine whether the ATPase activity of type IIB topoisomerases is stimulated in a DNA topology-dependent manner, we examined nucleotide turnover by wild-type topo VI in the absence and presence of linear sheared salmon-sperm DNA and supercoiled plasmid DNA substrates at varying ATP concentrations using a coupled assay. Hydrolysis was also measured for an ATPase-deficient topo VI construct (Top6BE44A) to identify non-specific activity arising from contaminating ATPases (Figure 2A, Figure 2—figure supplement 1). Although topo VI likely hydrolyzes ATP cooperatively, the data conformed to apparent Michalis-Menten behavior and were fit to this model (Figure 2—source datas 12). Topo VI showed negligible basal ATPase activity, and required the addition of a DNA substrate to hydrolyze ATP. Incubation with supercoiled DNA produced the maximal observed rate of hydrolysis (and decreased the Km,app for ATP), resulting in a ~5-fold increase in catalytic efficiency (kcat,app/Km,app) over that for sheared salmon-sperm DNA. The observation that supercoiled DNA is more effective than linear substrates in activating ATP turnover indicates that topo VI not only interrogates DNA for specific topological features, but that its activity is potentiated when such features are recognized.

Figure 2 with 3 supplements see all
Top6B dimerization and ATP hydrolysis activity are stimulated by supercoiled DNA.

(A) Rate of steady-state ATP hydrolysis catalyzed by topo VI alone, topo VI incubated with 400 μM basepairs of linear sheared salmon-sperm DNA (800:1 basepair to enzyme ratio), or topo VI incubated with 400 μM basepairs of 2.9 kb supercoiled plasmid DNA (800:1 basepair to enzyme ratio) as a function of ATP concentration. Rates were determined spectroscopically using an NADH-coupled assay. Data represent hydrolysis rates after subtracting a small contribution of non-specific ATPase activity from assays performed with an ATPase-deficient topo VI construct (Figure 2—figure supplement 1). Points and bars correspond to the mean and standard error of the mean of three independent experiments. Curves represent fits to a Michealis-Menten kinetics model reported in Figure 2—source data 1. (B) The ratiometric FRET efficiency of Alexa555/Alexa647-labeled topo VIcyslite-155C (see Figure 2—figure supplement 2) was monitored over time following the addition of AMPPNP for enzyme alone, enzyme bound to supercoiled DNA, or enzyme bound to short linear (sheared salmon-sperm) DNA. Numerical data for (A–B) are reported in Figure 2—source data 2. (C–D) Example fluorescence emission spectra produced by 530 nm excitation of Alexa555/Alexa647-labeled topo VIcyslite-155C assessing the conformation of the Top6B ATPase domain in the absence of nucleotide ((C), solid lines) and 120 min following addition of AMPPNP ((D), dashed lines). Spectral emission was normalized by total emission from 545 nm to 700 nm. This behavior contrasts that of a model type IIA topoisomerase, ScTop2 (see Figure 2—figure supplement 3).

https://doi.org/10.7554/eLife.31724.008

ATP binding and hydrolysis by topo VI and many other enzymes that share its GHKL ATPase fold (e.g. type IIA topoisomerases, Hsp90, MutL, and MORC ATPases) rely on nucleotide-dependent dimerization of ATP-binding domains to elicit biological activity (Ali et al., 2006; Ban et al., 1999; Ban and Yang, 1998; Dutta and Inouye, 2000; Li et al., 2016; Shiau et al., 2006; Wigley et al., 1991). A mechanism in which supercoiled DNA binding, in particular T-segment engagement, promotes Top6B dimerization could thus explain why supercoiled DNA stimulates ATP turnover. To test this idea, we developed a Förster Resonance Energy Transfer (FRET) assay to monitor ATPase domain dimerization in the context of the topo VI holoenzyme. We first identified and mutated surface cysteines to non-reactive residues to create a fully functional ‘cys-lite’ construct of the holoenzyme. Thr155 of Top6B was then substituted with cysteine (Figure 2—figure supplement 2A). Dual labeling with donor (Alexa 555-maleimide) and acceptor (Alexa 647-maleimide) fluorophores yielded an enzyme population containing an expected labeled mixture of correctly labeled donor-acceptor enzymes (50%), and both acceptor-acceptor (25%) and donor-donor (25%) labeled enzymes (Figure 2—figure supplement 2B; labeling efficiency was determined by spectral absorption). The labeled topo VI holoenzymes were able to fully relax DNA and showed only a slight impairment (~2 fold) of overall specific activity compared to wild-type topo VI (Figure 2—figure supplement 2C).

Using the labeled enzyme, bulk FRET efficiencies in the absence and presence of either linear or supercoiled DNA were first measured by scanning the spectral emission of both donor and acceptor fluorophores under excitation at 530 nm. The conformational response of the enzyme to AMPPNP, a non-hydrolyzable ATP analog, was then assessed for the enzyme alone and in the presence of each substrate over time (Figure 2B). The addition of sheared salmon-sperm DNA and to a greater extent supercoiled DNA, led to minor but reproducible increases in FRET efficiency (Figures 2C, 0 min time-point), suggesting that DNA binding alone alters the conformation of Top6B in the holoenzyme. By comparison, the addition of AMPPNP led to larger FRET responses, and FRET efficiency increased much more rapidly with supercoiled DNA compared to linear sheared salmon-sperm DNA. AMPPNP alone produced detectable but minor FRET changes when DNA was omitted, indicating that duplex binding is needed for ATPase domain dimerization (Figure 2D, 120 min time-point). In conjunction with the ATPase data, these observations show that – unlike type IIA topoisomerases, whose ATPase regions efficiently dimerize in the absence of DNA ([Gubaev and Klostermeier, 2011; Roca and Wang, 1992] and Figure 2—figure supplement 3) – topo VI utilizes the DNA geometries presented by supercoiled substrates to help favor nucleotide-dependent conformational changes associated with strand passage.

Three conserved elements in Top6B play a role in DNA binding, the sensing of DNA geometry, and the productive coupling of ATP hydrolysis to strand passage

Based on the ability of topo VI to recognize and utilize topological features in supercoiled DNA to promote activity, we set out to identify the structural elements responsible for this coupling. Working from an assumption that topology-sensing elements might consist in part of positively charged residues on the B subunit, we mapped both amino acid conservation (derived from a multiple sequence alignment of Top6B homologs) and electrostatic surface potential onto the known structure of M. mazei Top6B using ConSurf and ABPS (Figure 3A–C and (Ashkenazy et al., 2010; Baker et al., 2001)). By comparing positively charged interfaces against sequence conservation, we identified three different regions as candidate DNA interaction sites.

Figure 3 with 2 supplements see all
Identification of potential DNA-binding elements in Top6B.

(A) Primary and tertiary structure [Protein Data Bank (PDB) ID: 2Q2E] of the M. mazei topo VI heterotetramer. Domains for one Top6A-Top6B heterodimer are colored as shown in the primary structure and the partner Top6A-Top6B heterodimer is shown in grey. Catalytic function is denoted in italics under primary structure. (B) Mapping of sequence conservation in Top6B based on a PSI-BLAST multiple sequence alignment. Conserved surface-exposed arginine and lysine residues (ConSurf score of ≥6) are shown as sticks. Coloration from cyan to magenta denotes variable to conserved. Top6A is represented in yellow. (C) Electrostatic surface representation of topo VI. A conserved basic loop in the T-segment storage cavity, and a conserved basic interface stretching from the WKxY motif and C-terminal stalk of Top6B to the Helix-2-turn-helix (H2TH) domain are labeled. (D) View of the KGRR basic loop motif. (E) View of the C-terminal stalk/WKxY interface. (F) View of the H2TH DNA-binding interface, rotated 90° towards the point of view as compared to A–C. See Figure 3—figure supplement 1 for a more detailed rationale for the functional importance of this interface. In (D–F), residues mutated to alanine or to glutamate for functional studies are labeled. Mutations to these interfaces produced well-behaved functional mutants (see Figure 3—figure supplement 2).

https://doi.org/10.7554/eLife.31724.014

The first prospective locus consisted of a small loop of basic residues (KGRR186-189) (Figure 3D) within the predicted T-segment storage cavity of topo VI. A second feature comprised a trio of conserved basic residues (R457, K399 and K401) that are found within two spatially adjacent structural elements (Figure 3E): the C-terminal, α-helical stalk of Top6B (which connects the so-called ‘transducer’ domain of this subunit to Top6A), and a loop in the transducer domain containing the so-called ‘WKxY motif,’ which is conserved in both Top6B and many meiotic Top6B-like proteins (Robert et al., 2016). The third area of note, the H2TH domain, is embedded between the topo VI GHKL and transducer regions. The function of the topo VI H2TH domain has not been established, but this type of fold serves as a general nucleic-acid-binding element in a diverse number of proteins, including FpG/Nei DNA glycosylases, s13 ribosomal proteins, and sIHF type nucleoid-associated proteins (Brodersen et al., 2002; Sugahara et al., 2000; Swiercz et al., 2013; Zharkov et al., 2002). Comparison of nucleic-acid-bound H2TH domain structures with Top6B (Figure 3—figure supplement 1) highlighted R263, K268 and K308 as candidate residues that might interact with DNA (Figure 3F).

Having identified three potential sites for supercoil sensing on the surface of Top6B, six constructs were generated to assess the functional attributes of each region. Selected constructs included triple-neutral and triple-acidic mutations to the basic storage-cavity loop (KGRR→AGAA and EGEE, referred to as KGRRAAA and KGRREEE), the C-terminal stalk (Stalk/WKxYAAA and Stalk/WKxYEEE), and the H2TH domain (H2THAAA and H2THEEE). All six mutant topo VI holoenzymes were soluble upon expression, purified to homogeneity (as judged by SDS-PAGE), and appeared well-behaved based on gel-filtration chromatography profiles as compared to the wild-type enzyme (Figure 3—figure supplement 2).

To assess overall activity, we next looked at the supercoil relaxation activity of the mutant enzymes compared to wild-type topo VI as a function of enzyme concentration (Figure 4A). Both sets of KGRR and Stalk/WKxY mutants (neutral and acidic) proved completely unable to relax supercoiled substrate. By contrast, both sets of mutations to the H2TH region led to enzymes that were able to relax supercoiled DNA, but with ~20–30 fold lower efficiency than native topo VI. The activity profiles seen in enzyme titration assays were corroborated by timecourse assays at a fixed enzyme concentration (Figure 4—figure supplement 1). In some of these experiments, the open-circle (nicked) plasmid species increased over time; however, this increase was independent of both nucleotide and topo VI, and thus does not reflect an elevated nicking activity of the mutants. Collectively, these findings show that the KGRR loop and the Stalk/WKxY region are essential components for topo VI function, but that the H2TH domain, while important, is not strictly required for strand passage.

Figure 4 with 2 supplements see all
Effect of neutralization and charge reversal mutations to the KGRR loop, Stalk/WKxY region, or H2TH DNA-binding interface on supercoil relaxation activity and ATP hydrolysis by topo VI.

(A) Activity of mutant topo VI constructs for relaxing supercoiled DNA compared to wild type as a function of enzyme concentration. For the enzyme titrations (0.3–20 nM in two-fold steps), each assay proceeded for 30 min prior to quenching with EDTA and SDS and contained 3.5 nM plasmid (10.2 μM bp DNA). Similar behavior for each mutant may be observed by timecourse (Figure 4—figure supplement 1). The placement and nature of mutations in each construct are depicted in the cartoons above each titration (‘•••' - AAA; ‘- - -' – EEE). (B) The rate of steady-state ATP hydrolysis above basal levels (Figure 4—figure supplement 2) catalyzed by wild-type topo VI compared to mutant topo VI constructs, plotted as a function of the basepair concentration (µM) of sheared salmon-sperm DNA (pink), or a 2.9 kb supercoiled plasmid DNA (orange). ATP was held at 2 mM, and rates were determined spectroscopically using an NADH-coupled assay. Points and bars correspond to the mean and standard deviation of three independent experiments. Curves represent a fit to a Michealis-Menten type kinetics model reported in Figure 4—source data 1. Numerical data are reported in Figure 4—source data 2.

https://doi.org/10.7554/eLife.31724.017

To further investigate the role of each DNA-binding interface in the topo VI reaction cycle, the stimulatory effect of sheared salmon-sperm DNA and supercoiled DNA upon ATP hydrolysis activity of the six mutants was compared to wild-type enzyme. ATP hydrolysis rates were again measured using a coupled assay; however, ATP was held at 2 mM for these experiments, while the concentration of DNA substrate was varied to characterize the stimulatory effects of each substrate on each enzyme (Figure 4B, Figure 4—source datas 12). No DNA-stimulated ATP turnover was observed for either of the Stalk/WKxY mutants. Interestingly, the H2THAAA and H2THEEE mutants, which exhibited large defects in strand passage, showed similar levels of ATP hydrolysis stimulation by both DNA substrates as compared to wildtype topo VI. Moreover, whereas no additional ATP turnover was observed for the KGRRAAA and KGRREEE mutants on sheared salmon-sperm DNA, both variants showed an increased maximal rate of DNA-stimulated ATP hydrolysis compared to the wild-type enzyme on supercoiled DNA (albeit with a more weakly coupled response to DNA concentration than wildtype topo VI or the H2TH mutants as judged by Kstim,DNA). All six mutants exhibited basal hydrolysis rates similar to both wild-type topo VI and the ATPase-deficient Top6ABE44A construct (Figure 4—figure supplement 2), indicating that the DNA-stimulated responses of each topo VI mutant are directly attributable to the introduced alterations. Collectively, these data indicate that the abrogation of strand passage activity by the KGRR loop mutants stems in part from a loss of an essential DNA-sensing motif required to carry out strand passage. However, unlike the Stalk/WKxY mutants, the KGRR loop mutants retain some feature which allows supercoiled, but not short linear DNAs, to promote ATP hydrolysis. Mutations to the H2TH domain additionally appear to largely decouple strand passage from ATP hydrolysis, yet do not appreciably alter the DNA dependence of ATPase activity. This result implies a role for the H2TH domain in facilitating A- and B-subunit coordination to minimize futile cycling.

Since all three interfaces identified affect strand passage activity and its coupling to ATP turnover, we next tested whether the observed differences result directly from weakened binding to duplex DNA. Using fluorescence anisotropy, the affinity of each mutant was assessed for a range of duplex lengths (30, 40, 60, and 70 bp) found to exhibit moderate-to-tight binding to wild-type topo VI (Figure 5A and Figure 5—source datas 1 and 3). As with native topo VI, a single-site binding model adequately described the DNA-binding isotherms for the mutant panel; the one exception was the data for the H2THEEE mutant, which fit better to a cooperative model. This result suggests that charge reversal in the H2TH region may alter how longer duplexes are bound by the enzyme—although the direct binding data suggest that H2THEEE binds longer DNAs better than H2THAAA, both mutants display similar affinities for a 60 and 70 bp duplex in competitive binding experiments (Figure 5—figure supplement 1), indicating that differences in the fluorophore environment may underlie the higher Kd,app values seen in the direct binding study. Both Stalk/WKxY mutants were compromised for DNA binding overall (as judged by the maximum observed changes in anisotropy), with the magnitude of the binding defects proving more severe for the acidic substitutions. This finding highlights the Stalk/WKxY region of Top6B as an important DNA-binding interface, a finding that helps explain both why the binding affinity of topo VI is higher for DNAs whose length exceeds what is necessary to bind a Top6A dimer alone and why mutations in this region lead to defects in both strand passage and ATP hydrolysis. By comparison, the KGRR loop and the H2TH domain mutants showed either no change or only a moderate decrease (for the 60 and 70 bp duplexes) in DNA affinity compared to wildtype topo VI, suggesting that these regions potentially contribute a more peripheral or secondary site of DNA binding.

Figure 5 with 1 supplement see all
Effect of neutralization and charge reversal mutations to Top6B on DNA binding affinity and preferential engagement of supercoiled DNA.

