The genome of bacterial cells is in a negative-superhelical state through the combined activity of various DNA topoisomerases, enzymes that can alter the linking number between the two DNA strands constituting the bacterial genome (Bush et al., 2015; Vos et al., 2011). The only enzyme that can introduce negative supercoils is a type IIA topoisomerase, DNA gyrase. An active gyrase complex is constituted by two GyrA subunits, encoded by the gyrA gene, and two GyrB subunits, encoded by the gyrB gene, thus forming an A₂B₂ heterotramer (Figure 1b). The A₂B₂ complex can bind a segment of DNA (green, Figure 1b) sandwiched between the two TOPRIM domains of the GyrB subunits and the two tower domains of GyrA and sitting on the winged-helix domain (WHD) domain of GyrA. The enzyme is able to establish a transient double-strand break in this DNA segment through transesterification from a 5′-phosphate in the DNA backbone to a catalytic tyrosine (Y122 in Escherichia coli gyrase, Figure 1d). The DNA duplex that extends from the cleaved segment can wrap in a positive loop around the GyrA C-terminal domains (CTDs) and the other extremity of the loop can be passed through the various interfaces of the enzyme, including the DNA break, thereby converting the positive wrap into a negative one, and resulting in the introduction of negative supercoils by a step of 2, once the DNA break has been re-sealed (Brown and Cozzarelli, 1979; Mizuuchi et al., 1980; Figure 1b). Therefore, gyrase needs to be able to transport a DNA duplex through the whole enzyme, that is three interfaces (Figure 1a and b). The first one is constituted by the ATPase domain at the N-terminus of GyrB, whose dimerization is controlled by the binding of two ATPs and results in the trapping of the transported DNA (T) segment that enters through the open interface. This interface is dubbed the N-gate. The transport of DNA is coupled to the binding and hydrolysis of ATP. The second interface the duplex must go through is the DNA gate, which is constituted by the DNA break itself and a protein-protein interface between the two GyrA subunits (WHD domains). The duplex can then exit the enzyme through the C-gate (Figure 1b). There must be some coupling between the interfaces opening and closing since opening them all at the same time would cause the enzyme to come apart and the formation of a permanent double-stranded break (DSB). This is never observed at any detectable level with the purified enzyme in vitro.

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It has been suggested that the coupling between ATP binding and hydrolysis acts as a safety mechanism by only allowing DNA transport through the DNA gate and C-gate when the ATP interface (N-gate) is closed (Bates and Maxwell, 2007). However, gyrase can perform strand passage without ATP (and therefore with the N-gate opened), without introducing DSBs (Gellert et al., 1977; Sugino et al., 1977), suggesting that some allosteric coupling prevents the DNA gate and the C-gate from being opened at the same time. Consistent with this, the purified GyrA subunit forms stable dimers in solution, whereas the GyrB subunit can be observed as a monomer (see results) in the absence of ATP (Costenaro et al., 2007). While the N-gate is controlled by ATP, it is unclear what controls the opening of the DNA gate and the C-gate. However, clear structural evidence obtained, mainly with eukaryotic topoisomerase II, suggests coupling between motion of the WHD domains and the opening of the C-gate, pointing to a coordination between DNA cleavage status and the opening of the C-gate (Dong and Berger, 2007; Wendorff et al., 2012). In the case of gyrase, however, no structure with the C-gate open is available so it is difficult to conclude. It has also been suggested that the transported DNA segment itself controls the opening of the various interfaces for instance by ‘pushing open’ the DNA-gate (Bax et al., 2019; Roca, 2004).

DNA gyrase is the target for the clinically important antibiotics, the fluoroquinolones. These compounds bind the enzyme and stabilize the cleaved state, the so-called ‘cleavage complex’. The resulting DSBs are deleterious for the bacterial cell and are thought to be the basis for their antibacterial properties. However, in vitro, the cleavage complexes stabilized by fluoroquinolones are reversible, suggesting that the enzyme subunits do not come apart. It is therefore unclear how the reversible cleavage complexes are converted in vivo to irreversible DNA lesions. However, treatment of bacterial cells by fluoroquinolones clearly results in DSBs (Bush et al., 2020; Chen et al., 1996). In eukaryotic cells, pathways exist for the removal of the covalently bound topoisomerase II enzyme and exposure of the DNA lesion to regular DNA repair mechanisms (Pommier et al., 2014). In bacterial cells such pathways are not well known (but see Huang et al., 2021).

Some work has also suggested that in Escherichia coli gyrase cleavage complexes can induce illegitimate recombination between two DNA segments (Ikeda, 1994; Ikeda et al., 1981; Ikeda et al., 2004). One possible mechanism being the exchange of subunits between two cleavage complexes, followed by re-ligation. However, these experiments were conducted in a complex cellular extract and whether the observed recombination occurs through subunit exchange is not known. It has been shown that treatment with fluoroquinolones is associated with genomic rearrangements (López and Blázquez, 2009; López et al., 2007), potentially playing a role in their cytotoxicity, but again the exact mechanism remains unclear.

