Structural Dynamics and MechanochemicalCoupling in DNA Gyrase
, Angelica C. Parente2, 3 an
d Zev Bryant 3, 4
1 - Department of Physics, University of Illinois at Urbana-Champaign,Urbana, IL 61801, USA2 - Program in Biophysics, Stanford University, Stanford, CA 94305, USA3 - Department of Bioengineering, Stanford University, Stanford, CA 94305, USA4 - Department of Structural Biology, Stanford University Medical Center, Stanford, CA 94305, USA
Correspondence to Zev Bryant: Department of Bioengineering, Stanford, University, Stanford, CA, 94305, [email protected]://dx.doi.org/10.1016/j.jmb.2016.03.016Edited by James Berger
Abstract
Gyrase is a molecular motor that harnesses the free energy of ATP hydrolysis to perform mechanical work onDNA. The enzyme specifically introduces negative supercoiling in a process that must coordinate fuelconsumption with DNA cleavage and religation and with numerous conformational changes in both the proteinand DNA components of a large nucleoprotein complex. Here we present a current understanding ofmechanochemical coupling in this essential molecular machine, with a focus on recent diverse biophysicalapproaches that have revealed details of molecular architectures, new conformational intermediates,structural transitions modulated by ATP binding, and the influence of mechanics on motor function. Recentsingle-molecule assays have also illuminated the reciprocal relationships between supercoiling andtranscription, an illustration of mechanical interactions between gyrase and other molecular machines atthe heart of chromosomal biology.
DNA gyrase remodels the bacterial chromosomeby introducing negative supercoils, playing anessential role in compacting the genome and solvingtopological challenges associated with replicationand transcription [1]. The enzyme forms a dynamiccomplex with N100 bp of DNA, and must form andbreak protein-DNA interactions and rearrange thesharply bent DNA path during each ATP-fueledconformational cycle. As has been extensivelyreviewed elsewhere [2,3], many structural andbiochemical features of the gyrase mechanismhave been identified over the past several decadessince the discovery of the enzyme. However, aconcrete understanding of the structure and dynam-ics of the complete holoenzyme complex has beenelusive because of the size of the machine, theextent of its interactions with the DNA substrate, andthe number of moving parts. Over the past five years,biophysical measurements have provided newinsight into conformational states and kinetic path-ways that underlie the mechanochemical function of
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the gyrase motor. We discuss these recent results,related experiments, and remaining questions afterbriefly introducing some biochemical and structuralbackground.
DNA gyrase is a specialized type IItopoisomerase
Gyrase is an A2B2 tetramer (Fig 1a) that shares acore set of domains and a duplex strand passagemechanism with other members of the type IIAtopoisomerase family [3]. In this family, threeprotein–protein interfaces form gates that can openand close to allow passage of DNA through theenzyme. The segment of duplex DNA destined to becleaved, called the G- (or Gate-) segment, binds atthe central DNA gate formed by the Topoisomerase-Primase (TOPRIM) domain together with theWinged Helix Domain (WHD) containing the activesite tyrosine that forms a transient 5′-phosphotyrosyl
Fig. 1. Composition and basic mechanism of DNA gyrase. (a) Cartoon showing domain organization. Gyrase is an A2B2heterotetramer. Interfaces between the subunits form three gates that can be opened and closed. (b) Outline of theenzymatic cycle. The G-segment binds to the central DNA gate. Chiral wrapping presents a proximal T-segment within theN-gate cavity. ATP binding induces N-gate closure, followed by passage of the T-segment through the transiently cleavedG-segment and expulsion through the C-gate. One round of strand passage leads to the introduction of two negativesupercoils.