(A) Binding of a 30, 40, 60, or 70 bp fluorescein-labeled duplex (20 nM) to topo VI mutant constructs. Binding was observed as a change in fluorescence anisotropy (ΔFA) and measured in milli-anisotropy units (mA) as a function of enzyme concentration. Points and error bars correspond to the mean and standard deviation of three independent experiments. For the H2THEEE mutant, curves represent fits to a Hill-type cooperative binding model. All other curves represent fits to a single site ligand depletion binding model. Binding isotherms for the wildtype enzyme are reproduced from Figure 1A for reference. Apparent dissociation constants are reported in Figure 5—source data 1. (B) Binding assay assessing the ability of supercoiled DNA and sheared salmon-sperm DNA to compete a fluorescein-labeled 70 bp duplex (20 nM duplex, 0.14 μM bp) from 100 nM H2THAAA, H2THEEE, KGRRAAA or KGRREEE, topo VI enzyme. Non-labeled DNA was titrated from 0.1 μM bp to 106.5 μM bp with competition observed as a change in fluorescence anisotropy (ΔFA) measured in milli-anisotropy units (mA). Data are plotted as a function of the basepair concentration (µM) of competitor DNA. Points and error bars correspond to the mean and standard deviation of three independent experiments. Curves represent a fit to an explicit competitive displacement model (Figure 5—source data 2). Dashed curves corresponding to the competitive binding data for wildtype enzyme (Figure 1B) are shown for reference. Numerical data are reported in Figure 5—source data 3.

https://doi.org/10.7554/eLife.31724.022

Because the KGRR loop and H2TH domain mutants minimally impacted affinity for short duplex DNAs as compared to the Stalk/WKxY mutants, we wondered whether these motifs might instead contribute to the preferential binding of topo VI seen for supercoiled DNA (Figure 1B). To this end, the relative affinities of supercoiled plasmid and linear, sheared salmon-sperm DNA were assessed for both sets of KGRR and H2TH mutants, using the fluorescence anisotropy-based competition assay described earlier (Figure 5B, and Figure 5—source datas 23). The H2THAAA substitution minimally affected supercoiled DNA binding, whereas the H2THEEE and both KGRR substitutions resulted in a ~10–20 fold decrease of the overall affinity of topo VI for supercoiled DNA, with KGRREEE showing a greater defect than KGRRAAA. Both KGRR substitutions adversely impacted the binding of random linear DNA compared to wild type as well, a result concordant with this mutant’s negligible ATPase activity on sheared salmon-sperm DNA and which further suggests that this set of substitutions may ablate a secondary DNA-binding site on the holoenzyme. Together, these data indicate that both the KGRR loop and H2TH domain contribute to the preferential binding of topo VI to supercoiled substrates as compared to sheared salmon-sperm DNA, but that neither is solely responsible for this discrimination.

The KGRR loop acts as a DNA crossing sensor to regulate Top6B dimerization

Rather than contributing to overall DNA affinity, the biochemical and biophysical activities of our topo VI mutants implicate the KGRR loop and H2TH domain in recognizing supercoiled DNA and in coupling ATP hydrolysis to strand passage. Although these two motifs might recognize either the DNA crossings or bends present in plectonemic substrates, we hypothesized that the KGRR element in particular might sense T-segment occupancy directly due to its physical location in the holoenzyme (Figure 3). To address this question, we designed a fluorescently-labeled, 20 bp by 16 bp Holliday junction substrate that can form a stacked-X structure (Duckett et al., 1988; Ortiz-Lombardía et al., 1999) as a mimic of a prospective duplex DNA crossing (Figure 6A–B). Using fluorescence anisotropy, wildtype topo VI was found to bind this substrate nearly 4-fold more tightly than a single 20 bp DNA duplex (Figure 6C, Figure 6—source datas 12). We next asked whether mutations to the KGRR loop or the H2TH domain interfered with binding to the stacked-junction substrate. Whereas both H2TH mutants showed similar increases in affinity for the stacked-junction DNA as seen with native topo VI, the KGRRAAA mutant showed a clear decrease in affinity for this substrate compared to a 20 bp duplex (and little to no change in affinity for a 20 bp duplex alone, Figure 6C, Figure 6—source datas 12). The KGRREEE mutant displayed an even more pronounced defect in junction binding. Collectively, this response implicates the KGRR loop in the binding of DNA crossings by topo VI, potentially as a T-segment-sensing element.

Figure 6 with 1 supplement see all
Effect of neutralization and charge reversal mutations to the KGRR loop or H2TH domain on DNA crossing affinity and Top6B dimerization.

(A) A four-way junction folds into a stacked-X structure in the presence of divalent cations (PDB ID 1DCW) (Eichman et al., 2000). (B) Modeling of a prospective G-segment and T-segment DNA into a previously published structure of M. mazei topo VI (PDB ID 2Q2E)(Corbett et al., 2007). Domains are colored as in Figure 3A. The juxtaposition of the two DNAs in this intermediate closely mimic the stacked-X junction structure in Figure 6A. (C) Binding of a 20 bp fluorescein-labeled duplex (top) or 20 bp by 16 bp fluorescein-labeled stacked junction substrate (bottom, both 20 nM) to topo VI or mutant constructs. Binding was observed as a change in fluorescence anisotropy (ΔFA) and measured in milli-anisotropy units (mA) as a function of enzyme concentration. Points and bars correspond to the mean and error of three independent experiments. Curves represent fits to a single site ligand depletion binding model. In the plot of the enzyme-stacked junction binding isotherms, the fit of wildtype topo VI binding to the 20 bp duplex is displayed (---) for reference. Apparent dissociation constants are reported in Figure 6—source data 1. (D) Change in ratiometric FRET efficiency for the indicated Alexa555/647-labeled topo VI constructs incubated with supercoiled DNA was monitored over time following the addition of AMPPNP. As further detailed in Figure 6—figure supplement 1, incubation with supercoiled DNA alone increases the FRET efficiency for each construct. Numerical data are reported in Figure 6—source data 2.

https://doi.org/10.7554/eLife.31724.027

We next considered whether the binding of DNA crossings facilitated by the KGRR loop might affect how supercoiled DNA promotes the ATP-dependent dimerization of Top6B (as observed for native topo VI [Figure 2B]), or whether this activity might instead arise from an H2TH domain interaction with supercoiled DNA. To address this question, we added the KGRRAAA and H2THAAA mutations into the topo VI construct used to monitor the conformational status of the ATPase domain by FRET. Following purification and labeling, we measured the emission spectra of both mutants alone and bound to supercoiled DNA. Similar to the wild-type construct, both the KGRRAAA and H2THAAA mutants showed increased FRET efficiencies in the presence of supercoiled substrate, independent of nucleotide (Figure 6—figure supplement 1). Interestingly, the ‘enzyme alone’ spectra suggest that each mutant alters the resting conformational status of the Top6B dimer compared to wild type, with the H2THAAA mutant taking on a more open state, and the KGRRAAA mutant taking on a more closed state. Although the H2THAAA mutant displayed a lower FRET signal than wild type, both in the presence of supercoiled DNA alone and with supercoiled DNA and nucleotide, the addition of AMPPNP produced a rapid FRET increase in the H2THAAA mutant similar to that of native topo VI, indicating that the ATPase region of this mutant responds to supercoiled DNA and nucleotide in a wildtype-like manner. By contrast, the KGRRAAA mutant initially manifested a higher FRET signal than either wildtype topo VI or the H2THAAA mutant in the presence of supercoiled DNA alone; however, the addition of AMPPNP failed to elicit any further increase in FRET (Figure 6D). Given that ATP binding and hydrolysis rely on Top6B dimerization, and that the maximum observed ATPase rate of the KGRRAAA construct is actually greater than wild-type topo VI in the presence of supercoiled DNA (Figure 4B), the high initial FRET signal for this mutant suggests that its Top6B subunits can adopt a ‘pre-dimerized’ ATPase competent state in the presence of supercoiled DNA alone. As a consequence, the rapid ATP turnover by the KGRR mutants likely arises from the decoupling of ATP hydrolysis and product release from a slow conformational change necessary for strand passage (i.e. those that drive G-segment opening and T-segment release). In this view, the KGRR loop would serve not only as a sensor of DNA crossings, but also as an element that delays ATP turnover until T-segment binding or strand passage has occurred.

The H2TH interface engages an extended G-segment to couple nucleotide-dependent Top6B dimerization with DNA cleavage

Since the H2TH domain does not appear to participate in T-segment sensing (Figure 6D), yet is important for the strand passage activity of topo VI (Figure 4A), we considered whether this element might instead interact with the G-segment. The H2TH domains reside far from the site of G-segment cleavage in the Top6A dimer (Corbett et al., 2007; Graille et al., 2008); however, a prior AFM study has reported that topo VI can bend DNA by 100–140° (Thomson et al., 2015). Modeling DNAs with varying bend angles into structures of S. shibatae topo VI, which was captured in a splayed-open B-subunit conformation (Graille et al., 2008), suggested that a ~70 bp duplex with a ~100˚ bend could span both H2TH domains in a topo VI holoenzyme, running along the helical Stalk/WKxY region of the Top6B transducer domains and through the Top6A catalytic center (Figure 7A). Based on this model, a ~30 bp duplex would fully engage the Top6A dimer and one Stalk/WKxY element, whereas a ~40 bp duplex would be sufficient to span both Stalk/WKxY elements in a Top6A/Top6B heterotetramer (Figure 7B). An extended G-segment interface of this nature would not only provide a physical rationale for the marked increase in affinity of topo VI for duplex DNA as substrate length is increased from 20 to 30 bp (Figure 1A), but also would account for the observed DNA binding deficiencies exhibited by the Stalk/WKxY mutants (Figure 5A). Similarly, the impaired DNA binding of the KGRRAAA mutant (Figure 5—source data 1) may reflect its apparent altered conformational state (Figure 6—figure supplement 1), which might misalign the G-segment-binding surfaces of the B and A subunits to lower the affinity of the enzyme for duplexes that are not already pre-bent.

Figure 7 with 2 supplements see all
Topo VI requires H2TH-mediated, nucleotide-dependent bending of a 70 bp duplex G-segment to induce cleavage.

(A) Model of a 70 bp bent duplex which spans dimer-related H2TH domains through the TOPRIM/Winged Helix Domain cleavage site of Top6A (using a previously published SAXS model of S. shibatae topo VI with Top6B in an open conformation (Corbett et al., 2007) see also PDB ID: 2ZBK for a similar conformation, stabilized by the inhibitor radicicol (Graille et al., 2008)). Domains are colored as in Figure 3A. DNA was modeled with a continuous bend using web 3DNA (Zheng et al., 2009). (B) Schematic of the estimated duplex lengths needed to span across the H2TH, Stalk/WKxY, and Top6A dimer DNA-binding regions, using the G-segment path modeled in Figure 7A. Note that the Stalk/WKxY region may allow for the asymmetric binding of DNA in different registers, accounting for the jump in affinity seen between 20 and 30 bp DNA duplexes. (C) Nucleotide-dependent cleavage of fluorescein-labeled DNA duplexes by topo VI and mutant constructs. Length-dependent cleavage by wildtype (left), cleavage of a 70 bp duplex by basic-to-neutral mutants (middle), and cleavage by basic-to-acidic mutants (right) was tested. Cleavage reactions containing a 2:1 ratio of enzyme:duplex were run on denaturing PAGE to separate reaction products, and were visualized using a laser gel scanner. Enzyme construct (wildtype, KGRRAAA, KGRREEE, Stalk/WKxYAAA, Stalk/WKxYEEE, H2THAAA, or H2THEEE), duplex length (40 bp, 60 bp, or 70 bp), and addition of 1 mM ATP or 1 mM AMPPNP is noted above each lane. A no enzyme control containing 1 mM AMPPNP and a single strand DNA ladder consisting of 20, 30, 40, 60, 70, and 80 nt oligonucleotides were run for reference. Where present, the percentage of cleavage product relative to intact DNA is quantified above the lane. (D) Nucleotide-dependent bending of a Cy5/Cy5.5-labeled 70 bp duplex was assessed using bulk FRET. Fluorescence emission spectra (left) produced by 630 nm excitation of the Cy5-Cy5.5-labeled DNA show an increase in cy5.5 emission in the presence of topo VI (left, inset) and AMPPNP, but not in the presence of AMPPNP alone. Spectral emission was normalized by total emission from 645 nm to 850 nm. Ratiometric FRET efficiency was monitored over time upon addition of AMPPNP for the noted basic-to-neutral mutant (middle) or basic-to-acidic topo VI mutant (right). Wildtype and duplex alone are shown in each case for comparison. Figure 7—figure supplement 1 confirms FRET changes arise from DNA bending. Figure 7—figure supplement 2 further considers the gate closure activity of KGRRAAA on the 70 bp duplex substrate. Numerical data are reported in Figure 7—source data 1.

https://doi.org/10.7554/eLife.31724.031

If the H2TH domains do engage G-segment DNAs, they do not contribute appreciably to DNA binding, at least as judged by the affinity of the 70 bp duplex for wildtype topo VI compared to shorter duplexes (Figure 1A). We therefore considered whether the H2TH domains might instead help bend DNA, serve as sensors for pre-bent substrates, and/or help couple B-subunit dimerization to G-segment cleavage or strand passage. To test these ideas, we first assessed the minimal length of DNA required for nucleotide-dependent G-segment cleavage. Topo VI was incubated with 40, 60, or 70 bp long 5’-labeled duplexes in the absence of nucleotide, or with ATP or AMPPNP. Reactions were analyzed by denaturing urea-formamide PAGE to separate cleaved and uncleaved oligonucleotide products. Although the absence of a T-segment strongly inhibits G-segment scission, topo VI produced clear cleavage products in the presence of either ATP or AMPPNP on the 70 bp duplex. Faint cleavage products were also produced from the 60 bp duplex, but only in the presence of AMPPNP. The length of the cleavage products suggest M. mazei topo VI is cutting DNA slightly off-center from the preferred site identified for its S. shibatae homolog; these products are instead consistent with strand scission occurring at a secondary site six nucleotides upstream of this locus (Buhler et al., 2001). No cleavage was seen for any condition on the 40 bp duplex (Figure 7C).

We next assessed whether the H2TH domains play a role in the observed length dependence of the G-segment cleavage reaction by measuring the nucleotide-dependent cleavage activity of our functional mutant panel on a 70 bp duplex (Figure 7C). The KGRR mutants showed a slight decrease in AMPPNP-dependent cleavage, while the two Stalk/WKxY mutants displayed a greater decrease in this activity (the triple glutamate substitution proved the most severely compromised). These results are consistent with the impaired affinities that these mutant enzymes show for the 70 bp substrate (Figure 5A). By contrast, neither H2TH mutant proved capable of supporting short duplex cleavage. Collectively, these findings support the idea that for a G-segment to bind productively to the Top6A dimer, it ideally should be sufficiently long to engage both the stalk and H2TH regions of Top6B. The inability of a 40 bp duplex to support cleavage, even though this DNA binds with higher affinity than a 20 bp duplex and is long enough to reach both Stalk/WKxY regions, suggests G-segment DNAs must engage at least one H2TH domain before strand scission can be triggered.

One implication of H2TH contacts with the distal arms of an associated G-segment is that ATP-binding and ATPase domain dimerization might in turn alter G-segment bending. To test this prediction, we labeled opposing ends of the 70 bp duplex with Cy5 and Cy5.5 and monitored changes in the end-to-end distance by FRET for native topo VI and our panel of mutants. Bulk FRET efficiencies in the absence and presence of enzyme were measured by exciting Cy5 at 630 nm and scanning the spectral emission of both the donor and acceptor fluorophores (Figure 7D). The time-dependent conformational response to the addition of AMPPNP was also assessed. The addition of wild-type topo VI alone to the labeled DNA led to a modest FRET increase, a result indicative of G-segment bending that accords with prior AFM data (Thomson et al., 2015). The KGRR mutants produced a similar FRET increase; however, both sets of Stalk/WKxY and H2TH mutants yielded only a minor nucleotide-independent response (between that of wildtype topo VI and the free duplex). Upon adding AMPPNP, FRET efficiency rapidly increased further for the labeled DNA incubated with topo VI, or the KGRRAAA or KGRREEE mutants, indicating that nucleotide-driven dimerization of the ATPase regions leads to additional DNA bending. This FRET increase did not occur when Cy5 and Cy5.5 were placed on separate duplexes (Figure 7—figure supplement 1), allowing us to attribute the observed changes in FRET with the doubly labeled DNA to intramolecular bending, rather than the binding of two segments in trans. While this result for the KGRR constructs initially appeared to contradict the inability of nucleotide to alter Top6B conformation in the KGRRAAA mutant on supercoiled DNA (Figure 6D), we note that the substrate differed between these two experiments. Performing the Top6B dimerization experiment with excess, unlabeled 70 bp duplex showed that, similar to wildtype topo VI bound to linear DNA, the KGRRAAA mutant adopts a more open conformation when bound to the 70 bp duplex than when bound to supercoiled DNA, and that the addition of nucleotide can shift the conformational equilibrium of the enzyme toward a closed state (Figure 7—figure supplement 2). For their part, both Stalk/WKxY mutants produced a FRET increase in the presence of AMPPNP, albeit with substantially slowed kinetics that likely account for their negligible ATPase activities (Figure 4B). By contrast, the H2TH mutants did not support any nucleotide-dependent increase in FRET, indicating that Top6B dimerization in these mutants no longer introduces DNA bending to the distal ends of a bound G-segment. Together, these observations both suggest that topo VI engages a G-segment using an extended interface that runs from one H2TH domain to the other, and that strand scission is stimulated by bending induced by Top6B dimerization. The observation that the ATPase activity of the H2TH mutants is decoupled from strand passage, yet also substantially impaired when compared to the futile cycling of the KGRR mutants (Figure 4), further suggests that there is a feedback mechanism which couples nucleotide turnover to efficient G-segment deformation and cleavage.