In this work we purify E. coli gyrase complexes with a heterodimeric GyrA interface as pioneered in the Klostermeier group (Gubaev et al., 2016). The complexes are purified as a dimer of GyrA and a fusion of GyrB and GyrA (BA fusion), encoded by different plasmids and thus allowing the introduction of targeted mutations on one side only of the complex (Figure 1—figure supplement 1). An active enzyme is reconstituted by the addition of free purified GyrB. We show that mutating the catalytic tyrosine on one side completely abolishes cleavage on this side of the complex and severely compromises negative supercoiling activity by removing the ability to double-strand cleave DNA. We show that the complex can nevertheless reconstitute double-strand cleavage activity, underlying the reconstitution of negative supercoiling activity by strand-passage through the double-strand break. We suggest that this reconstitution of double-strand cleavage activity occurs when the DNA gate interface of a gyrase dimer is open, and then re-forms by association between one subunit from one dimer with another subunit coming from another dimer (Figure 1—figure supplement 2). The exchanged interface functionality would be conserved as the contacts and configuration of the exchanged interface remains identical to the unexchanged interface, and therefore this process, which we dub ‘interface swapping’ (IS), can reconstitute double-strand cleavage when the exchanged interface presents 2 active tyrosine. Interface swapping can lead to whole ‘subunit exchange’ depending on the interfaces involved (Figure 1b, Figure 1—figure supplement 2). For instance, swapping interfaces 2 and 3 leads to the whole GyrA subunit to be exchanged, whereas to exchange a BA fusion subunit all three interfaces needs to be exchanged. In addition, we show that interface swapping is favored by the presence of the free GyrB subunit and that these free GyrB subunits can multimerize and pull the heterodimers into higher order multimers. We show that mutating isoleucine 174 in GyrA (E. coli numbering, Figure 1c) abolishes DNA cleavage but not DNA binding and only lowers wrapping activity. Introducing the I174G mutation on one side of the heterodimer severely reduces cleavage activity suggesting that proper DNA bending and/or wrapping favors interface swapping. Altogether, our data suggest that DNA binding and wrapping and the accumulation of gyrase in higher order oligomers can favor rapid interface exchange between two active gyrase complexes brought into proximity. We discuss the possible mechanism and suggest that gyrase can recombine two DNA duplexes through interface exchange.

In this work, we have purified stable heterodimers of GyrA and a GyrBA fusion construct (Figure 2). We have shown that mutating the catalytic tyrosine on one side of the heterodimer completely abolished cleavage on this side only (Figure 4). We observed the reconstitution of double-strand cleavage activity from a BA_F.A and BA.A_F mutated heterodimers with a mutated catalytic tyrosine on one side (Figure 3). This activity was dependent on the addition of a high concentration of GyrB (Figure 5). The appearance of this double-strand cleavage was delayed by a few minutes compared to the unmutated heterodimer (Figure 7). We conclude that gyrase can rapidly (within a few minutes) exchange its DNA gate sides in vitro, thereby allowing the mutated heterodimer to reconstitute double-strand cleavage activity and therefore supercoiling activity (Figure 3). We call this process ‘interface swapping’. Interface swapping can result in whole subunit exchange depending on the number of interfaces involved (Figures 1 and 8, Figure 1—figure supplement 2).

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We have carefully controlled our heterodimer preparation for contaminations that could explain these results. Our heterodimer preparations do not show any contamination by GyrA dimers either on BN-PAGE, by mass photometry or native mass spectrometry (Figure 2, Figure 2—figure supplement 1). The low contamination by either (BA)₂ or BA.A.B would produce double-strand cleavage independent of GyrB addition. This background cleavage is minimal (Figure 3). Therefore, we can confidently conclude that the cleavage reconstituted with the mutated BA_F.A heterodimers does not arise from a contamination by GyrA dimers (see Supplementary discussion for the BA.A_F complex).

The observation that heterodimeric single-sided catalytic mutant preparations (e.g. BA.A_F, BA_F.A, BA_F.A₅₉) still have some supercoiling activity, which we have confirmed, has led others (Gubaev et al., 2016) to propose a mechanism by which gyrase introduces negative supercoiling through nicking of the DNA (which requires only one catalytic tyrosine) and swivel relaxation of the DNA wrapped around the CTD domain of GyrA. We conclude that such a mechanism is most likely incorrect for several reasons. The first is a theoretical consideration. It is known that gyrase can introduce positive supercoiling by relaxing the constrained positive wrap around the CTD in the absence of ATP. Therefore, the default position of the CTD is the constraint of positive supercoils and their relaxation necessarily implies release from the CTD, which is favored by the ATPase activity of the enzyme (Basu et al., 2018; Basu et al., 2012). On average, +0.74–0.8 supercoils is constrained by wrapping around the CTD (Basu et al., 2012; Kampranis et al., 1999). The relaxation of a single positive loop (with a~+1 contribution to the ΔLk) should therefore introduce negative supercoiling by steps of 1. Only the conversion of a positive loop (+1) to a negative (–1) by strand passage can introduce supercoiling by steps of 2. Since a gyrase complex has two CTDs, it could be assumed that the simultaneous relaxation of the two loops could introduce supercoiling by step of two, however, Klostermeier and co-workers have proved that only one CTD is needed for introducing supercoils by steps of two (Stelljes et al., 2018). Therefore, for the model to stand, it must be assumed that, in addition to the (+1) supercoils stabilized by the writhe of the loop, another +1 must be stabilized in the twist of the double helix within this short loop. We find this to be unlikely considering recent cryo-EM structures of gyrase with DNA (Vanden Broeck et al., 2019). However, a CTD-wrapped state with +1.7 supercoils has been detected in the absence of ATP (Basu et al., 2012). It is therefore conceivable that the relaxation of DNA wrapped in this state could lead to the introduction of supercoiling by step of 2 via swiveling. This state is much less abundant than the +1 state however and the efficiency of the swiveling mechanism would be very low. For an efficient mechanism involving relaxation by swiveling of this +1.7 state one would have to assume that ATP favors its formation, in addition to promoting the release of the DNA loop. These two activities contradict each other. In addition, the addition of nucleotides either did not favor the +1.7 state or did not change its abundance (Basu et al., 2018). Finally, this state could represent twin-wrapping around the 2 CTDs, and only one CTD is sufficient for supercoiling, as stated above. In conclusion, introduction of negative supercoiling by swiveling-mediated relaxation of the wrapped DNA loop would have to most likely occur by steps of 1, or be extremely inefficient when they occur by steps of two.