1834 Review: Dynamics and Coupling in DNA Gyrase
protein-DNA linkage in the cleaved state. DNA-bound structures of the binding and cleavage corefor gyrase and other type IIA topoisomerases showa dramatic bend in theG-segment [4,5]. A secondDNAsegment known as the T- (or transfer) segment entersthrough theN-gate, in an upper cavity formedbyGHKLATPase domains that can dimerize upon ATP bindingandmay communicate the nucleotide state to theDNAgate via conformational changes involving the inter-vening transducer domain [6,7]. After passing throughthe transiently cleaved G-segment and opened DNAgate into a lower cavity framed by coiled-coil domains,the T-segment can exit through a final reversibleinterface known as the C-gate. In a closed circularmolecule, the overall reaction inverts a node betweenthe T- and G-segments and therefore changes thelinking number of the DNA in a step of 2 [8].The globular C-terminal domain (CTD) of DNA
gyrase (Fig, 1a) diverges from other type IIAtopoisomerases [9] and is essential for the uniqueability of DNA gyrase to introduce, rather than merelyrelax, supercoils (Fig. 1b). The directionality of super-coiling can be enforced by chirally wrapping DNAbetween the G-segment and the T-segment, trapping(+) writhe and presenting a (+) node whose inversionchanges the linking number by −2 [8,10]. GyrasewrapsDNAasprobedby exonuclease, DNAase I, andhydroxyl radical footprinting experiments [11–13], and
constrains (+) writhe in the absence of nucleotide asshownby topoisomer footprinting [10] [14]. TheCTD isessential for these properties and for directionalsupercoiling; its deletion converts gyrase into aconventional type II topoisomerase [15,16]. Structuraland functional studies of the isolated CTD [17] [18]showed that is has a beta-pinwheel fold with a basicpatch around the outer edge, and can bind and bendDNA.
Structural investigations of globalarchitecture
As of this writing, there is no reported high-resolution structure of full-length gyrase in complexwith DNA. The arrangement of the CTDs relative tothe core and the path of the DNA around the CTDshave been the subject of study and debate. A crystalstructure of the related topoisomerase IV [9] showedthe CTDs in plane with the DNA gate, and smallangle X-ray scattering models have shown the CTDsin a lower position near the exit gate in GyrA [19] andthe holoenzyme [20], leading to proposals in whichthe CTDs are mobile during the conformational cycle[19]. An important goal for describing the DNAgyrase mechanism is to define conformations ofthe overall holoenzyme and the associated DNA
1835Review: Dynamics and Coupling in DNA Gyrase
visited during the mechanochemical cycle, andcharacterize transitions in this global architecturedependent on substeps in fuel consumption.A recent cryoelectron microscopy study [21]
provides the most complete picture to date of thearchitecture of full-length DNA gyrase, and the firstdirect visualization of DNA wrapped around theCTDs (Fig. 2a). A 23 Å map of the T. thermophilusgyrase holoenzyme was obtained in complex with155 bp DNA, stabilized with ciprofloxacin andAMPPNP. The N-gate is closed and the CTDs arein plane with the DNA gate, with clear densityattributable to DNA bent around them. The confor-mation was proposed to represent a state that trapsa T-segment prior to strand passage, although theincluded DNA appears to be of insufficient length to
Fig. 2. Recent structures illuminate the architectures of gyrmap of the T. thermophilus gyrase holoenzyme in complex wfrom [21]). The domain architecture can be seen together witCrystal structures of protein components and modeled DNA duis shown in a domain-swapped configuration first observed inenzyme S. cerevisiae topo II in complex with G-segment DNAalso seen in closed configurations in these structures.