Discussion

The ATPase elements of type IIB topoisomerases engage supercoiled DNA to regulate DNA strand passage

Using a broad range of functional and reporter assays (summarized in Tables 12), we show here that type IIB topoisomerases preferentially engage the DNA crossings and bends of supercoiled substrates, and that binding to supercoiled DNA in turn stimulates the nucleotide-dependent dimerization of Top6B and couples this movement to DNA cleavage and strand passage at a distance in Top6A. To recognize and exploit distinguishing features of supercoiled substrates, topo VI uses several previously unidentified DNA-binding elements integrated into Top6B, including: (1) a basic interface formed along the subunit’s C-terminal stalk and a conserved WKxY motif that is important for robust G-segment binding (Figure 3E, Figure 5A), (2) a basic ‘KGRR’ loop in the GHKL domain that aids DNA crossing recognition and links controlled ATP turnover to productive strand passage (Figure 3D, Figure 4, Figure 6), and (3) an H2TH DNA-binding domain that promotes nucleotide-dependent G-segment bending and links ATP turnover to DNA cleavage and strand passage (Figure 3F, Figure 7C–D). Collectively, our data highlight new intermediate steps in the topo VI catalytic cycle (Figure 8) and provide a molecular rationale for the essential role of Top6B in driving transesterase activity by Top6A (Buhler et al., 1998; Buhler et al., 2001). By demonstrating that efficient and productive Top6B dimerization requires nucleotide, supercoiled DNA, and an intact KGRR loop, our findings also suggest that the previously visualized, inactive conformation of the Top6A dimer (Corbett et al., 2007; Graille et al., 2008; Nichols et al., 1999) may represent a cleavage-suppression mechanism that can only be overcome when the regions identified here are occupied by the binding of an extended DNA crossing and when nucleotide induces the dimerization of Top6B.

Figure 8 with 1 supplement see all
A new model for the Type IIB topoisomerase catalytic cycle.

Free topo VI (A) binds to linear DNA (B), but preferentially engages DNA crossings (C). Binding to a hooked DNA crossing (C) by the KGRR loop and Stalk/WKxY region (step 1) induces a conformational change that presets Top6B dimerization. From this state ATP binding (step 2) introduces H2TH-dependent G-segment DNA bending and shifts the catalytic tyrosines on the Winged Helix Domain (WHD) into a cleavage-competent conformation, thus committing the enzyme to strand passage and ATP hydrolysis. While the KGRR/T-segment interaction stabilizes Top6B dimerization, T-segment capture potentiates DNA-gate opening by introducing strain in the storage cavity (step 3). T-segment release allows for DNA-gate closure and G-segment religation (step 4). Without a DNA crossing to stabilize Top6B closure, ADP and Pi are released, the WHDs relax to an inactive conformation (step 5) and Top6B returns to a relaxed, open conformation (step 6). From this G-segment bound state (B) topo VI tends to dissociate from DNA (to state A), but will infrequently capture another T-segment, regenerating a DNA crossing (C). The mechanistic implications of this model for meiotic recombination systems are considered in Figure 8—figure supplement 1.

https://doi.org/10.7554/eLife.31724.035
Table 1
Summary of topo VI functional activities and mutant effects.
https://doi.org/10.7554/eLife.31724.037
AssayWildtype activityEffect of mutation to:
KGRR loopStalk/WKxYH2TH
Supercoil relaxationdistributive strand passage activitykills strand passagekills strand passagegreatly impairs strand passage
ATPase activity
-on linear DNAstimulates above basal activityno activityno activity~wildtype activity
-on supercoiled DNAstimulates more than linear DNAincreased activity, futile cyclingno activity~wildtype activity, futile cycling
DNA binding
-short duplexesaffinity increases from 20 bp to 40 bp in lengthmoderately impairs binding for longer duplexesgreatly impairs bindingslightly impairs binding for longer duplexes
-sheared salmon-sperm DNAsimilar affinity as for 40–70 bp duplexesmoderately impairs bindingN.D.slightly impairs binding
-supercoiled DNAincreased affinity compared to linear DNAmoderately impairs bindingN.D.moderately impairs binding
-stacked junctiontighter binding than to duplexgreatly impairs bindingN.D.~wildtype affinity
Top6B dimerization
-on short DNA duplexesDNA promotes closure
AMPPNP promotes further closure
loss of substrate promoted closure
AMPPNP promotes some closure
N.D.N.D.
-on supercoiled DNApromotes greater closure than linear DNA
AMPPNP promotes further closure
supercoiled DNA promotes closure loss of AMPPNP promoted closureN.D.weaker substrate dependent closure than wildtype AMPPNP promotes further closure
Short duplex cleavageAMPPNP promotes cleavage on 60 and 70 bp duplexessimilar to wildtypegreatly impairs cleavageno cleavage
Short duplex bendingAMPPNP promotes bendingsimilar to wildtypegreatly slows bendingno bending
Table 2
Summary of enzyme, DNA and nucleotide conditions by type of experiment.
https://doi.org/10.7554/eLife.31724.038
Assay[enzyme][nucleotide][DNA]
DNA binding0, 3.9–4000 nMN/A20 nM (0.4–1.4 μM bp) probe duplex
Competitive binding100 nMN/A20 nM (1.4 μM bp) probe duplex
0, 0.1–106 μM bp DNA competitor/
0, 0.3–36 nM plasmid
Supercoil relaxation
titration
timecourse/chase
 
0, 0.3-20 nM
2.5 nM
 
1 mM ATP
1 mM ATP
2.9 kb primary plasmid-
10.2 μM bp DNA/3.5 nM plasmid
6.5 kb chase plasmid-
10.2 μM bp DNA/1.6 nM plasmid
ATP hydrolysis-
ATP titration
DNA titration
 
500 nM
500 nM
 
0, 0.06–4 mM ATP
2 mM ATP
 
400 μM bp DNA/136 nM plasmid
0, 3.1–800 μM bp DNA/
0, 1–273 nM plasmid
Top6B dimerization200 nM1 mM AMPPNP100 μM bp DNA/34 nM plasmid
Short duplex cleavage200 nM1 mM ATP or
AMPPNP
100 nM duplex (7 μM bp DNA)
Short duplex bending200 nM1 mM AMPPNP100 nM duplex (7 μM bp DNA)

Besides promoting Top6B closure, our data also imply that T-segment engagement may actively control both ATP turnover and DNA-gate opening to permit strand passage. As with wild-type topo VI, the binding of the KGRRAAA mutant to supercoiled DNA alone promotes Top6B dimerization (Figure 6—figure supplement 1) and supports ATP hydrolysis (Figure 4B); however, disruption of this region impairs the binding of a DNA crossover (Figure 6C), does not support additional conformational response to nucleotide (Figure 6D), blocks ATP hydrolysis on short linear DNA (Figure 4B), and abolishes strand passage overall (Figure 4A). This behavior suggests the bends or pre-formed crossings present in supercoiled DNA help to promote B-subunit dimerization and ATP hydrolysis, and may partially compensate for a loss of the KGRR element (Figure 4B, Figure 6D), but that the coupling of ATP hydrolysis to strand passage requires the productive binding of a T-segment DNA (Figure 2B–D, Figure 6D). Given that the T-segment storage cavity appears to be too small to accommodate DNA when fully closed (Corbett et al., 2007; Corbett and Berger, 2005; Graille et al., 2008), it has been proposed that T-segment engagement may potentiate opening of the Top6A dimer and separation of a cleaved G-segment. Considering this idea in light of our present findings suggests that the binding of the KGRR loops to a stored T-segment helps to suppress premature release of ATP hydrolysis products, which is normally linked to a slow conformational change associated with G-segment separation and subsequent T-segment release. This scheme offers a simple explanation for why the KGRR mutants rapidly hydrolyze ATP when bound to supercoiled DNA: weakening of the T-segment interaction with the KGRR loop allows for early ATP turnover, yet by not resolving the DNA crossing, Top6B remains pre-dimerized and primed to bind ATP again, leading to futile cycling without strand passage.

Topo VI’s enzymatic properties seem oddly mismatched to the expected demands of the cell

Certain biochemical properties of topo VI identified here are somewhat surprising when considering the demands placed on the cell by transcription and replication. For example, the highly distributive nature of supercoil relaxation observed for topo VI (Figure 1C–D) is at odds with a need to remove the continual local build-up of superhelical tension arising from RNA polymerase advancement or replication fork progression. The maximal observed ATP hydrolysis and strand passage rates for Mm topo VI in vitro (Figure 1C, Figure 2A) are also much slower (~50–100 fold) than rates generally observed for type IIA topoisomerases (Higgins et al., 1978; Lindsley and Wang, 1993; Osheroff et al., 1983; Sugino and Cozzarelli, 1980). These enzymatic properties raise important questions as to when and in which context topo VI acts in the cell. For instance, using estimates based on the genome size and generation time of M. mazei (Appendix 1), topo VI would appear to require ~50-fold greater specific activity to keep up with gene expression and chromosome duplication, or else be present at extremely high cellular concentrations.

Although the source of this discrepancy may be due to differences between in vitro vs. in vivo rates of strand passage, it could alternatively arise from a missing factor that enhances topo VI activity. This second explanation, if true, has intriguing ramifications. For example, if a secondary factor were to accelerate topo VI’s strand passage rate by increasing processivity, then the distributive action of topo VI might reflect an auto-inhibitory mechanism that is manifest until the enzyme is localized to the appropriate chromosomal context. Along this line, multiple protein factors have been identified to bind topo VI in Arabidopsis thaliana, and may be obligate components of the topo VI machinery in plants (Breuer et al., 2007; Forterre and Gadelle, 2009; Kirik et al., 2007; Sugimoto-Shirasu et al., 2005). If analogous factors exist for archaeal topo VI, it may be that Top6A and Top6B actually constitute the core of a larger type IIB topoisomerase complex. Eukaryotic topo IIIα, a type IA topoisomerase, exemplifies such a strategy, interacting with a RecQ family helicase and SSB/RPA-like factors to channel DNA strand passage into efficient Holliday junction resolution (Plank et al., 2006; Raynard et al., 2006; Singh et al., 2008; Wu et al., 2006; Xu et al., 2008).

Implications of functional features of type IIB topoisomerases for homologous ATPase, nuclease, and transesterase systems

The picture of the type IIB topoisomerase strand passage mechanism developed here reveals a rich set of regulatory mechanisms both shared with and divergent from type IIA topoisomerases. Of these, a central feature is the requirement that topo VI must engage a DNA crossing (as found in supercoiled or catenated DNA) to access a stable dimerized B-subunit conformation and productively turn over ATP (Figure 8). Contrariwise, Top6B takes on a predominantly open conformation when bound only to a prospective G-segment (or in the absence of DNA [Figure 2, Corbett et al., 2007; Graille et al., 2008]), even when nucleotide is present. This tight control over Top6B dimerization, and its reciprocal coupling to G-segment cleavage, likely helps compensate for the missing safeguard of a third dimerization interface – the so-called ‘C-gate’ – found in type IIA topoisomerases (Roca et al., 1996; Roca and Wang, 1994; Williams and Maxwell, 1999). For its part, the ATPase region of type IIA topoisomerases does possess potential T-segment-sensing elements (Tingey and Maxwell, 1996); however, in contrast to topo VI, the ATPase domains of type IIA topoisomerase holoenzymes readily dimerize upon binding ATP, even when DNA is absent (Gubaev and Klostermeier, 2011; Roca and Wang, 1992). Interestingly, in requiring the binding of a substrate T-segment for stable ATPase domain dimerization, topo VI echoes the behavior of Hsp90, whose related GHKL ATPase fold strongly depends on client protein or co-chaperone engagement to drive ATPase association (Ali et al., 2006; Hessling et al., 2009; Wolmarans et al., 2016). This co-dependency raises the possibility that other GHKL ATPases, such as MutL and MORC proteins, may similarly rely on substrate/cofactor interactions as a checkpoint to license nucleotide-dependent dimerization.

Following DNA crossover recognition, ATP binding by Top6B is needed to trigger G-segment scission by Top6A (Buhler et al., 1998). However, closing of the ATP gate also serves to further bend the G-segment through contacts mediated by the H2TH domain (Figure 7D). Although H2TH mutants are unable to cleave short duplex substrates (Figure 7C), they support weak strand passage activity on supercoiled DNA (Figure 4A), a substrate that constrains DNA bends independent of enzyme binding. This suggests that DNA bending itself, whether innate or H2TH-mediated, may help promote DNA breakage by Top6A. Although the H2TH domain is specific to type IIB topoisomerases, type IIA enzymes also bend G-segment DNAs to support cleavage (Dong and Berger, 2007; Laponogov et al., 2009; Lee et al., 2013; Lee et al., 2012; Wohlkonig et al., 2010). This dependency raises the possibility that other nucleases or transesterases that rely on the TOPRIM fold beside type II topoisomerases (e.g. OLD family enzymes and Spo11 (Aravind et al., 1998)) may similarly require DNA deformation to promote strand scission.

The discovery that Spo11 was related to the DNA-cleaving Top6A subunit of archaeal topo VI was a critical development in understanding how DNA breaks are formed to initiate meiotic recombination (Bergerat et al., 1997; Keeney et al., 1997). The realization that Top6A requires Top6B for DNA cleavage (Buhler et al., 1998) has in turn raised the question of whether Spo11 might partner with a similar regulatory factor during meiotic recombination. Recently, structurally homologous counterparts to Top6B have been recognized across a wide range of eukaryotic species (MTop6B in plants, Top6BL in mammals, Rec102 in S. cerevisiae, and Mei-P22 in Drosophila) (Robert et al., 2016; Vrielynck et al., 2016). Interestingly, the WKxY motif implicated here in G-segment binding is conserved between Top6B and some of its meiotic homologs (e.g. mammalian MTop6B and plant Top6BL) (Robert et al., 2016), suggesting that this region could assist Spo11 with DNA targeting, and contribute to the signals necessary to activate DNA cleavage during meiosis. In those Top6B homologs where the WKxY motif is poorly conserved, alternative features on the transducer stalk may participate in binding to DNA. For example, the prospective WKxY motif in budding yeast Rec102 is highly divergent in sequence (WEEQ), yet Spo11 hotspots from this organism display a sequence bias that extends beyond the predicted footprint of the Spo11 dimer. Interestingly, this bias maps to a distance of ±11–16 bp from the dyad of Spo11 (Pan et al., 2011), compared to the ~17–20 bp distance between the Top6B Stalk/WKxY region and the Top6A dyad, consistent with the notion that non-Spo11 DNA interaction sites may have shifted during evolution.

In topo VI, we find that Top6B dimerization further bends DNA to potentiate cleavage by Top6A. Surprisingly, components critical for Top6B-mediated dimerization are either highly divergent or missing in meiotic Top6B homologs. For example, both Topo6BL and MTopo6B contain a highly degenerate GHKL domain that lacks essential elements required for ATP binding (only purine-binding elements are conserved, see Figure 8—figure supplement 1), and Rec102 and Mei-P22 lack a GHKL domain entirely (Dutta and Inouye, 2000; Robert et al., 2016; Vrielynck et al., 2016). Insofar as DNA bending, the meiotic Top6B-like factors identified thus far also lack an H2TH domain (Robert et al., 2016; Vrielynck et al., 2016). Should Spo11, like Top6A, require both DNA bending and allosteric activation to achieve a cleavage-competent state, these differences indicate that it is not the newly identified Top6B-like subunits alone that are responsible for mediating this event. Candidate factors that might further regulate Spo11-dependent break formation include additional partner proteins, post-translational modifications, and tension on or deformation of the DNA itself by factors responsible for sister chromatid pairing (Lam and Keeney, 2014). Future studies focused on defining how topo VI and Spo11-type systems physically engage DNA strands, respond to possible partner factors, and switch between inactive and active DNA-cleavage states will be needed to help shed light on how these systems operate.