On top of the above consideration ours and others’ (Gubaev et al., 2016) results can be explained by DNA gate interface swapping that reconstitute the double-strand cleavage activity that is necessary for negative supercoiling activity. Figure 8—figure supplements 1–4, Figure 8—figure supplement 7 show schematics representing how the process of interface swapping and subunit exchange could happen for our various heterodimer constructions and some constructions reported elsewhere (Gubaev et al., 2016). This model allows some predictions about the abundance and activity of each species present in the prep, either initially or after IS and SE. These predictions are consistent with our experimental data and, we suggest, data reported by Gubaev et al., 2016.

In almost all heterodimer configurations, mutating one catalytic tyrosine results in an important diminution of supercoiling activity, accompanied by a drastic diminution of double-strand cleavage activity, suggesting that most of the activity is based on strand passage through a double-strand break. In a configuration where the reconstitution of double-strand cleavage activity remained important (such as BA_F.A, Figure 8—figure supplement 1), it correlates with a robust supercoiling activity, further supporting that supercoiling activity is based on the ability to cleave DNA in both strands. These results argue strongly against swiveling since the later only requires a single-strand break, and therefore should be unaffected by mutating one catalytic tyrosine.

A series of experiments (Gubaev et al., 2016) were interpreted as excluding subunit exchange and interface swapping (the authors do not distinguish between the two). Namely the absence of double-strand cleavage observed with heterodimers and FRET experiments interpreting the absence of FRET as showing that the DNA gates of different heterodimer never come into close proximity. We argue that these experiments do not fully exclude SE or IS. Negative supercoiling by DNA gyrase is an extremely efficient reaction and this activity can be detected with a very low concentration of the active enzyme. Consequently, it is possible to detect efficient negative supercoiling and not be able to detect the double-strand cleavage from which the activity originates because of the low concentration of active enzyme. We argue that DNA gate IS that reconstitute double-strand cleavage activity occurs but is not detected by cleavage assays or FRET (Gubaev et al., 2016) because of the low proportion of complexes where IS has occurred. Despite the low number of exchanged complexes able to double-strand cleave DNA, negative supercoiling can be observed because of the high efficiency of the reaction.

We have shown that efficient double-strand cleavage reconstitution by DNA gate IS only occurs in specific conditions, namely with the BA_F.A complex in the presence of an excess of GyrB. In Gubaev et al., 2016, cleavage experiments are only shown for BA.A_F, and display low double-strand cleavage. This agrees with our experiments that show barely detectable double-strand cleavage with the BA.A_F complex (Figure 8—figure supplement 8–supp. and 2). Considering that a low amount of DNA gate IS can explain results shown in Gubaev et al., more experiments are needed to exclude it. For instance, cleavage assays with the heterodimer are shown (Gubaev et al., 2016), and we argue that there is a very slight increase in double-strand breaks observed with BA.A_F +GyrB and not observed in the absence of GyrB. More experiments could have been done to investigate this, namely increase the GyrB concentration (it is known that GyrB favors cleavage) and purify the cleavage complexes. We have done this with the BA.A_F and shown unambiguously that a low amount of DNA gate IS occurs (see below for explanation of the difference between the two types of heterodimers).

In addition, the supercoiling activity seen with the heterodimer is delayed by several minutes compared to the wild-type homodimer (Gubaev et al., 2016). This is completely consistent with our experiment showing a delay of several minutes in the appearance of double-strand cleavage with the heterodimer (Figure 7). If the wild-type enzyme and the heterodimer use the same swiveling mechanism, why is this delay observed? Another way to ask the same question would be: if gyrase uses a swiveling mechanism that only uses one catalytic tyrosine, why does the efficiency of supercoiling go down when a single tyrosine is mutated on one side? This is not addressed (Gubaev et al., 2016); we suggest that the efficiency of supercoiling by heterodimers will be correlated to the efficiency of double-strand cleavage reconstitution. Consistently, complexes with a higher level of reconstituted double-strand cleavage (BA_F.A) have a higher supercoiling efficiency.

We have observed that heterodimers that should be incapable of reconstituting an active gyrase when the whole subunits are exchanged, namely BA_F.A₅₉, is still able to reconstitute a low supercoiling activity (Figure 6). We have interpreted this result by the observed low contamination by BA_F.A_endogenous, which undergoes subunit exchange (Figure 8—figure supplement 8–supp. and 3). Elsewhere it is reported that the reverse complex, BA₅₉.A_F (called BA_ΔCTD.A_F) can reconstitute supercoiling activity (Gubaev et al., 2016). However, the observed supercoiling is very low, much lower than the other heterodimers and the reason for this is not discussed. Such an amount of supercoiling could be explained by even the most minimal contamination by an active double-strand cleaving gyrase and therefore extreme caution should be used to interpret these data. Deleting the CTD on one side when two tyrosines are present does not reduces supercoiling activity as much and therefore, at the very least, one should conclude that the swiveling mechanism is much less efficient than the strand-passage mechanism which requires requires two tyrosines. We argue that BA₅₉.A_F (Gubaev et al., 2016) could also be contaminated with BA₅₉.A_endogenous, which would have a lower efficiency of subunit exchange than BA_F.A_endogenous (this study) since they lack a CTD on the BA fusion side (see Supplementary discussion and Figure 8—figure supplement 4). This would explain why the supercoiling activity of BA₅₉.A_F is so low. The final possibility would be that the exchanged DNA gate could use the CTD of the neighboring complex for wrapping. However, our BA_LLL.A₅₉ shows that it is not the case. BA_LLL.A₅₉ shows higher reconstitution of double-strand cleavage through interface swapping. The reconstituted active DNA gate is therefore between the two A₅₉ subunits and if this DNA gate could use the CTD from the neighboring BA fusion, we should expect this extra double-strand cleavage to produce negative supercoiling activity. The extra IS observed with BA_LLL.A₅₉ is not accompanied by supercoiling activity (Figure 6—figure supplement 1a) and therefore the data shows that an active DNA gate can only use a CTD from the same GyrA subunit for strand passage and negative supercoiling. We conclude that the low supercoiling reconstituted by BA_F.A₅₉ (this study) and BA₅₉.A_F (Gubaev et al., 2016) are likely to originate from low amount of contamination (see Supplementary discussion).