present a T-segment in the complex that wasobtained. The modeled DNA (Fig. 2a) includes ashallower G-segment bend than has been seen incrystal structures of various type IIA topoisomerasesin complex with DNA (Fig. 2b), including structures ofthe gyrase cleavage core bound to shorter DNAfragments [4, 5], suggesting an influence of theCTDs on the central DNA conformation. The authorsalso obtained a 17 Å reconstruction of the holoen-zyme in the absence of DNA, in which the CTDs arenot visible due to conformational heterogeneity. Forboth reconstructions, the closed N-gate is seen in a“domain swapped” configuration previously ob-served [22] in a recent structure of the full lengthS. cerevisiae topoisomerase II in complex withG-segment DNA and AMPPNP (Fig. 2b), which was
ase and related type II topoisomerases. (a) 23 Å CryoEMith 155 bp DNA, ciprofloxacin, and AMPPNP (reproducedh density attributable to DNA wrapped around the CTDs.plex (green) have been fit to the density. The closed N-gate(b) a crystal structure (reproduced from [22]) of the related(green) and AMPPNP. The DNA-gate and the C-gate are
1836 Review: Dynamics and Coupling in DNA Gyrase
proposed to represent a post-strand passage con-formation that prevents regression of the T-segmentback up through the DNA gate.Unlike crystallography and cryoelectron microsco-
py, single-molecule measurements are able to probeonly limited structural degrees of freedom. Thesemethods are nevertheless powerful tools for charac-terizing dynamic molecular machines because theycan be used on heterogenous populations insolution, report on conformational distributions,directly follow the dynamics of actively cyclingenzymes, and apply controlled mechanical pertur-bations. Three major single-molecule approacheshave been applied to DNA gyrase, each exploitingone or more of the attributes above: (1) FRETmeasurements have been used to characterizeconformational distributions under varying condi-tions relevant to the mechanochemical cycle, nota-bly contributing to our understanding of how proteindomains rearrange in response to DNA and nucle-otide binding; (2) a specialized magnetic tweezerstechnique known as rotor bead tracking (RBT) [23]has been used to directly follow the dynamics ofsupercoiling and nucleotide-dependent transitions
Fig. 3. Single-molecule FRET reveals DNA and nucleotid(a) Schematic of confocal smFRET microscopy (not to scale). Lvolume produce brief bursts of fluorescence that are collected oFRET efficiencies. (b) FRET labeling positions used for probingFRET histograms for gyrase using the S7C labeling position (rlabeled O (open), C (closed), and I (intermediate/narrowed). Fthan the intermediate state, explained by the N-terminal locat(c) No nucleotide (blue) vs ADPNP (black), in the absence of Dthe absence of nucleotide. (e) Cartoons of N-gate conformationsmFRETmeasurements between the gyrA CTD and the core egyrA dimer alone, move out slightly when gyrB is bound, and
between states that differ in their DNA conformation;and (3) “conventional” magnetic tweezers assays[24] have been used to probe the effects of force,torque, and DNA mechanics on the gyrase molec-ular motor.
FRET measurements of conformationalstates
Single-molecule fluorescence resonance energytransfer (smFRET) can provide a measurement ofthe distance between two dyes attached at knownpositions to a molecule of interest. Gyrase confor-mations have been extensively studied [25–31]using an smFRET experimental design in whichmolecules are observed when they diffuse through aconfocal volume [32], producing a brief burst offluorescence (Fig. 3). This smFRET approach allowsdistributions of FRET values to be recorded overmany molecules while avoiding the complicationsof surface interactions. The goal of the assay isprincipally to capture snapshots rather than dynam-ics, since the observation time for each molecule is
e dependent conformations of B. subtilis DNA gyrase.abeled complexes diffusing through the femtoliter confocaln donor and acceptor channels to measure distributions ofN-gate conformations (reproduced from [27]). (c-d) N-gateeproduced from [27]), showing three N-gate conformationsor this labeling position, FRET is lower in the closed stateion of S7C in the intertwined dimerized ATPase domains.NA. (d) No DNA (black) vs relaxed plasmid DNA (blue), ins probed by FRET. (f) Cartoons of CTD positions based onnzyme [30]. CTDs are positioned toward the exit gate in theswing up when the enzyme is complexed with DNA.