Materials and methods

Key resources table
Reagent type (species)
or resource
DesignationSource or referenceIdentifiersAdditional information
Gene (Methanosarcina
Mazei)
Top6AN/ANCBI Gene ID: 1480760
Gene (Methanosarcina
Mazei)
Top6BN/ANCBI Gene ID: 1480759
Strain, strain background
(E. coli)
BL21(DE3)-RILQB3-MacroLab
Strain, strain background
(E. coli)
XL1-BlueQB3-MacroLab
Recombinant DNA reagentM. mazei Top6AB
expression vector
PMID: 17603498
Recombinant DNA reagentM. mazei Top6AB-KGRRAAA
expression vector
this paperConstruct generated by introduction
of point mutations: K186A, R188A,
and R189A to Top6B gene on M. Mazei
Top6AB expression vector
Recombinant DNA reagentM. mazei Top6AB-KGRREEE
expression vector
this paperConstruct generated by introduction
of point mutations: K186E, R188E,
and R189E to Top6B gene on M. Mazei
Top6AB expression vector
Recombinant DNA reagentM. mazei Top6AB-Stalk/
WKxYAAA expression vector
this paperConstruct generated by introduction
of point mutations: K399A, K401A, and
R457A to Top6B gene on M. Mazei
Top6AB expression vector
Recombinant DNA reagentM. mazei Top6AB-Stalk/
WKxYEEE expression vector
this paperConstruct generated by introduction
of point mutations: K399E, K401E, and
R457E to Top6B gene on M. Mazei
Top6AB expression vector
Recombinant DNA reagentM. mazei Top6AB-H2THAAA
expression vector
this paperConstruct generated by introduction
of point mutations: R263A, K268A,
and K308A to Top6B gene on M. Mazei
Top6AB expression vector
Recombinant DNA reagentM. mazei Top6AB-H2THEEE
expression vector
this paperConstruct generated by introduction
of point mutations: R263E, K268E, and
K308E to Top6B gene on M. Mazei
Top6AB expression vector
Recombinant DNA reagentM. mazei Top6AB-cyslite-
155C expression vector
this paperConstruct generated by introduction
of point mutations: T155C, C267S,
C278A, C316A, and C550A to Top6B
gene on M. Mazei Top6AB
expression vector
Recombinant DNA reagentM. mazei Top6AB-KGRRAAA
cyslite-155C expression vector
this paperConstruct generated by introduction
of point mutations: K186A, R188A,
and R189A to Top6B gene on M. Mazei
Top6AB-cyslite-155C expression vector
Recombinant DNA reagentM. mazei Top6AB-H2THAAA
cyslite-155C expression vector
this paperConstruct generated by introduction
of point mutations: R263A, K268A,
and K308A to Top6B gene on M. Mazei
Top6AB-cyslite-155C expression vector
Recombinant DNA reagentM. mazei Top6AB-E44A
expression vector
this paperConstruct generated by introduction
of point mutations: E44A to Top6B
gene on M. Mazei Top6AB
expression vector
Recombinant DNA reagentpSG483 (plasmid DNA)PMID: 160236702.9 kb plasmid used as
supercoiled substrate
Sequence-based reagent
(13 oligonucleotides)
See Figure 1—source data 1Integrated DNA
Technologies
Chemical compound, drugsalmon sperm DNA, shearedThermo Fisher ScientficThermoFisher:AM9680
Chemical compound, drugAlexa Fluor 555 C2 MaleimideThermo Fisher ScientficThermoFisher:A20346
Chemical compound, drugAlexa Fluor 647 C2 MaleimideThermo Fisher ScientficThermoFisher:A20347
Software, algorithmConSurf ServerPMID: 20478830RRID:SCR_002320
Software, algorithmw3DNA serverPMID: 19474339
Software, algorithmPyMolSchrödinger, LLCRRID:SCR_000305
Software, algorithmPrism 7Graphpad SoftwareRRID:SCR_015807

Cloning of M. mazei topo VI functional mutant vectors

Cloning of the M. mazei Top6B gene in frame with an N-terminally fused His6-tobacco etch virus (TEV) protease-cleavable tag and the M. mazei Top6A gene into a polycistronic expression vector was previously described (Corbett et al., 2007). Oligonucleotides used for site directed mutagenesis were obtained from Integrated DNA Technology (IDT, Coralville, IA). Mutant constructs were generated either by PCR amplification of the expression vector using primers containing the desired point substitutions followed by blunt-end ligation, or by quick-change mutagenesis (Agilent, Santa Clara, CA). The following mutations were added to generate the ‘Cys-lite’ construct: C267S, C278A, C316A and C550A, all in Top6B. Mutagenesis was verified by Sanger sequencing (Genewiz LLC, South Plainfield, NJ).

Protein expression and purification

Topo VI and functional mutant variants were overexpressed in E. coli BL21(DE3)Codon +RIL cells (QB3-Macrolab, University of California-Berkeley, CA) grown in ZYM-5052 auto-induction media (Studier, 2005). Wild-type topo VI was expressed in cultures grown at 37°C, whereas cultures expressing functional mutant constructs were shifted to 25°C upon reaching an OD600 of 0.4–0.6. The KGRRAAA FRET assay construct was grown at 37°C to an OD600 of 2–3 in M9ZB media (Studier, 2005), cooled to 18°C, and then induced with IPTG (250 μM final concentration) and grown overnight. Cultures were harvested by centrifugation at 24 hr following inoculation, resuspended in buffer A [20 mM HEPES-KOH pH 7.5, 800 mM NaCl, 20 mM Imidazole, 10% (v/v) glycerol, 1 μg/mL pepstatin A, 1 μg/mL leupeptin, 1 mM PMSF], and frozen drop-wise into liquid nitrogen for storage at −80°C.

Proteins were purified as previously described (Corbett et al., 2007). Harvested cells were lysed by sonication, and lysate was clarified by centrifugation. Clarified lysate was applied to a 5 mL HiTrap Ni2+ column (GE Healthcare Life Sciences, Marlborough, MA, USA) and washed with buffer A [20 mM HEPES-KOH pH 7.5, 800 mM NaCl, 20 mM Imidazole, 10% (v/v) glycerol, 1 μg/mL pepstatin A, 1 μg/mL leupeptin, 1 mM PMSF]. Following a subsequent wash with buffer B [20 mM HEPES-KOH pH 7.5, 150 mM NaCl, 20 mM Imidazole, 10% (v/v) glycerol, 1 μg/mL pepstatin A, 1 μg/mL leupeptin, 1 mM PMSF], bound proteins were eluted by a 15-column volume gradient from buffer B to buffer C [20 mM HEPES-KOH pH 7.5, 150 mM NaCl, 20 mM Imidazole, 10% (v/v) glycerol, 1 μg/mL pepstatin A, 1 μg/mL leupeptin, 1 mM PMSF]. Fractions containing the topo VI heterotetramer were applied to a 5 mL HiTrap SP cation-exchange column and 5 mL HiTrap Q anion-exchange column (GE Healthcare Life Sciences) in series and washed with buffer B. The HiTrap SP column was removed, and protein bound to the HiTrap Q column was eluted with a 10-column volume gradient from buffer B to buffer A. Peak fractions were concentrated by centrifugation (Millipore Amicon Ultra 30K MWCO) and incubated with 1.5 mg of His6-TEV protease (QB3-Macrolab, University of California, Berkeley) overnight at 4°C to remove His6 tags. Uncleaved proteins and His6-TEV protease were removed by applying the protease cleavage reaction to a HiTrap Ni2+ column equilibrated in buffer B. Flow-through was concentrated and applied to an Sephacryl-300 HR gel filtration column (GE Healthcare Life Sciences) equilibrated and run in sizing buffer [20 mM HEPES-KOH pH 7.5, 300 mM KCl, 10% (v/v) glycerol] and concentrated by centrifugation (Millipore Amicon Ultra 10K MWCO). Purity of peak fractions was assessed by SDS-PAGE and coomassie blue staining, and the concentration of tetramer was determined by absorbance at 280 nm using extinction coefficients determined by the ExPASY ProtParam webserver (Gasteiger et al., 2005). Proteins were flash frozen in a final storage buffer [20 mM HEPES-KOH pH 7.5, 300 mM KCl, 30% (v/v) glycerol, 1 mM Trisphosphine hydrochloride (TCEP)] and stored in aliquots at −80°C for use in subsequent biochemical and biophysical studies.

DNA binding and competition

DNA substrates were resuspended in ddH2O and annealed from single strand DNA oligomers of complementary sequence (Figure 1—source data 1) obtained from IDT. Annealing of the stacked-junction DNA substrate followed published protocols (Duckett et al., 1988) with a few modifications. The junction was prepared in 25 mM Tris HCl pH 7.9, 25 mM NaCl, 10 mM MgCl2 and annealed by heating at 70°C for 2 hr, followed by cooling at 0.5°C/min to 4°C. Annealing reaction products were loaded onto a 5 mL HiTrap-Q anion exchange column (GE Healthcare Life Sciences) equilibrated in stacked junction (SJ) buffer A [25 mM NaCl, 25 mM Tris-HCl pH 7.9, 10 mM MgCl2]. Contaminants were removed by washing with 55%/45% mix of SJ buffer A to SJ Buffer B [1 M NaCl, 25 mM Tris 7.9, 10 mM MgCl2]. Correctly annealed substrate was eluted with 45%/55% Buffer A/Buffer B, pooled and dialyzed back into SJ buffer A, and concentrated by centrifugation (Amicon Ultra 3K MWCO, EMD Millipore, Billerica, MA). Proper annealing for all substrates was assessed by native 15% PAGE run in 0.5x Tris-Borate-EDTA (TBE) buffer at 4°C.

DNA binding by topo VI and functional mutants was assessed using fluorescence anisotropy. Protein was serially diluted in two-fold steps in binding assay dilution buffer [250 mM potassium glutamate, 5% (v/v) glycerol, 50 mM HEPES-KOH pH 7.5 and 1 mM TCEP] and incubated with fluorescein-labeled DNA substrate in the dark and on ice for 5 min. Reactions were diluted to final binding assay conditions [27 μL, 0, 0.3–4000 nM enzyme, 20 nM labeled duplex, 50 mM potassium glutamate, 5% (v/v) glycerol, 20 mM HEPES-KOH pH 7.5, 1 mM TCEP, 10 mM MgCl2 and 0.1 mg/mL BSA], and incubated on ice an additional 10 min. Fluorescence anisotropy was measured at ambient temperature using a Clairiostar microplate reader (BMG Labtech GmbH, Ortenberg, Germany) by exciting at 482 nm (band pass 16 nm) and measuring parallel and perpendicular emission intensity at 530 nm (band pass 40 nm), with an inline 504 nm long pass dichroic filter. Data are the average of three independent experiments, with all points normalized to the DNA alone condition and fit to the following single-site binding model:

(1) FA=FAmax(L+P+Kd,app-(L+P+Kd,app)2-4[L][P]2[L])

where ΔFAmax is the maximal specific change in anisotropy, [L] is DNA substrate concentration, [P] is the concentration of topo VI construct, and Kd,app is the apparent dissociation constant for DNA substrate and enzyme. To test for cooperativity, binding isotherms were also fit to a Hill equation-type model:

(2) FA=FAmax([P]hKd,apph+[P]h)

where ΔFAmax is the maximal specific change in anisotropy, [P] is the concentration of topo VI construct, h is the apparent Hill coefficient, and Kd,app is the apparent dissociation constant for DNA substrate and enzyme.

Competition assays were carried out similarly to binding assays, with protein diluted in binding assay dilution buffer and incubated with the 70 bp fluorescein-labeled duplex and either negatively supercoiled pSG483 plasmid DNA (pBluescript SK derivative, 2927 bp) or linear sheared salmon-sperm DNA (ThermoFisher Scientific, Waltham, MA). Reactions were diluted to final binding assay conditions, except enzyme concentration was set at 100 nM, and competitor concentration varied from 0.1 μM bp to 106.5 μM bp DNA. Anisotropy data were fit to an explicit competition model (Wang, 1995), which fits to the parameters: [A], total concentration of the competitor DNA substrate; [B], total concentration of the labeled DNA probe; [P], total topo VI concentration; KA, dissociation constant of the competitor DNA substrate; KB, dissociation constant of the labeled DNA probe; and ΔFAmax, the maximal specific change in fluorescence anisotropy for the probe.

Supercoiled DNA relaxation

Topo VI holoenzyme was thawed and diluted in series with relaxation assay dilution buffer [300 mM potassium glutamate, 10% (v/v) glycerol, 20 mM HEPES-KOH pH 7.5 and 1 mM TCEP] and incubated with negatively supercoiled pSG483 plasmid DNA for 5 min on ice before dilution into final relaxation assay conditions [30 μL reactions, 0, 0.3–20 nM topo VI for titration, 2.5 nM topo VI for timecourses, 50 mM potassium glutamate, 10% (v/v) glycerol 20 mM bis-tris-propane-HCl (BTP-HCl) pH 7.5, 2 mM HEPES pH 7.5, 1 mM TCEP, 10 mM MgCl2, 0.1 mg/mL BSA, 3.5 nM pSG483 (10.2 μM bp DNA), and 1 mM ATP]. Reactions were initiated by addition of ATP, incubated at 30°C, and quenched by addition of SDS and EDTA to final concentrations of 1% and 10 mM respectively. Glycerol-based loading dye was added to samples which were run on a 1% (w/v) TAE agarose gel (40 mM sodium acetate, 50 mM Tris-HCl, pH 7.9 and 1 mM EDTA, pH 8.0) for 15 hr at ~2 V/cm. For visualization, gels were stained for 30 min with 0.5 μg/mL ethidium bromide in running buffer, de-stained in running buffer for 30 min, and exposed to UV trans-illumination. Experiments were carried out similarly for the plasmid-chase experiments, except that a 6.5 kb chase plasmid (p1C) was added with ATP to a final concentration of 10.2 μM bp when initiating reactions.

Steady state ATP hydrolysis

ATP hydrolysis was measured using an established NADH-coupled assay (Morrical et al., 1986; Tamura and Gellert, 1990). Topo VI was thawed and diluted with 300 mM potassium glutamate, 10% (v/v) glycerol, 50 mM BTP-HCl pH 7.5 and 5 mM TCEP to 3.75 μM enzyme, mixed 1:2 with sheared salmon-sperm DNA, supercoiled pSG483, or ddH2O, and incubated for 5 min on ice. Enzyme/substrate mixes were diluted with NADH-PK/LDH coupling mix to final ATP hydrolysis assay conditions [100 μL reactions, 3.75 mM phosphoenolpyruvate, 150 μM NADH, 24 U pyruvate kinase and 36 U lactate dehydrogenase (PK/LDH from rabbit muscle in buffered, aqueous glycerol solution, Sigma Aldrich, St Louis, MO), 0.1 mg/mL BSA, 50 mM BTP-HCl, pH 7.5, 50 mM potassium glutamate, 5 mM TCEP, 10 mM MgCl2, 5% (v/v) glycerol, 500 nM topo VI holoenzyme]. ATP titration reactions contained either 400 μM bp sheared salmon-sperm DNA, 400 μM bp negatively supercoiled pSG483 or no DNA, and were initiated by addition of ATP to a final concentration of 0 mM or 62.5 μM-4 mM diluted in two-fold steps. DNA titrations containing 3.12–800 μM bp DNA diluted in two-fold steps were initiated by addition of ATP to a final concentration of 2 mM. Reactions were incubated at 30°C and followed in clear 96-well plates (Corning Inc, Corning, NY) by absorbance at 340 nm using a Clairiostar microplate reader. Raw absorbance values were converted to NADH molar concentrations based on measurements from NADH standards in the final ATP hydrolysis assay condition. ATP hydrolysis rates were determined by fitting to the linear portion of NADH consumption curves. Data representing three independent experiments were fit to a standard Michaelis-Menten model:

(3) V0=kcat,appET[S]Km,app+[S]

where V0 is the observed turnover rate, kcat,app is the maximum turnover rate, [Et] is the total topo VI holoenzyme concentration, [S] is the concentration of ATP, and Km,app is the Michaelis constant for ATP. For DNA titration experiments, [S] is the concentration of DNA and the kcat-stim,DNA and Kstim,DNA parameters substitute for kcat,app and Km,app.