The reasons for the asymmetry between BA_F.A and BA.A_F are unclear so far. With the BA_F.A heterodimer subunit exchange produces an active GyrA dimer and an inactive (BA_F)₂ dimer. In the case of BA.A_F, subunit exchange reconstitutes an active (BA)₂ dimer and an inactive GyrA_F dimer. Therefore, the difference in the level of cleavage and supercoiling activity reconstituted could stem from a difference in activity between the GyrA dimer (with free GyrB) and the (BA)₂ dimer. We have directly tested this and compared both cleavage and supercoiling activity between GyrA dimer (‘wild type’), (BA)₂ dimer and BA.A heterodimer (Figure 8—figure supplement 5). We found that the activity of (BA)₂ is reduced by a factor 100. Interestingly, the heterodimer has a level of activity intermediate between the GyrA dimer and the (BA)₂ dimer. This is true for both cleavage and supercoiling. These results can explain the asymmetry between BA_F.A and BA.A_F since the subunit exchange with the former produces a more active complex by subunit exchange compared to the later. These results also show that fusing GyrB to GyrA limits both cleavage and supercoiling activity. We did not add a linker to our fusion and therefore the position of GyrB with respect to GyrA and DNA is expected to be more constrained than with free GyrB. It has been reported that the GyrB dimer is leaning to one side with respect to the dyad axis in the gyrase-DNA complex (Vanden Broeck et al., 2019). Therefore, in this complex, the position of each GyrB subunit with regard to GyrA is different and this flexibility of GyrB in the gyrase complex might be important for its activity. We speculate that with a GyrB fused to GyrA limits the leaning ability of the GyrB dimer and therefore reduces both the supercoiling activity and the ability to form cleavage complexes.

This leads us to discuss the exact mechanism of interface swapping and subunit exchange. It is a priori a surprising result considering the stability of the gyrase dyad interface and its importance in ensuring genome stability. However, during strand passage all three interfaces must be broken and the opening of the interfaces necessarily implies the possibility of exchange between two complexes prior to their reformation. The integrity of the complex is ensured through sequential opening and closing of the three interfaces. This sequential opening and closing allows the transported DNA to go through the whole complex without it coming apart. Similarly, subunit exchange could also occur sequentially, one interface after the other, allowing the partial or whole exchange to occur without the complex coming apart (Figure 1b, Figure 1—figure supplement 2). This model would only require gyrase complexes being able to come together and interact in an appropriate configuration and we indeed observe that both GyrA dimers and GyrB monomers can multimerize, each on their own in solution, supporting this hypothesis. In fact, our results only show that the DNA-cleaving interface is exchanged (DNA gate IS); it is possible that this is not followed by the exchange of the C-gate interface, producing a sort of helical super-complex where one interface interacts with a different subunit compared to the other interface (Figure 1—figure supplement 2, Figure 8 – Figure 7). Structural evidence exists for such a complex. In earlier work (Rudolph and Klostermeier, 2013) a GyrA structure with the DNA gate opened was obtained. The DNA gate interacts with the next GyrA subunit forming a helical oligomer of GyrA within the crystal. We suggest that this structure could underly IS in our system, with the C-gate interface remaining closed, while the DNA gate interface opens and reforms with the neighboring complex. It is even possible that such a super-complex would retain negative supercoiling activity. The opening of the DNA gate does not occur by the pulling apart of the GyrA WHD domains but rather by a sliding of the two GyrA domains against one another along an axis close to being perpendicular to a plane formed by the dyad axis and the DNA axis (Chen et al., 2018; Germe et al., 2018; Wendorff et al., 2012). Therefore, the freed DNA-gate interface could be available to dimerize with a neighboring interface similarly freed by the opening of the DNA gate. The only requirement being that the neighboring complex is positioned parallel to the other one, in such a way that the simultaneous opening of the DNA gates results in the meeting of the two DNA gate interfaces between the complexes, thereby achieving interface swapping (Figure 8, Figure 1—figure supplement 2). If within this complex only the DNA-gate is swapped with a neighboring heterodimer, one would expect the reconstitution of an active super-complex where the swapped DNA-gate reconstitutes double-strand DNA cleavage while the unswapped ATPase domains retain strand capture activity (see Supplementary discussion).

We have tried and failed to directly observe subunit exchange by mass photometry. We tried to observe the reconstitution of a GyrA dimer from a heterodimer. However, this experiment is complicated because of several factors. The first is the necessity of a high concentration of GyrB. This will mask any GyrA monomer that might be produced and will bind any GyrA dimer that is produced reconstituting a trimer (with a MW equivalent to the heterodimer) or a tetramer (with a MW equivalent to GyrB bound to a heterodimer) that are indistinguishable from the original complex. We have not observed a significant change of the mass photometry profile of a mixture of GyrB +BA .A heterodimer over time. In addition, our data suggest that proper DNA binding and bending is necessary for interface swapping. Adding a long piece of DNA only scrambles the mass photometry profile as the complexes binds to it to form high MW particles. We have tried to add a short, purified DNA fragment (170 bp) which gives a nice peak on its own in mass photometry but again failed to see a difference. It could be that longer DNA is needed to allow the wrapping of several CTDs along the DNA and position the complexes for subunit exchange (see below). It is also possible that only the DNA-gate interfaces are exchanged and therefore no GyrA dimers are ever produced, only higher MW oligomers that are difficult to detect. Indeed, interface swapping does not necessarily result in the coming apart and complete separation of the whole complex (see above). Therefore, due to its partial nature, complexes with swapped interfaces are likely difficult to observe. This could further account for the lack of subunit exchange observed by FRET. FRET can only be observed on a stable complex, and we note that it was tested either before or after the supercoiling reaction not during the reaction when they would occur transiently. The only condition that showed stable, exchanged interfaces is with the BA_F.A +GyrB reaction in the presence of ciprofloxacin which produces stable, reversible double-strand cleavage. We surmise that cross-linking the DNA interface in these could provide direct evidence of interface swapping. Gubaev et al., 2016 have done no cross-linking experiments; these experiments are not straightforward since they require specific cross-linking at a functional DNA gate and a general cross-linker cannot be used.