1837Review: Dynamics and Coupling in DNA Gyrase
short in comparison with the timescale of themechanochemical cycle. This may be contrastedwith the longer observation times enabled by acommon alternative approach [33] in which mole-cules are affixed to a coverslip and typicallyilluminated using total internal reflection (TIRF).smFRET measurements on B. subtilis gyrase have
shown how both the N-gate conformation and theCTD position depend on the nucleotide and the DNA.These studies have revealed unanticipated confor-mational states, and haveprovided snapshots that aresuggestive of a series of conformational changesleading to directional strand passage. To monitor theN-gate, Gubaev et al. [27] produced gyrB-gyrA fusionproteins labeled at each of several alternativepositions on the ATPase domain, and measuredFRET between equivalent positions on symmetricdimers (Fig. 3b). As expected, FRET values areconsistent with an open N-gate in the apo state,shifting to a closed state in presence of AMPPNP(Fig. 3c). Unexpectedly, the authors discovered anintermediate (narrowed) conformation of the N-gatethat is populated when DNA is bound even in theabsence of nucleotide (Fig. 3d). Formation of theintermediate state requires N110 bp of DNA, sug-gesting that the narrowed gate depends on DNAwrapping around the CTDs. A chiral wrapping modelfor this state is supported by the observation of areduced population of the intermediate state in thepresence of negative DNA supercoiling, which isexpected to oppose positive superhelical wrapping.The results were interpreted to suggest an orderedprogression that coordinates DNA wrapping withN-gate closure, in which the N-gate partially closeswhen the chiral wrap is formed, then closes complete-ly upon ATP binding to trap a T-segment poised forstrand passage (Fig. 3e).For directly measuring CTD movements, hetero-
dimeric gyrA mutants were purified to enableintramolecular labeling with a donor/acceptor dyepair on the CTD and body of a single gyrA subunit[30]. By measuring differences in FRET distributionswhen exposed to gyrB and different lengths of DNA,the authors found that the CTDs are positionedtoward the exit gate in the gyrA dimer, move slightlyfurther away from the body in the gyrA:gyrBholoenzyme, and move upward when DNA isbound (Fig. 3f). The inferred change in CTD positionbetween the gyrA dimer and the DNA-boundholoenzyme is qualitatively similar to a comparisonbetween the earlier SAXS structure for gyrA [19] andthe T. thermophilus cryoEM structure that appearedafter these FRET studies [34]; the results with DNAcontrast with the SAXSmodel obtained for the E. coliholoenzyme [20]. smFRET histograms for someCTD-body labeling locations were bimodal, suggest-ing there may be either structural asymmetrybetween the two CTDs or heterogeneity of a singleCTD position. Repositioning the CTDs requires
neither nucleotide, nor cleavage, nor a long enoughDNA template to present a T-segment, suggestingan ordered progression in which CTD movement isan early step in the catalytic cycle, positioning theCTD for chiral wrapping to present a T-segment inthe narrowed N-gate followed by N-gate closure.Solution confocal smFRET measurements can
provide richly detailed information about conforma-tions accessible to DNA gyrase. The use of multipleFRET pair positions places constraints on the 3Darchitectures of complexes that have resistedconventional structural characterization, and ex-haustive sampling of conditions has helped deter-mine coupling between protein conformations andchemical states relevant to the mechanochemicalcycle of the enzyme. However, because the assaydoes not provide either dynamic tracking of confor-mations over the timescale of the cycle or a readoutof progress of the supercoiling reaction, the.temporal ordering, kinetics, and functional context
of conformational states cannot be directly deter-mined. Rotor bead tracking (RBT), which measuresreal-time changes in the extension and linkingnumber of a single tethered DNA molecule [23], isa complementary technique that has been exploitedto directly monitor the mechanical output of thegyrase supercoiling reaction while tracking rear-rangements of the nucleoprotein complex manifest-ed as dynamic changes in the conformation of theDNA (Fig. 4).
RBT measurements of nucleoproteindynamics
RBT relies on measuring the angular position of asubmicron bead (the “rotor”) attached to the side of asingle stretched DNA molecule (Fig. 4a). In assays ofDNA gyrase, the introduction of two negative super-coils causes the rotor to spin by two full rotations perenzymatic cycle, while a distal swivel prevents thepermanent accumulation of torsional strain. Structuralintermediates within the cycle appear as angulardwells that can be placed along the natural repeatingtwo-rotation reaction coordinate. Initial RBTmeasure-ments of DNA gyrase showed processive bursts instrict multiples of two rotations (similar to Fig. 4b,above) as expected [35], and found that the rate-limiting angular dwell occurs at the ~0 (even) rotationmark, implying that the enzymepredominantly waits ina state that does not trap any writhe. This state wasinitially assumed to have released the wrapped DNA,but later RBT measurements [36] — which includedtracking of the rotor height (z) as a direct measure ofDNA sequestration (Fig. 4b, below) for the first time—showed that although no writhe is trapped, theenzyme in fact sequesters N100 bp of DNA in thisunanticipated nucleoprotein conformation nowdubbed the Ω state.