Top6B dimerization assessed by FRET

Following purification, topo VI FRET constructs were labeled by reacting enzyme with 5-fold molar excess to enzyme of both Alexa Fluor 555 C2 maleimide and Alexa Fluor C2 647 maleimide (ThermoFisher Scientific) in sizing buffer overnight at 4°C. TCEP was also added at 50-fold molar excess to enzyme. Reactions were quenched with 5 mM DTT and applied to a HiPrep 26/10 Desalting column (GE Healthcare Life Sciences) to separate protein from unreacted dye. Proper labeling was imaged by SDS PAGE using a Typhoon FLA 9500 laser scanner (GE Healthcare Life Sciences). Labeling efficiencies were determined by comparing absorption at 280 nm for protein to absorption at 555 nm for Alexa555 and 650 nm for Alexa647. Proteins were brought to storage buffer conditions, flash frozen as aliquots in liquid nitrogen and stored at −80°C.

For gate closure assays, labeled protein was diluted in 250 mM potassium glutamate, 10% (v/v) glycerol and 20 mM HEPES-KOH pH 7.5 to 1 μM, mixed 1:1 with 500 μM bp DNA substrate or ddH2O, incubated on ice for 5 min, and diluted to final assay conditions [20 μL reactions, 200 nM topo VI, 0 or 100 μM bp DNA, 50 mM potassium glutamate, 1 mM TCEP, 10% (v/v) glycerol, 20 mM HEPES-KOH pH 7.5, 10 mM MgCl2 and 0.1 mg/mL BSA]. Fluorescence emission spectra were measured by exciting samples at 530 nm and measuring emission from 545 nm to 700 nm using a Fluoromax Fluorometer 4 (HORIBA Jobin Yvon, Edison, NJ). Adenylyl-imidodiphosphate (AMPPNP) was added to a final concentration of 1 mM and changes to emission spectra were measured over time. Spectra were normalized by total emission intensity. Plotted FRET efficiencies (E) were determined ratiometrically from donor (ID) and acceptor (IA) peak intensities:

(4) E=IAID+IA

Short DNA duplex cleavage

Topo VI was diluted in [250 mM potassium glutamate, 10% (v/v) glycerol, 10 mM MgCl2 and 20 mM HEPES-KOH pH 7.5] to 1 μM, mixed 1:1 with 500 nM fluorescein-labeled duplex (Figure 1—source data 1) and incubated 5 min on ice. Reactions were diluted to a final cleavage reaction condition [20 μL reactions, 200 nM topo VI construct, 100 nM FAM-labeled duplex, 50 mM potassium glutamate, 1 mM TCEP, 10% (v/v) glycerol, 16 mM BTP-HCl pH 7.5, 4 mM HEPES-KOH pH 7.5, 10 mM MgCl2, 0.1 mg/mL BSA and 15% DMSO]. ATP, AMPPNP or ddH2O were added to initiate reactions. Reactions were incubated at 30°C for 2 hr then quenched with SDS to a final concentration of 1%. Proteinase K was added to reactions at a final concentration of 0.3 mg/mL and incubated at 45°C for 1 hr. Formamide was added 1:1 to samples and cleavage products were separated on 7 M Urea-Formamide 0.5x TBE 12% PAGE. Gels were visualized using a Typhoon FLA 9500 laser scanner.

DNA bending assessed by FRET

DNA bending experiments used the same 70 bp duplex sequence from binding and cleavage experiments, except the substrate was modified to have a Cy5 replace the 5’-fluorocein on strand one and Cy5.5 was added to the 5’ end of strand 2 (Figure 1—source data 1). Reactions were prepared exactly as described for the DNA cleavage assays. Fluorescence emission spectra were measured by exciting samples at 630 nm and measuring emission from 645 nm to 850 nm using a Fluoromax Fluorometer 4. AMPPNP was added to a final concentration of 1 mM and changes to emission spectra were measured over time. Spectra were normalized by total spectral emission. Plotted FRET efficiencies were calculated as for the gate closure assays.

Expression and purification of S. cerevisiae topoisomerase II1-1177 (ScTop2ΔCTR) and ScTop2 ΔCTR-cyslite-180C

A ScTop2 construct containing labeling sites on the ATP gate (ScTop2ΔCTR-cyslite-180C) was generated from a previously described ScTop2 construct with a C-terminal truncation (coding for residues 1–1177, ScTop2ΔCTR, [Schmidt et al., 2012]) cloned in frame with an N-terminally fused His6-TEV protease-cleavable tag by introducing the following mutations: C48A, C381A, C471A, C731A. Proteins were overexpressed and purified as previously described (Schmidt et al., 2012). In brief, S. cerevisiae strain BCY123 was transformed with a GAL1 shuttle vector containing the ScTop2ΔCTR ORF and grown in CSM-Ura- media with a 2% lactic acid and 1.5% glycerol carbon source at 30°C. Overexpression was induced by the addition of 2% galactose at A600 = 0.8. Six hours following induction, cells were centrifuged, resuspended in 1 mM EDTA and 250 mM NaCl (1 mL/L liquid culture), and flash frozen drop-wise in liquid nitrogen.

For purification, frozen cells were first lysed under liquid nitrogen using an SPEX SamplePrep 6870 Freezer Mill (SPEX SamplePrep, Metuchen, NJ), and resultant powder was thawed and re-suspended in Buffer A300 [20 mM Tris-HCl pH 8.5, 300 mM KCl, 20 mM imidazole, and 10% (v/v) glycerol, 1 μg/mL pepstatin A, 1 μg/mL leupeptin, and 1 mM PMSF]. Lysate was clarified by centrifugation and applied to a 5 mL HiTrap Ni2+ column equilibrated in buffer A. Following washing with buffer A, protein was eluted with buffer B [20 mM Tris-HCl pH 8.5, 100 mM KCl, 200 mM imidazole, and 10% (v/v) glycerol, 1 μg/mL pepstatin A, 1 μg/mL leupeptin, and 1 mM PMSF], and applied to a 5 mL HiTrap SP cation-exchange column. Bound protein was eluted with buffer C [20 mM Tris-HCl pH 8.5, 500 mM KCl, 10% (v/v) glycerol, 1 μg/mL pepstatin A, 1 μg/mL leupeptin and 1 mM PMSF]. Peak fractions were concentrated by centrifugation (Millipore Amicon Ultra 30K MWCO) and incubated with 1.5 mg of His6 TEV protease overnight at 4°C. Uncleaved proteins and TEV protease were removed by applying the protease reaction to a HiTrap Ni2+ column equilibrated in buffer A. Flow-through was concentrated and applied to an Sephacryl-300 HR gel filtration column (GE) equilibrated and run in ScTop2 sizing buffer [20 mM Tris-HCl pH 7.9, 500 mM KCl, 10% (v/v) glycerol]. Peak fractions were collected and concentrated (Millipore Amicon Ultra 30K MWCO). Purity was estimated by SDS-PAGE and concentration was determined by absorbance at 280 nm. ScTop2 was flash frozen in a final storage buffer containing [20 mM Tris-HCl pH 7.9, 500 mM KCl, 30% (v/v) glycerol] and stored in aliquots at −80°C.

Supercoiled DNA relaxation by ScTop2

Plasmid relaxation assays and chase assays with ScTop2ΔCTR were carried out as described for topo VI, except that ScTop2ΔCTR was diluted in [500 mM KCl, 10% (v/v) glycerol, 20 mM Tris-HCl pH 7.9] and final relaxation assay conditions were [30 mM Tris-HCl pH 7.9, 10 mM MgCl2, 0.05 mg/mL BSA, 0.5 mM TCEP, 100 mM KCl, 10% (v/v) glycerol, 1 mM ATP, 3.5 nM (10.2 μM bp DNA) pSG483, and 2.5 nM topo II], with 10.2 μM bp of the 6.5 kb plasmid added with ATP to initiate reactions for chase experiments.

S. cerevisiae topo II ATPase domain dimerization assessed by FRET

The ATP gate of ScTop2ΔCTR-cyslite-180C was labeled on a native cysteine residue (180C) with the Alexa Fluor 555 C2 maleimide and Alexa Fluor C2 647 maleimide FRET pair following the same procedure as for Top6B, except that the reaction was carried out in ScTop2 sizing buffer and samples were flash frozen in the topo II storage buffer conditions.

Gate closure assays were performed similarly as with topo VI, except protein was diluted in 500 mM KCl, 10% (v/v) glycerol and 20 mM Tris-HCl pH 7.9, and final assay conditions were 200 nM topo II, 0 or 100 μM bp DNA, 100 mM KCl, 2% (v/v) glycerol, 10 mM Tris-HCl pH 7.9, 5 mM MgCl2 and either 0 mM or 1 mM AMPPNP. Fluorescence emission spectra were measured as with topo VI.

Data analysis and figure preparation

All data were plotted and fit using Prism Version 7 (RRID: SCR_015807, (GraphPad Software, La Jolla, CA)). Mapping of sequence conservation in relation to tertiary structure was aided by the Consurf web server (RRID: SCR_002320, [Ashkenazy et al., 2010]). Coordinates for bent DNA models were generated using the 3DNA web server (Zheng et al., 2009). Pymol was used for structure visualization and comparison (RRID: SCR_000305, [The PyMOL Molecular Graphics System, Schrödinger, LLC]).

Appendix 1

Supplementary material

Comparing the expected workload of M. mazei topo VI to model type IIA topoisomerases

To assess whether the enzymatic activity reported for Mm topo VI could satisfy its cellular workload in archaea, we began by estimating the in vitro strand passage rate for the enzyme. Assuming an average super-helical density of σ = −0.06 for the plasmid substrate used in our assays (Lockshon and Morris, 1983; Pruss et al., 1982), each DNA molecule should contain ~17 supercoils (Lko = 2927 bp plasmid ×1 turn/10.5 bp = 280 turns/plasmid; ΔLk = σ ×Lko = −0.06×280 turns/plasmid=~17 negative supercoils/plasmid). Based on our time-course data (Figure 1C), at a 1:1.4 enzyme:plasmid ratio it takes between 10 and 15 min (600–900 s) for topo VI to fully relax this substrate, indicating that one strand passage event (equivalent to the removal of two supercoils) occurs every 50–75 s (i.e.: one strand passage event/2 supercoils removed ×17 supercoils/plasmid×1.4 plasmids/enzyme/600 s = 0.02 events/sec or one event every 50 s). This rate agrees well with the maximal ATPase rates we observe (~3 ATP/min on supercoiled DNA, corresponding to one strand passage event every ~40 s for an enzyme without futile cycling). Both ATPase and supercoil relaxation rates for Mm topo VI are at least 50-fold slower than speeds reported for a broad range of bacterial and eukaryotic type IIA topoisomerases, which catalyze strand passage events at a rate of ~1–2 s−1 (Basu et al., 2012; Higgins et al., 1978; Lindsley and Wang, 1993; Osheroff et al., 1983; Sugino and Cozzarelli, 1980; Vos et al., 2013).

Because replication and chromosome segregation demarcate an absolute minimum requirement for topoisomerase activity in the cell, we next asked whether the genome size and doubling time of M. mazei under optimal growth conditions might allow for topo VI’s slow activity. M. mazei have a ~ 8–16 hr doubling time in optimal conditions (Mah, 1980), and possess a ~4 Mbp genome (Deppenmeier et al., 2002). However, the closely related Methanoscarina acetivorans maintains a chromosome copy number of ~16 in similar conditions (Hildenbrand et al., 2011), and the maintenance of moderate to high polyploidy appears to be a common trait in euryarchaea (Samson and Bell, 2011). Assuming M. mazei possess a similar genome copy number as M. acetivorans during growth, each doubling would require copying ~64 Mbp of DNA per cell. M. mazei chromosomes are circular; thus, assuming ~10 bp/turn of DNA, replicating 64 Mbp requires the removal of ~6.4 M DNA links per cell cycle. If doubling takes 10 hr, replication in M. mazei requires a minimum unlinking rate of ~200 s−1 (6.4 M DNA links/36,000 s = 178 DNA links/sec). By comparison, E. coli requires a similar unlinking rate to replicate and separate its single ~4 Mbp chromosome over a 40 min cell cycle (0.4M DNA links/2400 s = 167 links/sec) (Blattner et al., 1997; Schaechter, 1962). A similar minimum unlinking rate also holds for S. cerevisiae (12 Mbp genome/10 bp/turn of DNA/90 min doubling time yields ~220 links/sec (Goffeau et al., 1996; Sherman, 2002)). Thus, topo VI’s slow activity is not compensated for by slow replication or delayed chromosome segregation when compared to bacterial and eukaryotic counterparts.

Finally, we considered whether topo VI’s activity might reflect the set of encoded topoisomerases in M. mazei, which in addition to topo VI include topo III and a DNA gyrase gained from bacteria by horizontal gene transfer (Forterre et al., 2007). If M. mazei employs its assortment of topoisomerases similarly to other organisms, then M. mazei topo III would be expected to play a role in hemicatenane resolution (Mm topo III may provide a modest degree of negative supercoil relaxation as well, but generally this reaction is inefficient for this subfamily of type IA enzymes) (DiGate and Marians, 1988; Harmon et al., 1999; Wallis et al., 1989). M. mazei gyrase is expected to help remove positive supercoils and to further negatively-supercoil the chromosome (Charbonnier and Forterre, 1994; Drlica and Snyder, 1978; Gellert et al., 1976; Higgins et al., 1978). This division of labor suggests that Mm topo VI serves as the major decatenating factor of the cell and likely also assists in counterbalancing the build-up of negative supercoils arising from transcription and gyrase activity. This role is comparable to that of topo IV in E. coli (Zechiedrich and Cozzarelli, 1995). Because the unlinking rate required by replication does not solely fall upon cellular decatenase activity (removal of both positive supercoils ahead of the replication fork and hemicatenanes also contribute (Hiasa and Marians, 1996; Peter et al., 1998)), the cellular copy number of ~5–30 topo IV heterotetramers in E. coli (Taniguchi et al., 2010; Wiśniewski and Rakus, 2014) may estimate the workload required of topo VI in M. mazei. Such a workload would require ~1500 copies of topo VI (considering 30 copies of topo IV, a topo IV strand passage rate of 1 s−1, and a topo VI strand passage rate of 0.02 s−1) to keep up with replication. This calculation does not begin to account for any transcriptional demands that might be also placed on topo VI, which if compared to E. coli would likely need to compensate for the missing activity of E. coli topo I in M. mazei, and would push the necessary copy number even higher. Collectively, these considerations indicate that topo VI either needs to be present at extremely high concentrations to fulfill its expected role in vivo, or else be significantly stimulated by as yet unidentified factors.