We have shown that efficient IS requires the CTD or wrapping domain of GyrA. IS is quick in our system and removing the CTD produced much less double-strand cleavage within the 30 min timeframe of our experiment. We suggest that the wrapping of the DNA is necessary for the positioning of two or several gyrase complexes in such a way that interface exchange by the mechanism proposed above is favored. It is also possible that the CTD favors the opening of the DNA-gate interface, an idea supported by the observation that full-length gyrase has ATP-independent strand passage activity but not the A₅₉ dimer +GyrB.

We also showed that mutating isoleucine 174 drastically diminished the amount of IS. This could be interpreted in various ways. First, proper DNA bending is necessary for DNA cleavage (Dong and Berger, 2007; Lee et al., 2013), and X-ray structures have shown that DNA binding and bending correlate with a slight opening of the DNA gate (Germe et al., 2018; Wendorff et al., 2012). Therefore, it could be that the binding of the DNA loosens the DNA gate interface and favors its opening. The second interpretation is that, although the isoleucine mutant can still bind and wrap DNA, the wrapping efficiency is reduced and therefore indirectly affects interface swapping since the CTD is important for IS.

We also observed that mutating the two residues neighboring the catalytic tyrosine favored IS. The mutation converts polar side chains to aliphatic ones and likely affects interface interaction. Indeed, the catalytic tyrosine and its neighboring residues are in proximity to the TOPRIM domain in the gyrase apo structure (Bax et al., 2019). Upon binding DNA, the distance between them increases and DNA cleavage is accompanied by the full disengagement of the catalytic tyrosine from the TOPRIM domain. The opening of the DNA gate fully separates the catalytic loop, which contains the catalytic tyrosine, from the TOPRIM domain by a sliding motion. Therefore, mutations disfavoring interaction between the catalytic loop and the TOPRIM domain are likely to favor DNA gate opening and therefore subunit exchange.

Subunit exchange, which necessitates IS, has been proposed as the mechanism behind DNA gyrase-mediated illegitimate recombination (Ikeda et al., 2004). Gyrase was shown to be involved in the transfer of an antibiotic resistance cassette from a plasmid to lambda DNA in a packaging reaction, in the presence of a quinolone (Ikeda et al., 1982; Ikeda et al., 1981). This result was independent of any homologous recombination pathway in either the phage or bacterium (Ikeda et al., 1982; Ikeda et al., 1981). The authors suggested that gyrase-mediated recombination in one of two ways: either (1) by dissolution of the gyrase complex into heterodimers (GyrA:GyrB) that still had the DNA bound in the phosphotyrosyl bond, which could reassociate with another GyrA:GyrB heterodimer in a similar situation, allowing for recombination, or (2) by the formation of higher order oligomers (GyrA₄GyrB₄) and interface swapping, which upon dissolution of the higher-order complex could result in subunit exchange and recombination (Ikeda, 1994; Ikeda et al., 2004). We suggest the former may be the cause of slower subunit exchange observed in the formation of GyrA and GyrA59 heterodimers over several days (Figure 2—figure supplement 2). This slow subunit exchange is inhibited by DNA and ciprofloxacin in contrast with the rapid subunit exchange observed with heterodimers. Therefore, the two are mechanistically distinct. Indeed, solution-based slow subunit exchange requires the complete separation of the complex for the formation of free monomers but does not require the formation of higher-order oligomers. In contrast, rapid subunit exchange by interface swapping does not require the complete separation of the complex but does involve the formation of higher-order oligomers. It therefore makes sense that one, and not the other would be inhibited by DNA and ciprofloxacin. Ciprofloxacin-induced cleavage complexes do not come apart in solution, which is consistent with ciprofloxacin inhibiting the formation of free monomers. Ciprofloxacin does not prevent rapid interface swapping and it has been shown that it does not prevent DNA-gate opening (Chan et al., 2017). Finally, while deleting the CTD affects rapid subunit exchange, it does not prevent, slow, solution-based exchange (Figure 2—figure supplement 2). We surmise that illegitimate recombination may occur within higher-order oligomer formation followed by interface swapping (the DNA gate specifically). This is consistent with the timescale of illegitimate recombination and explains how two DNA duplexes are brought together for recombination by two gyrase complexes.