1838 Review: Dynamics and Coupling in DNA Gyrase
The properties of the Ω state are explained by aschematic model in which the DNA flanking theG-segment is bent around the CTDs without forminga chiral wrap [36]. A major remodeling transition isrequired in order to reach a chirally wrapped
Fig. 4 (legend o
configuration, now dubbed the α state. Both ofthese structural intermediates may be observed inRBT experiments and visualized as populations intwo dimensional histograms of (angle, z) coordinatesvisited during active supercoiling (Fig. 4c). The
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1839Review: Dynamics and Coupling in DNA Gyrase
Ω-to-α transition, which dominates the kinetics ofsupercoiling, was proposed to involve CTD motionand correspond to T-segment capture. ATP dramat-ically accelerates the rate of this remodeling transi-tion, revealing a new role for nucleotide binding inpromoting the formation of the chiral wrap. Quanti-tative analysis of [ATP]-dependent substep kinetics[36] led to a branched kinetic model for early eventsin the gyrase cycle (Fig. 4e): the central Ω-to-αremodeling transition can occur slowly via thermalsampling in the absence of ATP, or quickly when2 ATP molecules are bound. The α state is chirallywrapped and contains a poised T-segment; it requires2 boundATPmolecules to progress forward via strandtransfer, and can otherwise thermally revert to the Ωstate. Notably, thermal interconversion of Ω and αquantitatively explains bulk topoisomer footprintingassays, in which ~0.8 positive supercoils were foundto be trapped per gyrase enzyme in the absence ofnucleotide [10,14]. RBT, a single-molecule analog oftopoisomer footprinting, recapitulates a similar valueas an average over three dynamically exchangingconformations: theΩ state, which traps ~0 supercoils,and two isoforms of the α state in which either ~1 or~1.7 supercoils are trapped. This result illustrates thepower of single-molecule measurements to resolveheterogeneous populations [36], which should beconsidered when interpreting other bulk measure-ments of the complex such as DNA protection studies[11–13] that may similarly reflect averages overconformational ensembles.RBT provides an incomplete picture of nucleopro-
tein dynamics because of limited degrees of freedom(only the angle and extension of the DNA aremonitored) and also because of finite spatiotemporalresolution: Brownian noise of the rotor obscures thedetection of short-lived states or small conforma-tional changes [23]. An important advance in RBTtechnology was therefore the introduction of AuRBT,which uses evanescent darkfield imaging to track
Fig. 4. Rotor bead tracking reveals new conformations androtor bead tracking (RBT) assay. DNA is stretched using a maside of the molecule and tracked using fluorescence [35] [36] ochanges in DNA angle and extension (z) in real time. (b) RBT trunder in 1 mM ATP (above) or 75 μM ATP (below). Individucorresponding to processive negative supercoiling. [ATP]-depean intermediate angle (*) corresponding to a chirally wrappedpaired (angle,z) values during gyrase activity in presence of 75the enzyme. Angles are shown modulo 2 rotations. The Ω stamark, which is explained by sequestering DNA contour withourotation mark, corresponding to trapping (+) writhe prior to stranATP using gold rotor bead tracking (reproduced from [37]). A sindwells are interrupted by brief excursions to a state (*) that relefor structural transitions and ATP coupling in DNA gyrase [36,37the transition from Ω to α, which can occur slowly and reversiblSubsequent strand passage also requires the presence of 2recaptured to begin a new round of supercoiling.