References

  1. 1
  2. 2
  3. 3
  4. 4
  5. 5
  6. 6
  7. 7
  8. 8
  9. 9
  10. 10
  11. 11
  12. 12
    Purification of a DNA topoisomerase II from the hyperthermophilic archaeon Sulfolobus shibatae. A thermostable enzyme with both bacterial and eucaryal features
    1. A Bergerat
    2. D Gadelle
    3. P Forterre
    (1994)
    The Journal of Biological Chemistry 269:27663–27669.
  13. 13
  14. 14
  15. 15
  16. 16
  17. 17
  18. 18
  19. 19
  20. 20
  21. 21
  22. 22
  23. 23
    The genome of Methanosarcina mazei: evidence for lateral gene transfer between bacteria and archaea
    1. U Deppenmeier
    2. A Johann
    3. T Hartsch
    4. R Merkl
    5. RA Schmitz
    6. R Martinez-Arias
    7. A Henne
    8. A Wiezer
    9. S Bäumer
    10. C Jacobi
    11. H Brüggemann
    12. T Lienard
    13. A Christmann
    14. M Bömeke
    15. S Steckel
    16. A Bhattacharyya
    17. A Lykidis
    18. R Overbeek
    19. HP Klenk
    20. RP Gunsalus
    21. HJ Fritz
    22. G Gottschalk
    (2002)
    Journal of Molecular Microbiology and Biotechnology 4:453–461.
  24. 24
    Identification of a Potent Decatenating Enzyme from Escherichia coli
    1. RJ DiGate
    2. KJ Marians
    (1988)
    The Journal of Biological Chemistry 263:13366–13373.
  25. 25
  26. 26
  27. 27
  28. 28
  29. 29
  30. 30
  31. 31
  32. 32
  33. 33
    The Proteomics Protocols Handbook
    1. E Gasteiger
    2. C Hoogland
    3. A Gattiker
    4. S Duvaud
    5. MR Wilkins
    6. RD Appel
    7. A Bairoch
    (2005)
    571–608, Protein Identification and Analysis Tools on the ExPASy Server, The Proteomics Protocols Handbook, 10.1385/1-59259-890-0:571.
  34. 34
  35. 35
  36. 36
  37. 37
  38. 38
  39. 39
  40. 40
  41. 41
  42. 42
  43. 43
  44. 44
  45. 45
  46. 46
  47. 47
  48. 48
  49. 49
  50. 50
  51. 51
  52. 52
  53. 53
  54. 54
    On the coupling between ATP usage and DNA transport by yeast DNA topoisomerase II
    1. JE Lindsley
    2. JC Wang
    (1993)
    The Journal of Biological Chemistry 268:8096–8104.
  55. 55
  56. 56
  57. 57
  58. 58
  59. 59
  60. 60
  61. 61
  62. 62
  63. 63
  64. 64
  65. 65
    DNA Topoisomerase II from Drosophila melanogaster. Relaxation of supercoiled DNA
    1. N Osheroff
    2. ER Shelton
    3. DL Brutlag
    (1983)
    The Journal of Biological Chemistry 258:9536–9543.
  66. 66
  67. 67
  68. 68
  69. 69
  70. 70
  71. 71
  72. 72
  73. 73
  74. 74
  75. 75
  76. 76
  77. 77
  78. 78
  79. 79
  80. 80
    The PyMOL Molecular Graphics System
    1. Schrödinger, LLC
    (2012)
    The PyMOL Molecular Graphics System, 1.5.x.
  81. 81
  82. 82
  83. 83
  84. 84
  85. 85
  86. 86
  87. 87
  88. 88
  89. 89
    The Intrinsic ATPase of DNA Gyrase
    1. A Sugino
    2. NR Cozzarelli
    (1980)
    The Journal of Biological Chemistry 255:6299–6306.
  90. 90
  91. 91
    Characterization of the ATP binding site on Escherichia coli DNA gyrase. Affinity labeling of Lys-103 and Lys-110 of the B subunit by pyridoxal 5'-diphospho-5'-adenosine
    1. JK Tamura
    2. M Gellert
    (1990)
    The Journal of Biological Chemistry 265:21342–21349.
  92. 92
  93. 93
  94. 94
  95. 95
  96. 96
    Covalent Bonds between Protein and DNA-Formation of phosphotyrosine linkage between certain DNA topoisomerases and DNA
    1. YC Tse
    2. K Kirkegaard
    3. JC Wang
    (1980)
    The Journal of Biological Chemistry 255:5560–5565.
  97. 97
  98. 98
  99. 99
  100. 100
  101. 101
  102. 102
  103. 103
  104. 104
  105. 105
  106. 106
  107. 107
  108. 108
  109. 109
  110. 110
  111. 111
  112. 112
  113. 113
  114. 114

Decision letter

  1. Geeta J Narlikar
    Reviewing Editor; University of California, San Francisco, United States

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

Thank you for submitting your article "Topoisomerase VI senses and exploits both DNA crossings and bends to facilitate strand passage" for consideration by eLife. Your article has been reviewed by three peer reviewers, and the evaluation has been overseen by a Reviewing Editor and Andrea Musacchio as the Senior Editor. The following individuals involved in review of your submission have agreed to reveal their identity: Anthony Maxwell (Reviewer #2).

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

Summary of the work:

This paper describes mechanistic studies on a type IIB DNA topoisomerase (topo VI) from the archeon Methanosarcina mazei. This type of topoisomerase has some similarities to the better studied Type IIA topoisomerases, but significant differences in domain organization suggest there may be mechanistic difference between the IIA an IIB topoisomerase. The authors perform ATPase activity assays, FRET-based studies of dimerization of the N-terminal ATPase, direct and competition-based assays of topo IV binding to different types of DNA substrates (supercoiled DNA, a four-way junction or crossing, and linear DNAs of different lengths), and supercoil relaxing and processivity assays. A major part of this work is the identification of three regions of the enzyme with potential roles in DNA interaction and an exploration of these interactions using site-directed mutagenesis. Several mutants were made and tested in the same assays used with WT enzyme.

Based on these analyses, the authors conclude that topo VI senses and exploits both DNA crossings and bends to facilitate strand passage. Overall the manuscript provides new insights into how type IIB DNA topoisomerases tightly couple ATP hydrolysis and DNA cleavage to recognition of the appropriate DNA substrate. The results also have implications for understanding the mechanisms of paralogs of TopIIA such as Spo11, which helps initiate meiotic recombination.

Essential revisions:

Several issues should be clarified by the authors prior to publication.

1) Although WT topo VI binds the four-way junction (crossing or stacked junction) with a Kd of 122 nM, which is better than its binding to a 20 bp linear DNA (with a Kd of 427nM), the Kds of topo VI for 30 and 40 bp DNA are 84 and 49 nM, respectively, much better than that for the stacked junction (a total of 36 bp) (Figures 5 and 6). Why do the authors conclude that topo VI senses DNA crossing? Furthermore, does a four-way junction mimic a DNA crossing or catenate?

2) Is there any explanation for the decreased apparent affinity of topo VI for a 70 bp linear DNA compared to 40 or 60 bp DNA (Figure 1)? The minimal DNA length for topo VI binding appears to be 30 bp and the difference in Kds between 20 and 30 bp is five-fold. But why does increasing from 40 bp to 70 bp not increase DNA binding by topo VI? The increase in the Kd, app between 60 bp and 70 bp exceeds the uncertainties in the measurements. Can the authors comment on this decrease in affinity that occurs at about the length at which the model predicts that potentially both H2TH domains could interact with the duplex?

3) The ability of the H2TH region to facilitate bending a 70bp DNA substrate is strongly based on the FRET signal (shortened end-to-end distance of DNA) in the presence of H2TH but no FRET without H2TH. Is it possible that FRET arises from topo VI capturing two DNA molecules in the presence of H2TH?

4) The increased ATPase activity of the KGRR mutant compared to WT topo VI in the presence of supercoiled DNA and yet the lack of topoisomerase activity of the mutant (Figure 4) are not explained by its low affinity for supercoiled DNA and reduced ATPase dimerization, both of which are pre-requisites for ATPase activity. The supercoiled DNA-dependent ATPase domain-dimerization (FRET measurements) is initially high and decreases with time (Figure 7—figure supplement 1 and Figure 6), which is abnormal compared to the behavior of WT and the H2TH mutant topo VI, which increase with time (Figure 6B). The suggestion that the ATPase domain dissociates does not agree with the high ATPase activity, nor with the FRET data. At 150 min, the FRET signal due the ATPase domain dimerization of the KGRR mutant is at approximately the same level as the H2TH mutant topo VI and no worse. The Km of the KGRR mutant for ATP (~300µM) is much higher than WT (~3X) and H2TH topo VI (up to 10 X). KGRR is located in the ATPase domain. Could the mutation be influencing ATP binding? Do the authors know the KGRR mutant's affinity for AMPPNP (1 mM), which is used in the dimerization assay?

5) Is there any residual supercoiled DNA nicking activity of the KGRR mutant topo VI over time (Figure 4, and Figure 4—figure supplement 1)? Is there any explanation for the different "relaxed" circular DNAs in WT and three mutants? Is one a nicked band? The KGRR product is dominated by the second slowest migrating band, while the H2TH mutant predominantly produced the slowest migrating relaxed circular DNA. 10. It would be helpful if the authors could indicate linear and nicked bands on the gel for clarification. In a related point, it appears that there is an increase in the fraction of nicked or linear DNA in the gels testing the KGRR mutant in comparison to the other mutants or the wild-type enzyme. Is this significant and if so does it provide any additional insight into the regulation of cleavage by the KGRR motif?

6) It is not clear that sheared salmon sperm DNA is a suitable surrogate for linear DNA. As far as is known this preparation contains a mixture of DNAs, including single-stranded forms. Why didn't the authors just use linearized plasmid DNA in this experiment?

7) One difficult issue with the ATPase assays is the possible contribution of non-specific ATPases that may contaminate the topo VI prep. This difficulty can be addressed with other enzymes, e.g. DNA gyrase, where specific ATPase inhibitors are available. In this case the authors need to be cautious about the level of 'basal' ATPase activity, and this is particularly problematic with the mutants, where a different level of contaminating ATPase is possible. One possibility is to cut the topo VI bands out of a gel, re-fold and show that the ATPase activity is intrinsic to the enzyme. In this context it is important that the authors comment on the basal ATPase rates of the mutated enzymes.

8) Quoting kcat and KM values for this enzyme may not be straight forward (Figure 2—figure supplement 1). Type II topoisomerases are, in general, non-Michaelian enzymes, due to the possession of two ATPase active sites and the likely cooperativity that occurs. Although they can conform to M-M kinetics this is only 'apparent'. The ATPase data here are not really refined enough to distinguish Michaelian from non-Michaelian behaviour. However, as this is not essentially an enzyme kinetics study it does not matter too much. The authors could point this out in the text and add a footnote to the table in Figure 2—figure supplement 1 saying that these are likely to be apparent values.

9) It was surprising that the authors did not test DNA cleavage very much in this work, except in Figure 7C in the context of the mutants. Could a systematic study of cleavage couple with the binding studies have been more informative? In this context there are gels in the manuscript that show cleavage of DNA that is not commented upon, e.g. Figure 1D (large plasmid), Figure 4—figure supplement 1). Indeed the level of cleavage (linearization) is quite high in some tracks and seems to occur without ATP, which is not usually the case for topo VI. Is this a contaminant? The authors need to comment on this.

10) Subsection “Three conserved elements in Top6B play a role in DNA binding, the sensing of DNA geometry, 2 and the productive coupling of ATP hydrolysis to strand passage”, last paragraph:

The authors did the competitive binding experiments (Figure 5B) only with H2THEEE and KGRREEE but not with H2THAAA or KGRRAAA. Would it be possible to include the competitive binding experiments with H2THAAA and KGRRAAA? The authors tested these two mutants with the short-stacked junction discussed later (Figure 6B) and it would be more informative to compare the supercoil binding to these junction data for all four mutants.

11) "Together, these data indicate that both the KGRR loop and H2TH domain contribute to the preferential binding to supercoiled DNA, but that neither is solely responsible for this discrimination."

If these two domain mutations only affected supercoil DNA binding, then it could be argued that these two regions are important for supercoil DNA sensing. However, the linear DNA binding is also adversely affected for both mutations (KGRRAAA/EEE and H2THAAA/EEE) particularly for longer DNA (>50bp) (Figure 5—figure supplement 1). This adverse effect can be explained for H2TH since it can stabilize binding of longer DNA, but it is puzzling why KGRRAAA shows lower binding affinity for longer DNA. In addition, if a positive patch is important for DNA interaction stabilized through electrostatic interactions, it is surprising that H2THEEE exhibited a lower Kd than that of H2THAAA.

12) Subsection “The KGRR loop acts as a DNA crossing sensor to promote Top6B dimerization”, last paragraph:

In Figure 6—figure supplement 2, the authors compare the emission spectra of wild type, KGRRAAA and H2THAAA with and without supercoiled DNA to probe the extent of Top6B dimerization. Interestingly, the spectra of the apo-enzymes KGRR and H2TH differ from that of the WT enzyme. For KGRR, the emission at 670 nm is higher than that of wild type while that of H2TH is lower. Is it possible that the site-specific mutation can lead to change in protein conformation? For example, two Top6B are closer or further apart for KGRR and H2TH respectively. In line with the previous comment, is it possible that this conformational change in KGRR could result in reduction in binding and or sensing of both linear DNA (G-segment) as well as the crossing DNA (T-segment)?

13) Subsection “The H2TH interface engages an extended G-segment to couple nucleotide-dependent Top6B dimerization with DNA cleavage”, second paragraph:

Regarding Figure 7: Based on the image shown in Figure 7C for the wild type case, the amounts of DNA loaded on the gel for different length of DNA appears different: the 70 bp duplex lanes appear darker than those for 60 bp, suggesting more of the 70 bp product was loaded. Is this just image artifact? Can the authors quantify the relative cleavage relative to the total in each lane? This is an important point since the argument for a critical ~70 bp length requirement for cleavage hinges on this data. Do the authors think that both H2TH should interact with DNA in order to achieve DNA cleavage or would interaction with H2TH on one side suffice? In addition, considering the fact that 20-30-40 oligo bands are well separated on the ssDNA ladder lane, the cleavage product would be expected to be located somewhere between 30-40 nt (mixture of 37 and 33 due to 2-nt stagger cut) but they appear to close to 30 nt. Do the authors have any explanation? Some of the lanes show high molecular weight bands? Are they intact duplex substrate?

14) "The inability of a 40bp duplex to support cleavage, even though this DNA binds with higher affinity than a 20bp duplex and is long enough to reach both Stalk/WKxY regions, suggests G-segment DNAs must engage both H2TH regions before strand scission can be triggered."

The authors' argument is not fully convincing. As 40bp DNA contains a preferential cleavage site in the middle (Figure 1—figure supplement 1), even if Topo VI can bind one end of H2TH as shown in Figure 7B (fifth from top configuration), the cleavage product may be too low to detect due to extremely low cleavage efficiency. Thus, the 40bp cleavage data is not sufficient to draw the conclusion of two H2TH requirement. In addition, the 60bp substrate cleavage data may suggest one-H2TH interaction is sufficient.

15) Considering the complexity of this manuscript, it is recommended the authors make things clearer regarding the conditions of experiments presented in the different sections – such as ATP, AMPPNP, DNA, enzyme concentrations and if they are all same for the same type of experiments, state as such at the beginning. Otherwise a table listing the conditions of the various measurements in one place would be helpful. It would also be useful to know how the authors decided on the reaction conditions for the topo VI assays, and how these compare with the in vivo environment of M. mazei. In addition it is confusing that the authors use the bp scale to convolve the length and concentration of DNA instead of indicating the length of DNA and its concentration separately for supercoiled DNA. In line with this, "800bp supercoiled DNA:enzyme" in Figure 2A can be confused as 800bp length of supercoiled DNA rather than bp scaled concentration. Whereas the rational for the scaling of the amount of DNA in bp is understandable, it would be helpful to provide the length and concentration in addition to this relative measure. It would further be useful to comment on the choice of oligos used in these experiments (Figure 1—figure supplement 1); these are based on a preferred cleavage sequence for the sulfolobus enzyme. Is there evidence that the M. mazei enzyme has a similar sequence preference?

https://doi.org/10.7554/eLife.31724.060

Author response

Essential revisions:

Several issues should be clarified by the authors prior to publication.

1) Although WT topo VI binds the four-way junction (crossing or stacked junction) with a Kd of 122 nM, which is better than its binding to a 20 bp linear DNA (with a Kd of 427nM), the Kds of topo VI for 30 and 40 bp DNA are 84 and 49 nM, respectively, much better than that for the stacked junction (a total of 36 bp) (Figures 5 and 6). Why do the authors conclude that topo VI senses DNA crossing? Furthermore, does a four-way junction mimic a DNA crossing or catenate?

The four-way junction serves as a mimic for the crossings present in plectonemic supercoiled DNA and catenated DNA. A four-way junction will fold into a stacked-X structure at the divalent cation concentrations used in our binding experiments (see (Duckett et al., 1988; Eichman, Vargason, Mooers, and Ho, 2000; McKinney, Declais, Lilley, and Ha, 2003; Ortiz-Lombardía, González, Aymamí, Azorín, and Coll, 1999). We noticed the geometry taken on by a prospective T-segment and G-segment DNA when modeled into topo VI closely mimics this stacked-X structure. We have added two panels to Figure 6 to illustrate how a stacked junction mimics this intermediate (A and B), and have revised the text in the first paragraph of the subsection “The KGRR loop acts as a DNA crossing sensor to regulatepromote Top6B dimerization” to reference these panels.