Considering the ubiquity and abundance of gyrase in bacterial cells, the rapid interface swapping mechanism is expected to occur in vivo where DNA is present and gyrase is highly concentrated. If interfaces are swapped when gyrase is covalently bound to cleaved DNA on both sides, followed by re-ligation with the swapped DNA-gate, this would result in illegitimate recombination. The way the two gyrase complexes interact would bring together two DNA duplexes for recombination. The presence of fluoroquinolones, by extending the lifespan of the cleaved state, would make this process much more likely. We have observed that GyrB on its own can oligomerize in a regular fashion where the intensity of the dimer, trimer, tetramer and so forth diminishes following roughly an exponential until the oligomers can no longer be resolved (carefully consider Figure 2—figure supplement 2a, lane B). This observation strongly suggests a regular oligomerization and tentatively a linear oligomer of a GyrB monomer repetition. We have observed that a high gyrase/DNA ratio in the presence of ciprofloxacin produces cleavage complexes ~200 bp apart (Figure 8—figure supplement 6). Considering that gyrase can interact with ~150 bp of DNA through DNA wrapping, it suggests that the DNA exiting a gyrase complex is immediately engaged in another neighboring gyrase-DNA complex. This linear repetition of cleavage complexes on DNA (spanning the whole plasmid in this instance) mirror the linear repetition of GyrB monomers within GyrB oligomers. We suggest that GyrB can drive the close repetition of many cleavage complexes on linear DNA. Again, the high concentration of gyrase in bacterial cells is consistent with the possibility of a long-range linear gyrase oligomer forming on the genome of bacteria. The tight packing of gyrase, observed on plasmids, and potentially driven by GyrB oligomerization, therefore opens the possibility of micro-deletions or -inversions by interface swapping of the cleavage complexes on the bacterial genome, induced by fluoroquinolones. The necessary re-ligation can happen if the compound is washed-out, and we speculate that temporary exposure to high dose of fluoroquinolones could induce these rearrangements. Longer-range genomic rearrangements are also conceivable depending on the organization of gyrase clusters in the bacterial cell. This interface swapping mechanism could therefore have profound consequences for antibiotics treatment of infections and more generally the evolution of the bacterial lineage, within which DNA gyrase is ubiquitous.

All the raw data are freely available at https://doi.org/10.5061/dryad.w6m905qwn. Reagents are freely available by writing to thomas.germe@cuanschutz.edu.

1. 1. Germe T

   (2024) Dryad Digital Repository

   Rapid, DNA-induced interface swapping by DNA gyrase.

   https://doi.org/10.5061/dryad.w6m905qwn

1. 1. Basu A
   2. Schoeffler AJ
   3. Berger JM
   4. Bryant Z

   (2012) ATP binding controls distinct structural transitions of Escherichia coli DNA gyrase in complex with DNA

   Nature Structural & Molecular Biology 19:538–546.

   https://doi.org/10.1038/nsmb.2278

   • PubMed
   • Google Scholar
2. 1. Basu A
   2. Hobson M
   3. Lebel P
   4. Fernandes LE
   5. Tretter EM
   6. Berger JM
   7. Bryant Z

   (2018) Dynamic coupling between conformations and nucleotide states in DNA gyrase

   Nature Chemical Biology 14:565–574.

   https://doi.org/10.1038/s41589-018-0037-0

   • PubMed
   • Google Scholar
3. 1. Bates AD
   2. Maxwell A

   (2007) Energy coupling in type II topoisomerases: why do they hydrolyze ATP?

   Biochemistry 46:7929–7941.

   https://doi.org/10.1021/bi700789g

   • PubMed
   • Google Scholar
4. 1. Bax BD
   2. Murshudov G
   3. Maxwell A
   4. Germe T

   (2019) DNA Topoisomerase Inhibitors: Trapping a DNA-Cleaving Machine in Motion

   Journal of Molecular Biology 431:3427–3449.

   https://doi.org/10.1016/j.jmb.2019.07.008

   • PubMed
   • Google Scholar
5. 1. Brown PO
   2. Cozzarelli NR

   (1979) A sign inversion mechanism for enzymatic supercoiling of DNA

   Science 206:1081–1083.

   https://doi.org/10.1126/science.227059

   • PubMed
   • Google Scholar
6. 1. Bush NG
   2. Evans-Roberts K
   3. Maxwell A

   (2015) DNA Topoisomerases

   EcoSal Plus 6:14.

   https://doi.org/10.1128/ecosalplus.ESP-0010-2014

   • PubMed
   • Google Scholar
7. 1. Bush NG
   2. Diez-Santos I
   3. Abbott LR
   4. Maxwell A

   (2020) Quinolones: Mechanism, lethality and their contributions to antibiotic resistance

   Molecules 25:5662.

   https://doi.org/10.3390/molecules25235662

   • PubMed
   • Google Scholar
8. 1. Chan PF
   2. Germe T
   3. Bax BD
   4. Huang J
   5. Thalji RK
   6. Bacqué E
   7. Checchia A
   8. Chen D
   9. Cui H
   10. Ding X
   11. Ingraham K
   12. McCloskey L
   13. Raha K
   14. Srikannathasan V
   15. Maxwell A
   16. Stavenger RA

   (2017) Thiophene antibacterials that allosterically stabilize DNA-cleavage complexes with DNA gyrase

   PNAS 114:E4492–E4500.

   https://doi.org/10.1073/pnas.1700721114

   • PubMed
   • Google Scholar
9. 1. Chen CR
   2. Malik M
   3. Snyder M
   4. Drlica K

   (1996) DNA gyrase and topoisomerase IV on the bacterial chromosome: quinolone-induced DNA cleavage

   Journal of Molecular Biology 258:627–637.

   https://doi.org/10.1006/jmbi.1996.0274

   • PubMed
   • Google Scholar
10. 1. Chen SF
    2. Huang NL
    3. Lin JH
    4. Wu CC
    5. Wang YR
    6. Yu YJ
    7. Gilson MK
    8. Chan NL

    (2018) Structural insights into the gating of DNA passage by the topoisomerase II DNA-gate

    Nature Communications 9:3085.

    https://doi.org/10.1038/s41467-018-05406-y

    • PubMed
    • Google Scholar
11. 1. Costenaro L
    2. Grossmann JG
    3. Ebel C
    4. Maxwell A

    (2007) Modular structure of the full-length DNA gyrase B subunit revealed by small-angle X-ray scattering

    Structure 15:329–339.

    https://doi.org/10.1016/j.str.2007.01.013

    • PubMed
    • Google Scholar
12. 1. Dong KC
    2. Berger JM

    (2007) Structural basis for gate-DNA recognition and bending by type IIA topoisomerases

    Nature 450:1201–1205.