The increased affinity of topo VI for a 30 bp or 40 bp duplex may be attributed to these DNAs engaging an extended G-segment binding interface (e.g., through the Stalk/WKxY region). The Holliday junction substrate folds as a 16 bp duplex stacked on a 20 bp duplex, and should bind no better than a simple 20 bp duplex if there were no preference for topo VI to bind to a crossing. However, the affinity does increase for the junction, indicating that topo VI favorably recognizes two DNA segments. It is from the relative increase in wildtype affinity for the stacked junction, as well as the relative decrease in affinity for this substrate with the KGRR mutants, that we conclude topo VI recognizes prospective T-segment/G-segment crossings.

2) Is there any explanation for the decreased apparent affinity of topo VI for a 70 bp linear DNA compared to 40 or 60 bp DNA (Figure 1)? The minimal DNA length for topo VI binding appears to be 30 bp and the difference in Kds between 20 and 30 bp is five-fold. But why does increasing from 40 bp to 70 bp not increase DNA binding by topo VI? The increase in the Kd, app between 60 bp and 70 bp exceeds the uncertainties in the measurements. Can the authors comment on this decrease in affinity that occurs at about the length at which the model predicts that potentially both H2TH domains could interact with the duplex?

Two lines of evidence indicate that the slightly higher Kd,app of topo VI for a 70 bp duplex compared to a 40 or 60 bp duplex is an effect of 5’ end-labeled fluorophore position on that particular probe DNA. First, the Kd,app determined by fluorescence anisotropy for an 80 bp duplex is also weaker (~2x) than the Kd,app determined for a 60bp duplex (see Author response image 1A). However, in a competitive binding experiment, the 70 bp and 80 bp duplex compete progressively better against a fluorescently labeled probe than the 60 bp duplex (see Author response image 1B). We have not included these findings with the revised manuscript, but we are happy to do so as supplements to Figure 1 if the reviewers deem it necessary.

3) The ability of the H2TH region to facilitate bending a 70bp DNA substrate is strongly based on the FRET signal (shortened end-to-end distance of DNA) in the presence of H2TH but no FRET without H2TH. Is it possible that FRET arises from topo VI capturing two DNA molecules in the presence of H2TH?

The DNA bending experiments were carried out at a 2:1 molar excess of enzyme to duplex DNA. Thus, the likelihood of two 70 bp duplexes binding to the same enzyme to induce a FRET response is small. However, to more explicitly rule out the potential capture of two duplexes as a cause for changes in FRET, we have repeated the DNA bending experiment using an equimolar mixture of two 70 bp duplexes, one labeled with Cy5 and the other with Cy5.5. In this setup, any observed FRET increase must occur between two duplexes, rather than by bending of a single DNA. We observe no nucleotide-dependent increase in FRET with this experimental setup, allowing us to rule out duplex capture as the source of the FRET signal. This control has been added as Figure 7—figure supplement 1 and is referenced in the last paragraph of the subsection “The H2TH interface engages an extended G-segment to couple nucleotide-dependent Top6B dimerization with DNA cleavage.”.

4) The increased ATPase activity of the KGRR mutant compared to WT topo VI in the presence of supercoiled DNA and yet the lack of topoisomerase activity of the mutant (Figure 4) are not explained by its low affinity for supercoiled DNA and reduced ATPase dimerization, both of which are pre-requisites for ATPase activity. The supercoiled DNA-dependent ATPase domain-dimerization (FRET measurements) is initially high and decreases with time (Figure 7—figure supplement 1 and Figure 6), which is abnormal compared to the behavior of WT and the H2TH mutant topo VI, which increase with time (Figure 6B). The suggestion that the ATPase domain dissociates does not agree with the high ATPase activity, nor with the FRET data. At 150 min, the FRET signal due the ATPase domain dimerization of the KGRR mutant is at approximately the same level as the H2TH mutant topo VI and no worse. The Km of the KGRR mutant for ATP (~300µM) is much higher than WT (~3X) and H2TH topo VI (up to 10 X). KGRR is located in the ATPase domain. Could the mutation be influencing ATP binding? Do the authors know the KGRR mutant's affinity for AMPPNP (1 mM), which is used in the dimerization assay?

The increased maximal ATPase rates of the KGRR mutants in the presence of supercoiled DNA can be explained if the release of hydrolysis products by these mutants is premature. Wildtype topo VI takes ~40 s to complete each ATPase cycle. The KGRR-AAA mutant takes ~15 s, but fails to perform strand passage. This difference suggests that under normal circumstances, a delay in product release is an important component of the topo VI strand passage mechanism. In this view, the high ATPase rates of the KGRR mutants do not reflect a ‘better’ ATPase activity, but rather an uncoupling of the ATPase cycle (and specifically, product release) from a set of slow conformational changes that are necessary for strand passage.

The experiments in Figure 4B do not measure the Km,app of each topo VI mutant for ATP. Rather, as noted in the manuscript and Figure 4—figure supplement 2, these experiments look at the stimulation of ATP hydrolysis as a function of DNA concentration at a single (2 mM) ATP concentration. We apologize for any confusion our discussion of this experiment might have caused, and we have modified the parameter names to Kstim,DNA and kcat-stim,DNA. The weaker Kstim,DNA of the KGRR mutants agrees with the lower affinity of these mutants for supercoiled DNA.

To address whether the KGRR mutations affect ATP engagement, we have measured ATP hydrolysis by KGRRAAA in the presence of supercoiled DNA (400 μM basepairs DNA) as a function of ATP concentration (see below). This is analogous to the experiment with wildtype topo VI in Figure 2A. The Km,app for ATP is ~1.5x higher for the KGRR-AAA mutant (350 μM) than for wildtype topo VI (240 μM). This result indicates that there is a slight decrease in affinity of the KGRR-AAA mutant for ATP, but also suggests that 1 mM AMPPNP is still an appropriate concentration to use for the dimerization assay.

In the dimerization assay, the KGRRAAA construct responds to AMPPNP in the presence of a 70 bp duplex, however, this response does not exceed the FRET signal generated by supercoiled DNA alone (Figure 7—figure supplement 1). While the KGRRAAA construct does not respond to AMPPNP in the presence of supercoiled DNA, it does show an increased FRET signal with supercoiled DNA alone that is innately higher than wildtype topo VI or the H2TH-AAA mutant (Figure 6—figure supplement 2). These data, together with the ATPase results, suggest that upon binding supercoiled DNA, the KGRR-AAA mutant enzymes ‘pre-dimerize’ into a state competent for binding and hydrolyzing ATP (and that the intact KGRR loop of wild-type topo VI prevents this conformational change from occurring until ATP also binds). If the KGRR-AAA mutant inappropriately releases hydrolysis products before strand passage (perhaps because a bound T-segment can no longer suppress the release of these products), the enzyme will remain pre-dimerized. Interestingly, Klostermeier and colleagues have noted that ATPase pre-dimerization occurs for bacterial gyrase upon DNA binding, albeit in the wild-type enzyme (Gubaev and Klostermeier, 2011).

Finally, as noted by the reviewers, both H2TH-AAA and KGRR-AAA produce similar FRET signals in the presence of supercoiled DNA and AMPPNP that are lower than the wildtype signal in the same condition. Given both mutants are impaired for strand passage, this raises the possibility that the higher wildtype signal reflects a conformational response to nucleotide binding that is not fully accessible to either mutant. Such a model directly relates to comments raised by the reviewers in point 12 (below). We have revised the text in the last paragraph of the subsection “The KGRR loop acts as a DNA crossing sensor to regulate Top6B dimerization” to convey this interpretation more clearly.

5) Is there any residual supercoiled DNA nicking activity of the KGRR mutant topo VI over time (Figure 4, and Figure 4—figure supplement 1)? Is there any explanation for the different "relaxed" circular DNAs in WT and three mutants? Is one a nicked band? The KGRR product is dominated by the second slowest migrating band, while the H2TH mutant predominantly produced the slowest migrating relaxed circular DNA. 10. It would be helpful if the authors could indicate linear and nicked bands on the gel for clarification. In a related point, it appears that there is an increase in the fraction of nicked or linear DNA in the gels testing the KGRR mutant in comparison to the other mutants or the wild-type enzyme. Is this significant and if so does it provide any additional insight into the regulation of cleavage by the KGRR motif?

We apologize for any confusion over the identity of DNA species in Figure 4A and Figure 4—figure supplement 1. We have more clearly annotated the position of linear and nicked plasmid DNA species for these experiments. The slowest-migrating band seen for a number of mutants in Figure 4A is open-circle/nicked DNA present in the plasmid preparation. Of all the constructs, only wildtype topo VI produces a fully relaxed, closed-circle topoisomer distribution.

We do not believe excessive nicking to be a feature of the KGRR mutants. In Figure 4A, there is no dose-dependent nicking for KGRR-EEE. Any apparent dose-dependent nicking for KGRR-AAA is an artifact of gel illumination due to our old scanner; a replicated gel on a newer system with more uniform illumination displays no dose-dependent nicking (see Author response image 3).

In Figure 4—figure supplement 1, all experiments show a slowly-migrating band that increases with time. This is the open-circle/nicked DNA species. The reviewers note an additional slower-running band in the KGRR time-courses. This is contaminating plasmid concatemer in the specific plasmid preparation used in those experiments. We have replicated the time-course gels for both KGRR mutants (see Author response image 4), which verify the band that increases over time is the same nicked species that increases in the Stalk/WKxY and H2TH mutant experiments.

The KGRR timecourse replicates also show that increased nicking over time is both nucleotide and enzyme-independent. This increase may reflect either a contaminant in an assay component, or a chemical nicking activity of our 1x assay conditions over the 2 h incubation at 30 oC. We have explicitly commented on this point in the manuscript (subsection “Three conserved elements in Top6B play a role in DNA binding, the sensing of DNA geometry, and the productive coupling of ATP hydrolysis to strand passage”, fourth paragraph).

6) It is not clear that sheared salmon sperm DNA is a suitable surrogate for linear DNA. As far as is known this preparation contains a mixture of DNAs, including single-stranded forms. Why didn't the authors just use linearized plasmid DNA in this experiment?

Other groups in the topoisomerase field have found sheared salmon-sperm DNA to be a suitable general dsDNA substrate (e.g., Wang, Lindsley, Nitiss and Austin groups (Baird, Gordon, Andrenyak, Marecek, and Lindsley, 2001; Harkins and Lindsley, 1998; Morris, Harkins, Tennyson, and Lindsley, 1999; Olland and Wang, 1999; Vaughn et al., 2005; Walker et al., 2004; West and Austin, 1999)). Following this precedent, we have frequently used sheared salmon-sperm DNA in previous studies and found no issue with the substrate (e.g., (Schmidt, Osheroff, and Berger, 2012; Schoeffler, May, and Berger, 2010; Tretter and Berger, 2012)). Based on agarose gel electrophoresis, the sheared salmon-sperm DNA used in this study consists of DNAs ranging from ~75-500 basepairs in length (primarily ~75-100 basepairs, see Author response image 5A). This DNA mixture competes for DNA binding to topo VI similarly to a defined DNA substrate in the same length range (60 bp), whereas single stranded DNA competes quite poorly (see Author response image 5B). According to the manufacturer, the sheared salmon-sperm DNA has not been boiled and should therefore be double-stranded. We confirmed that our sheared salmon-sperm DNA preparations are resistant to S1 single-stranded DNA nuclease treatment, but that when boiled, it is degraded, confirming the manufacturer’s claims (see Author response image 5C). While we do not believe these controls would add to the manuscript, we are happy to include them as supplements to Figure 1 if the reviewers deem it necessary.

The DNAs in the sheared salmon-sperm sample are shorter than the persistence length of DNA and thus less likely to form bends or crossings. This characteristic makes sheared salmon-sperm DNA a good substrate to compare against supercoiled plasmid DNA, which would have such features. Because linearized plasmid DNA (~3 kb) is sufficiently long to form random bends and intramolecular crossings, we expect it to have an intermediate effect upon topo VI compared to sheared salmon sperm DNA. Two tests confirm this prediction. In the first, we have found that linearized plasmid competes better than sheared salmon-sperm DNA for topo VI binding, nearly as well as supercoiled DNA (see Author response image 6D). This result comports with our model in that topo VI preferentially binds better to crossovers than single duplexes; in the absence of ATP (which is omitted in the binding assays), the crossovers present in the supercoiled and linearized plasmids will not be resolved, and will act as a sink to pull topo VI (which is present in limiting concentrations) away from the labeled 70 bp duplex. In the second test, we found that linearized plasmid has an overall stimulatory effect on ATP hydrolysis that is intermediate compared to that measured for sheared salmon-sperm DNA and supercoiled DNA (see Author response image 6E). This result is interesting, because at low supercoiled plasmid:enzyme ratios, where the crossover density is low (~1 crossover/enzyme), the ATPase stimulation is similar to that of linearized plasmid. However, at higher supercoiled plasmid:enzyme ratios, where the crossover density increases, the ATPase rate improves concordantly. Overall, these data are consistent with the conclusion that topo VI senses and responds to crossings present in DNA substrates. We also have not added these data to the text, but can do so if the reviewers feel it necessary.

7) One difficult issue with the ATPase assays is the possible contribution of non-specific ATPases that may contaminate the topo VI prep. This difficulty can be addressed with other enzymes, e.g. DNA gyrase, where specific ATPase inhibitors are available. In this case the authors need to be cautious about the level of 'basal' ATPase activity, and this is particularly problematic with the mutants, where a different level of contaminating ATPase is possible. One possibility is to cut the topo VI bands out of a gel, re-fold and show that the ATPase activity is intrinsic to the enzyme. In this context it is important that the authors comment on the basal ATPase rates of the mutated enzymes.

We appreciate the potential for contamination by non-specific ATPases may contribute to the overall ATPase rates reported. We have added a supplemental figure (Figure 4—figure supplement 2) reporting the ‘basal’ ATPase rates for all mutants as a function of ATP concentration (see Figure 4—figure supplement 2). All mutants exhibit basal ATPase rates similar to each other and to wild-type topo VI.

To determine whether co-contaminants in our topo VI preparations contribute to measured ATP hydrolysis rates, we cloned, expressed, and purified a construct of topo VI that lacks a catalytic glutamate necessary for ATPase activity in GHKL ATPases (Top6BE44A). As expected, this mutant shows no supercoil relaxation activity (see Figure 2—figure supplement 1B,top). While there is some background ATPase activity in the Top6ABE44A prep, it is not DNA stimulated (see Figure 2—figure supplement 1B, bottom). Thus, the basal rates reported for topo VI do appear to contain a contribution from a non-specific ATPase contaminant. By contrast, the ability of DNA to stimulate (or fail to stimulate) the ATPase activity of our topo VI mutants is directly attributable to the specific mutations we have introduced. To more accurately represent the topo VI-specific ATPase activity reported in Figure 2, we have subtracted out the background ‘basal’ ATPase rate measured for Top6BE44A, and reprocessed the kinetic parameters reported in Figure 2—figure supplement 1. This adjustment is not necessary for the experiments reported in Figure 4B, as they measure stimulation above the basal rate. We have added the experiments in panel A and B as a supplement to Figure 2 and 4. We have also added callouts to each supplement in the first paragraph of the subsection “Topo VI actively uses DNA crossings to couple ATP hydrolysis with DNA strand passage” and in “Three conserved elements in Top6B play a role in DNA binding, the sensing of DNA geometry, and the productive coupling of ATP hydrolysis to strand passage”.

8) Quoting kcat and KM values for this enzyme may not be straight forward (Figure 2—figure supplement 1). Type II topoisomerases are, in general, non-Michaelian enzymes, due to the possession of two ATPase active sites and the likely cooperativity that occurs. Although they can conform to M-M kinetics this is only 'apparent'. The ATPase data here are not really refined enough to distinguish Michaelian from non-Michaelian behaviour. However, as this is not essentially an enzyme kinetics study it does not matter too much. The authors could point this out in the text and add a footnote to the table in Figure 2—figure supplement 1 saying that these are likely to be apparent values.

We agree that ATP hydrolysis by topo VI is likely cooperative, but the reported data do not sufficiently assess turnover at low ATP concentrations to distinguish between a cooperative model and ‘apparent’ Michaelis-Menten behavior. We have modified both the text (subsection “Topo VI actively uses DNA crossings to couple ATP hydrolysis with DNA strand passage”, first paragraph) and Figure 2—figure supplement 1 to explicitly state that the reported kinetic parameters are apparent, and are not meant to imply that the enzyme acts in a truly Michaelian regime.