    https://doi.org/10.1038/nature06396

    • PubMed
    • Google Scholar
13. 1. Gellert M
    2. Mizuuchi K
    3. O’Dea MH
    4. Itoh T
    5. Tomizawa JI

    (1977) Nalidixic acid resistance: a second genetic character involved in DNA gyrase activity

    PNAS 74:4772–4776.

    https://doi.org/10.1073/pnas.74.11.4772

    • PubMed
    • Google Scholar
14. 1. Germe T
    2. Vörös J
    3. Jeannot F
    4. Taillier T
    5. Stavenger RA
    6. Bacqué E
    7. Maxwell A
    8. Bax BD

    (2018) A new class of antibacterials, the imidazopyrazinones, reveal structural transitions involved in DNA gyrase poisoning and mechanisms of resistance

    Nucleic Acids Research 46:4114–4128.

    https://doi.org/10.1093/nar/gky181

    • PubMed
    • Google Scholar
15. 1. Gubaev A
    2. Weidlich D
    3. Klostermeier D

    (2016) DNA gyrase with a single catalytic tyrosine can catalyze DNA supercoiling by a nicking-closing mechanism

    Nucleic Acids Research 44:10354–10366.

    https://doi.org/10.1093/nar/gkw740

    • PubMed
    • Google Scholar
16. 1. Hartmann S
    2. Gubaev A
    3. Klostermeier D

    (2017) Binding and Hydrolysis of a Single ATP Is Sufficient for N-Gate Closure and DNA Supercoiling by Gyrase

    Journal of Molecular Biology 429:3717–3729.

    https://doi.org/10.1016/j.jmb.2017.10.005

    • PubMed
    • Google Scholar
17. 1. Huang S-YN
    2. Michaels SA
    3. Mitchell BB
    4. Majdalani N
    5. Vanden Broeck A
    6. Canela A
    7. Tse-Dinh Y-C
    8. Lamour V
    9. Pommier Y

    (2021) Exonuclease VII repairs quinolone-induced damage by resolving DNA gyrase cleavage complexes

    Science Advances 7:eabe0384.

    https://doi.org/10.1126/sciadv.abe0384

    • PubMed
    • Google Scholar
18. Conference

    1. Ikeda H
    2. Moriya K
    3. Matsumoto T

    (1981) In vitro study of illegitimate recombination: involvement of DNA gyrase

    Cold Spring Harbor Symposia on Quantitative Biology. pp. 399–408.

    https://doi.org/10.1101/sqb.1981.045.01.054

    • Google Scholar
19. 1. Ikeda H
    2. Aoki K
    3. Naito A

    (1982) Illegitimate recombination mediated in vitro by DNA gyrase of Escherichia coli: structure of recombinant DNA molecules

    PNAS 79:3724–3728.

    https://doi.org/10.1073/pnas.79.12.3724

    • PubMed
    • Google Scholar
20. 1. Ikeda H

    (1994) DNA topoisomerase-mediated illegitimate recombination

    Advances in Pharmacology 29A:147–165.

    https://doi.org/10.1016/s1054-3589(08)60544-x

    • PubMed
    • Google Scholar
21. 1. Ikeda H
    2. Shiraishi K
    3. Ogata Y

    (2004)

    Illegitimate recombination mediated by double-strand break and end-joining in Escherichia coli

    Advances in Biophysics 38:3–20.

    • PubMed
    • Google Scholar
22. Software

    1. Jones E
    2. Oliphant T
    3. Peterson P

    (2001) "Scipy: open source scientific tools for python

    Python.

    http://www.scipy.org/
23. 1. Kampranis SC
    2. Bates AD
    3. Maxwell A

    (1999) A model for the mechanism of strand passage by DNA gyrase

    PNAS 96:8414–8419.

    https://doi.org/10.1073/pnas.96.15.8414

    • PubMed
    • Google Scholar
24. 1. Lee I
    2. Dong KC
    3. Berger JM

    (2013) The role of DNA bending in type IIA topoisomerase function

    Nucleic Acids Research 41:5444–5456.

    https://doi.org/10.1093/nar/gkt238

    • Google Scholar
25. 1. López E
    2. Elez M
    3. Matic I
    4. Blázquez J

    (2007) Antibiotic-mediated recombination: ciprofloxacin stimulates SOS-independent recombination of divergent sequences in Escherichia coli

    Molecular Microbiology 64:83–93.

    https://doi.org/10.1111/j.1365-2958.2007.05642.x

    • PubMed
    • Google Scholar
26. 1. López E
    2. Blázquez J

    (2009) Effect of subinhibitory concentrations of antibiotics on intrachromosomal homologous recombination in Escherichia coli

    Antimicrobial Agents and Chemotherapy 53:3411–3415.

    https://doi.org/10.1128/AAC.00358-09

    • PubMed
    • Google Scholar
27. 1. Maxwell A
    2. Howells AJ

    (1999) Overexpression and purification of bacterial DNA gyrase

    Methods in Molecular Biology 94:135–144.

    https://doi.org/10.1385/1-59259-259-7:135

    • PubMed
    • Google Scholar
28. 1. Mizuuchi K
    2. Fisher LM
    3. O’Dea MH
    4. Gellert M

    (1980) DNA gyrase action involves the introduction of transient double-strand breaks into DNA

    PNAS 77:1847–1851.

    https://doi.org/10.1073/pnas.77.4.1847

    • PubMed
    • Google Scholar
29. 1. Pommier Y
    2. Huang SN
    3. Gao R
    4. Das BB
    5. Murai J
    6. Marchand C

    (2014) Tyrosyl-DNA-phosphodiesterases (TDP1 and TDP2)

    DNA Repair 19:114–129.

    https://doi.org/10.1016/j.dnarep.2014.03.020

    • PubMed
    • Google Scholar
30. 1. Reece RJ
    2. Maxwell A

    (1989)

    Tryptic fragments of the Escherichia coli DNA gyrase A protein

    The Journal of Biological Chemistry 264:19648–19653.

    • PubMed
    • Google Scholar
31. 1. Reece RJ
    2. Maxwell A

    (1991)

    Probing the limits of the DNA breakage-reunion domain of the Escherichia coli DNA gyrase A protein

    The Journal of Biological Chemistry 266:3540–3546.

    • PubMed
    • Google Scholar
32. 1. Roca J

    (2004) The path of the DNA along the dimer interface of topoisomerase II

    The Journal of Biological Chemistry 279:25783–25788.

    https://doi.org/10.1074/jbc.M402555200

    • PubMed
    • Google Scholar
33. 1. Rudolph MG
    2. Klostermeier D

    (2013) Mapping the spectrum of conformational states of the DNA- and C-gates in Bacillus subtilis gyrase

    Journal of Molecular Biology 425:2632–2640.

    https://doi.org/10.1016/j.jmb.2013.04.010

    • PubMed
    • Google Scholar
34. 1. Shuman S
    2. Kane EM
    3. Morham SG

    (1989) Mapping the active-site tyrosine of vaccinia virus DNA topoisomerase I

    PNAS 86:9793–9797.

    https://doi.org/10.1073/pnas.86.24.9793

    • PubMed
    • Google Scholar
35. 1. Stelljes JT
    2. Weidlich D
    3. Gubaev A
    4. Klostermeier D

    (2018) Gyrase containing a single C-terminal domain catalyzes negative supercoiling of DNA by decreasing the linking number in steps of two

    Nucleic Acids Research 46:6773–6784.

    https://doi.org/10.1093/nar/gky470

    • PubMed
    • Google Scholar
36. 1. Sugino A
    2. Peebles CL
    3. Kreuzer KN
    4. Cozzarelli NR

    (1977) Mechanism of action of nalidixic acid: purification of Escherichia coli nalA gene product and its relationship to DNA gyrase and a novel nicking-closing enzyme

    PNAS 74:4767–4771.

    https://doi.org/10.1073/pnas.74.11.4767

    • PubMed
    • Google Scholar
37. 1. Vanden Broeck A
    2. Lotz C
    3. Ortiz J
    4. Lamour V

    (2019) Cryo-EM structure of the complete E. coli DNA gyrase nucleoprotein complex

    Nature Communications 10:4935.

    https://doi.org/10.1038/s41467-019-12914-y

    • PubMed
    • Google Scholar
38. 1. Vos SM
    2. Tretter EM
    3. Schmidt BH
    4. Berger JM

    (2011) All tangled up: how cells direct, manage and exploit topoisomerase function

    Nature Reviews. Molecular Cell Biology 12:827–841.

    https://doi.org/10.1038/nrm3228

    • PubMed
    • Google Scholar
39. 1. Wendorff TJ
    2. Schmidt BH
    3. Heslop P
    4. Austin CA
    5. Berger JM

    (2012) The structure of DNA-bound human topoisomerase II alpha: conformational mechanisms for coordinating inter-subunit interactions with DNA cleavage

    Journal of Molecular Biology 424:109–124.

    https://doi.org/10.1016/j.jmb.2012.07.014

    • PubMed
    • Google Scholar
40. 1. Young G
    2. Hundt N
    3. Cole D
    4. Fineberg A
    5. Andrecka J
    6. Tyler A
    7. Olerinyova A
    8. Ansari A
    9. Marklund EG
    10. Collier MP
    11. Chandler SA
    12. Tkachenko O
    13. Allen J
    14. Crispin M
    15. Billington N
    16. Takagi Y
    17. Sellers JR
    18. Eichmann C
    19. Selenko P
    20. Frey L
    21. Riek R
    22. Galpin MR
    23. Struwe WB
    24. Benesch JLP
    25. Kukura P

    (2018) Quantitative mass imaging of single biological macromolecules

    Science 360:423–427.

    https://doi.org/10.1126/science.aar5839

    • Google Scholar

Author details

1. Thomas RM Germe

   Department Biochemistry & Metabolism, John Innes Centre, Norwich Research Park, Norwich, United Kingdom

   Present address

   Dept. of Biochemistry and Molecular Genetics, University of Colorado Anschutz, Aurora, United States

   Contribution

   Conceptualization, Formal analysis, Supervision, Validation, Investigation, Visualization, Methodology, Writing – original draft, Project administration, Writing – review and editing

   For correspondence

   thomas.germe@cuanschutz.edu

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-6885-0971

2. Natassja G Bush

   Department Biochemistry & Metabolism, John Innes Centre, Norwich Research Park, Norwich, United Kingdom

   Present address

   Inspiralis Ltd., Innovation Centre, Norwich Research Park, Norwich, United Kingdom

   Contribution

   Investigation, Methodology, Writing – review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-3479-1261

3. Victoria M Baskerville

   Department Biochemistry & Metabolism, John Innes Centre, Norwich Research Park, Norwich, United Kingdom

   Contribution

   Visualization, Methodology

   Competing interests

   No competing interests declared

4. Dominik Saman

   Department of Chemistry, Biochemistry Building, University of Oxford, Oxford, United Kingdom

   Contribution

   Investigation, Visualization, Methodology

   Competing interests

   No competing interests declared

5. Justin LP Benesch

   Department of Chemistry, Biochemistry Building, University of Oxford, Oxford, United Kingdom

   Contribution

   Funding acquisition, Methodology

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-1507-3742

6. Anthony Maxwell

   1. Department Biochemistry & Metabolism, John Innes Centre, Norwich Research Park, Norwich, United Kingdom
   2. Department of Molecular Microbiology, John Innes Centre, Norwich Research Park, Norwich, United Kingdom

   Contribution

   Conceptualization, Supervision, Funding acquisition, Investigation, Methodology, Writing – review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-5756-6430

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