9) It was surprising that the authors did not test DNA cleavage very much in this work, except in Figure 7C in the context of the mutants. Could a systematic study of cleavage couple with the binding studies have been more informative? In this context there are gels in the manuscript that show cleavage of DNA that is not commented upon, e.g. Figure 1D (large plasmid), Figure 4—figure supplement 1). Indeed the level of cleavage (linearization) is quite high in some tracks and seems to occur without ATP, which is not usually the case for topo VI. Is this a contaminant? The authors need to comment on this.

Our goal in this work was to determine how topo VI engages DNA and how DNA binding may coordinate activities related to strand passage. The experiments performed in Figure 7C directly contribute to resolving these questions by determining both the DNA binding elements and minimal DNA length required for topo VI-mediated DNA cleavage. Given all the mutants under investigation, a truly systematic study of cleavage activities would greatly enlarge this study: DNA topology, oligo lengths, and ATP dependence would all need to be assessed for seven different constructs (eight when including our new ATPase-defective mutant to look at the role of hydrolysis). Then there is also the question of preferred cleavage sequences, both within the central cleavage region and more peripherally toward the H2TH domains. We agree that it would be useful to more systematically investigate DNA cleavage by topo VI, but as part of a future effort.

Regarding the DNA gels noted by the reviewers – in Figure 1D, topo VI produces a relaxed distribution for the large plasmid (13.5 kb) that runs very close to the nicked plasmid species. Since it is difficult to unequivocally distinguish this distribution from open-circle (nicked) plasmid, we repeated the experiment using a smaller chase plasmid (6.5 kb). There is better separation between relaxed and nicked bands for the 6.5 kb chase plasmid, which verifies topo VI does not produce a level of nicking or linearization beyond the 20 min control lacking ATP. We have substituted this experiment for Figure 1D and Figure 1—figure supplement 1.

Regarding the increase in open-circle/nicked plasmid DNA for gels in Figure 4—figure supplement 1, we have addressed this in the response to point 5, above.

10) Subsection “Three conserved elements in Top6B play a role in DNA binding, the sensing of DNA geometry, 2 and the productive coupling of ATP hydrolysis to strand passage”, last paragraph:

The authors did the competitive binding experiments (Figure 5B) only with H2THEEE and KGRREEE but not with H2THAAA or KGRRAAA. Would it be possible to include the competitive binding experiments with H2THAAA and KGRRAAA? The authors tested these two mutants with the short-stacked junction discussed later (Figure 6B) and it would be more informative to compare the supercoil binding to these junction data for all four mutants.

We have added competition assays with the H2TH-AAA and KGRR-AAA mutants to Figure 5 as requested. The KGRR-AAA mutant shows intermediate effects between wildtype and the KGRR-EEE mutant. The H2TH-AAA mutant shows minimal impairment for both sheared salmon-sperm DNA and supercoiled DNA binding when compared to the H2TH-EEE mutant. We have revised the text in the subsection “Three conserved elements in Top6B play a role in DNA binding, the sensing of DNA geometry, and the productive coupling of ATP hydrolysis to strand passage”.

11) "Together, these data indicate that both the KGRR loop and H2TH domain contribute to the preferential binding to supercoiled DNA, but that neither is solely responsible for this discrimination."

If these two domain mutations only affected supercoil DNA binding, then it could be argued that these two regions are important for supercoil DNA sensing. However, the linear DNA binding is also adversely affected for both mutations (KGRRAAA/EEE and H2THAAA/EEE) particularly for longer DNA (>50bp) (Figure 5—figure supplement 1). This adverse effect can be explained for H2TH since it can stabilize binding of longer DNA, but it is puzzling why KGRRAAA shows lower binding affinity for longer DNA. In addition, if a positive patch is important for DNA interaction stabilized through electrostatic interactions, it is surprising that H2THEEE exhibited a lower Kd than that of H2THAAA.

The ability of a site to sense supercoiled DNA does not necessarily preclude it from also impacting the binding of linear DNAs. As pointed out, the impact of the H2TH domain on linear DNA binding can be explained by the interaction of this element with longer DNA segments. As for the KGRR mutants, it may be that these substitutions affect the conformational disposition of subunits in the tetramer, which in turn makes it more difficult for DNA bound in the active site of a Top6A dimer to access the flanking Top6B WKxY and H2TH regions. The discussion outlined for points 4 and 12 comports with this reasoning.

Regarding the Kd values of the 60 and 70 bp DNAs for the H2TH mutants, these point mutations lie at the outer edge of the G-segment binding tract and may therefore differentially affect either DNA binding register (discussed further in point 13) and/or the mobility of the fluorophore label. To control for such effects, we have measured the affinity of both H2TH-AAA and H2TH-EEE for unlabeled 60 or 70 bp duplexes in a competition experiment that uses the fluorescently labeled 70 bp duplex as a probe. As can be seen from the data, the H2TH-EEE mutant does not exhibit greater affinity for either DNA compared to the H2TH-AAA mutant. We have added these experiments as a supplement to Figure 5.

Interestingly, whereas a 70 bp duplex competes better than a 60 bp duplex for binding to wildtype topo VI (see point 2), competitive binding of the 70 bp duplex is relatively impaired for both H2TH mutants. As we now note in the revised manuscript, this result suggests that each set of substitutions is sufficient to ablate DNA-H2TH interactions, but that they minimally affect DNA engagement by Top6A or the Stalk/WKxY region. We also note that in the competitive binding experiments reported in the new Figure 5, the H2TH-AAA mutant binds supercoiled DNA more strongly than the H2TH-EEE mutant. This finding provides additional evidence that the H2TH site engages DNA primarily in a role as a bend sensor. These considerations are explicitly discussed in the subsection “Three conserved elements in Top6B play a role in DNA binding, the sensing of DNA geometry, and the productive coupling of ATP hydrolysis to strand passage”.

12) Subsection “The KGRR loop acts as a DNA crossing sensor to promote Top6B dimerization”, last paragraph:

In Figure 6—figure supplement 2, the authors compare the emission spectra of wild type, KGRRAAA and H2THAAA with and without supercoiled DNA to probe the extent of Top6B dimerization. Interestingly, the spectra of the apo-enzymes KGRR and H2TH differ from that of the WT enzyme. For KGRR, the emission at 670 nm is higher than that of wild type while that of H2TH is lower. Is it possible that the site-specific mutation can lead to change in protein conformation? For example, two Top6B are closer or further apart for KGRR and H2TH respectively. In line with the previous comment, is it possible that this conformational change in KGRR could result in reduction in binding and or sensing of both linear DNA (G-segment) as well as the crossing DNA (T-segment)?

Based on the FRET spectra of the mutant enzymes alone, we agree that these point mutations could alter the underlying conformational states sampled by the enzyme, and that any such behavior might well contribute to the effect of each set of mutations on DNA binding (a concept also noted in response to points 4 and 11, above). We have revised the manuscript (subsection “The KGRR loop acts as a DNA crossing sensor to regulate Top6B dimerization”) to more explicitly comment on this possibility.

13) Subsection “The H2TH interface engages an extended G-segment to couple nucleotide-dependent Top6B dimerization with DNA cleavage”, second paragraph:

Regarding Figure 7: Based on the image shown in Figure 7C for the wild type case, the amounts of DNA loaded on the gel for different length of DNA appears different: the 70 bp duplex lanes appear darker than those for 60 bp, suggesting more of the 70 bp product was loaded. Is this just image artifact? Can the authors quantify the relative cleavage relative to the total in each lane? This is an important point since the argument for a critical ~70 bp length requirement for cleavage hinges on this data. Do the authors think that both H2TH should interact with DNA in order to achieve DNA cleavage or would interaction with H2TH on one side suffice? In addition, considering the fact that 20-30-40 oligo bands are well separated on the ssDNA ladder lane, the cleavage product would be expected to be located somewhere between 30-40 nt (mixture of 37 and 33 due to 2-nt stagger cut) but they appear to close to 30 nt. Do the authors have any explanation? Some of the lanes show high molecular weight bands? Are they intact duplex substrate?

The apparent loading difference is an artifact of our previous gel scanner. A repeat of the experiment shows that loading is constant from lane to lane and that the cleavage product is still more pronounced for the 70bp oligo. We have also determined that the high molecular weight band in some lanes in Figure 7C (e.g., lanes containing wildtype topo VI and 70 bp DNA substrate) represent DNA bound to incompletely digested topo VI. These bands disappear with longer Proteinase K treatment. We have substituted these gels and quantified the% cleavage product above each lane where this product is distinguishable. Full digestion reveals that the wildtype enzyme cleaves the 70 bp DNA at a slightly higher level than either KGRR mutant. This result is concordant with the relative affinities of each construct for the 70 bp DNA. We have modified the text (subsection “The H2TH interface engages an extended G-segment to couple nucleotide-dependent Top6B dimerization with DNA cleavage”, third paragraph) to reflect these quantification data.

We agree that the cleavage bands are running at a position that would suggest topo VI is cutting off-center. Our oligo sequence is based on an optimal cleavage site identified by the Forterre group for S. shibatae topo VI (Buhler, Lebbink, Bocs, Ladenstein, and Forterre, 2001). The same group also identified a weaker cleavage site ~6 bp away from this locus; we suspect our M. mazei enzyme is cutting at this secondary site. We also agree that the data are consistent with the idea that the engagement of a single H2TH domain can be sufficient to promote cleavage. We have revised the text to reflect both points in the subsection “The H2TH interface engages an extended G-segment to couple nucleotide-dependent Top6B dimerization with DNA cleavage”.

14) "The inability of a 40bp duplex to support cleavage, even though this DNA binds with higher affinity than a 20bp duplex and is long enough to reach both Stalk/WKxY regions, suggests G-segment DNAs must engage both H2TH regions before strand scission can be triggered."

The authors' argument is not fully convincing. As 40bp DNA contains a preferential cleavage site in the middle (Figure 1—figure supplement 1), even if Topo VI can bind one end of H2TH as shown in Figure 7B (fifth from top configuration), the cleavage product may be too low to detect due to extremely low cleavage efficiency. Thus, the 40bp cleavage data is not sufficient to draw the conclusion of two H2TH requirement. In addition, the 60bp substrate cleavage data may suggest one-H2TH interaction is sufficient.

We agree with this assessment of our data, and we have revised the text (subsection “The H2TH interface engages an extended G-segment to couple nucleotide-dependent Top6B dimerization with DNA cleavage”) accordingly.

15) Considering the complexity of this manuscript, it is recommended the authors make things clearer regarding the conditions of experiments presented in the different sections – such as ATP, AMPPNP, DNA, enzyme concentrations and if they are all same for the same type of experiments, state as such at the beginning. Otherwise a table listing the conditions of the various measurements in one place would be helpful. It would also be useful to know how the authors decided on the reaction conditions for the topo VI assays, and how these compare with the in vivo environment of M. mazei. In addition it is confusing that the authors use the bp scale to convolve the length and concentration of DNA instead of indicating the length of DNA and its concentration separately for supercoiled DNA. In line with this, "800bp supercoiled DNA:enzyme" in Figure 2A can be confused as 800bp length of supercoiled DNA rather than bp scaled concentration. Whereas the rational for the scaling of the amount of DNA in bp is understandable, it would be helpful to provide the length and concentration in addition to this relative measure. It would further be useful to comment on the choice of oligos used in these experiments (Figure 1—figure supplement 1); these are based on a preferred cleavage sequence for the sulfolobus enzyme. Is there evidence that the M. mazei enzyme has a similar sequence preference?

The reaction conditions (salts, buffers, etc.) for each type of experiment are the same throughout the manuscript. The only exception is between the ATP hydrolysis experiments in Figure 2 (where DNA concentration is maintained and ATP concentration is varied) and the ATP hydrolysis experiments in Figure 4B (where ATP concentration is maintained and DNA concentration is varied). As requested, we have added a table listing the enzyme concentration, nucleotide concentration and identity (i.e. ATP vs. AMPPNP), and DNA concentration for each set of experiments to aid the reader.

Assay conditions for M. mazei topo VI were previously optimized by screening monovalent salt identity, salt concentration, and buffer identity from a panel of common biochemical reagents (Corbett, Benedetti, and Berger, 2007). The current study uses essentially the same conditions, except for the addition of 10% (v/v) glycerol and 0.1 mg/mL BSA, which we found improved M. mazei topo VI activity at low enzyme concentrations. Our in vitro conditions are comparable to the optimal growth conditions and known physiology of M. mazei, including temperature (30 oC to 40 oC), pH (pH 6-8), and salinity (M. mazei mayharbor potassium glutamate concentrations up to 400 mM) (Deppenmeier et al., 2002; Mah, 1980; Spanheimer and Müller, 2008).

A 2.9 kb plasmid, pSG483, is used throughout the text as a supercoiled substrate. We have added this size identification to figures and text where it is missing. In cases where multiple DNA substrates are being used (i.e., ATPase assays, competition assays and gate closure assays), we have used a μM basepair DNA concentration scale. We believe this is the most straightforward scale for comparing between DNA substrates and DNA-dependent characteristics of topo VI.

The short duplexes used in this study (Figure 1—figure supplement 1) are based on a preferred sequence for S. shibatae topo VI. We have not explicitly tested whether M. mazei has a similar preference. Nevertheless, we reasoned that the S. shibatae topo VI cleavage sequence may contain general features recognized by type IIB topoisomerases, and using such a sequence was preferable to picking at random. We have modified the text (subsection “Topo VI is a distributive DNA relaxase that preferentially recognizes DNA crossings”, first paragraph) to highlight this point. Interestingly, in pilot experiments for cleavage of short duplexes (not discussed in the manuscript), M. mazei topo VI produced a small but detectable amount of cleavage of the 60 bp duplex bearing the S. shibatae sequence, but it did not cleave a 60 bp duplex with an unrelated sequence. We did not repeat this experiment with a 70bp oligo of random sequence.

https://doi.org/10.7554/eLife.31724.061

Article and author information

Author details

  1. Timothy J Wendorff

    Biophysics Graduate Program, University of California, Berkeley, Berkeley, United States
    Contribution
    Conceptualization, Formal analysis, Funding acquisition, Validation, Investigation, Visualization, Methodology, Writing—original draft, Writing—review and editing
    Competing interests
    No competing interests declared
  2. James M Berger

    Department of Biophysics and Biophysical Chemistry, Johns Hopkins University School of Medicine, Baltimore, United States
    Contribution
    Conceptualization, Resources, Supervision, Funding acquisition, Visualization, Methodology, Project administration, Writing—review and editing
    For correspondence
    jberge29@jhmi.edu
    Competing interests
    Reviewing editor, eLife
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-0666-1240

Funding

National Institutes of Health (RO1 CA077373)

  • James M Berger

National Science Foundation (DGE 1106400)

  • Timothy J Wendorff

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

The authors thank current and former members of the Berger lab, as well as Scott Keeney and Corentin Claeys Bouuaert for helpful discussions and critical feedback. The authors also thank Kevin Corbett for assisting with the use of models based on previously-published SAXS data. This work was supported by a National Science Foundation graduate research fellowship (DGE 1106400 to TJW) and the National Institute of Health (RO1 CA077373 to JMB).

Reviewing Editor

  1. Geeta J Narlikar, University of California, San Francisco, United States

Publication history

  1. Received: September 3, 2017
  2. Accepted: March 28, 2018
  3. Accepted Manuscript published: March 29, 2018 (version 1)
  4. Version of Record published: April 27, 2018 (version 2)
  5. Version of Record updated: May 11, 2018 (version 3)

Copyright

© 2018, Wendorff et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

Metrics

  • 1,174
    Page views
  • 271
    Downloads
  • 0
    Citations

Article citation count generated by polling the highest count across the following sources: Crossref, PubMed Central, Scopus.

Download links

A two-part list of links to download the article, or parts of the article, in various formats.

Downloads (link to download the article as PDF)

Download citations (links to download the citations from this article in formats compatible with various reference manager tools)

Open citations (links to open the citations from this article in various online reference manager services)

Further reading

    1. Biochemistry and Chemical Biology
    2. Microbiology and Infectious Disease
    Bridgette M Cumming et al.
    Tools and Resources Updated
    1. Biochemistry and Chemical Biology
    Adam J Pollak et al.
    Research Article