gold nanoparticles employed as high-speed probesof DNA angle and extension [37], offering dramaticimprovements over previous RBT methods. In aninitial application of AuRBT to DNA gyrase (Fig. 4d),not only are individual steps between dominantdwells in the Ω state very clearly visualized even atsaturating [ATP], but a new transient state betweenthese dwells can also be seen for the first time [37].In this “ν state”, substantial DNA contour length isreleased from the enzyme, leading to a model inwhich DNA is briefly released from one or both CTDsafter strand passage, and then recaptured to beginthe next cycle. In the picture that emerges from RBTand AuRBT measurements (Fig. 4e), the formationof the chiral wrap during each cycle is a multistepprocess: beginning from the ν state, DNA is firstquickly bent around the CTDs to form the Ω state,then goes through a slower rearrangement relyingon CTD motion to reach the chirally wrapped α state.While RBT measurements have illuminated major
global remodeling transitions in the gyrase:DNAcomplex, more work is needed to establish themolecular details of these structural transitions. TheDNA path for each state has only been depicted inschematic cartoons; RBT measurements are insuf-ficient to precisely define this path, and states mayalso contain variable structures, substates, andunresolved intermediates that contribute to thespread observed in angle and z values (Fig. 4c).The model for the DNA path presented in the recentcryoEM study resembles cartoons of the ~1.7supercoil-trapping isoform of the α state, but shouldbe interpreted with caution since (1) there is no directvisualization of a trapped T-segment; (2) thecomplex is stabilized using a nucleotide and drugcombination in a state with an unknown relationshipto the functional conformational cycle; and (3) theT. thermophilus enzyme used for cryoEM may havedistinct properties from the distantly related E. colienzyme used in RBT measurements.
ATP-dependent dynamics of E. coli DNA gyrase. (a) Thegnetic bead, and a submicron rotor bead is attached to ther evanescent scattering [37] videomicroscopy to measureaces (reproduced from [36]) in the presence of DNA gyraseal gyrase encounters lead to bursts of stepwise rotation,ndent dwells are seen at the even rotation mark and also atintermediate. (c) 2D histogram (reproduced from [36]) ofμM ATP, showing distinct conformational states visited byte is significantly contracted in z but lies at the ~0 rotationt trapping writhe. The α state can also be seen at the ~1d passage. (d) High-resolution dynamics of gyrase at 1 mMgle processive burst is shown in the (angle, z) plane. Majorases significant contour length. (e) Branched kinetic model]. The kinetics of processive supercoiling are dominated byy in the absence of ATP or quickly when 2 ATP are bound.ATP. DNA is partially released after strand passage and
1840 Review: Dynamics and Coupling in DNA Gyrase
Common themes and outstandingquestions fromsingle-molecule dynamics
Descriptions of gyrase structural dynamics inferredfrom smFRET and RBT studies have some similarfeatures, includingmobile CTDs, ordered progressionsof conformational intermediates leading to chiralwrapping and T-segment capture, and coupling be-tween the conformations of the DNA and the N-gate.However, it is unclear that any one-to-onemapping canbe made between protein conformational statesidentified using FRET and DNA conformational statesidentified using RBT. It was suggested [36] that theATP-free chirally wrapped α state contains a narrowedN-gate,whichmight partially trap aT-segment and helpexplain long dwells in this state in RBT experimentsas well as stabilization of the narrowed N-gate byconditions favoring chiral wrapping in FRET experi-ments [27]. In this model, the narrowed N-gate favorsproductive strand passage by a creating a binding siteto retain the T-segment awaiting complete N-gateclosure and strand passage.The Ω conformation has also been proposed to
inhibit N-gate closing [36], helping gyrase avoid futilecycles of ATP hydrolysis before a T-segment isbound. In this model, the closed N-gate occurs onlytransiently during active supercoiling, which agreeswith smFRET measurements using hydrolysableATP [27]. The Ω conformation has been also beendepicted with upwardly positioned CTDs [36], butdefinitive determination of the correspondence be-tween DNA and protein conformations awaitsexperiments that measure these degrees of freedomsimultaneously, as might be accomplished using amultimodal single-molecule approach [38–40] sucha proposed combination of AuRBT with FRET [37].Comparisons between the confocal smFRET and
RBT measurements are further complicated by theuse in these two experiments of gyrase from twodivergent species (B. subtilis and E coli, respectively),which may have differences in the proportion of thecycle spent in different substates, or possibly moredramatic mechanistic differences. Significant differ-ences are hinted at by the apparent failure ofB. subtilisgyrase to perform a single round of strand passagesupported by AMPPPNP, in contrast to E. coli [31,41]and by differing functional requirements for theC-terminal tail of the CTD, which has been implicatedas a physical element responsible for coordinationbetween chiral wrapping and theATPase cycle in bothspecies and is absolutely required for supercoiling inE. coli [42] but not in B. subtilis [29]. Directcomparisons between the two species in identicalsingle-molecule assays will be important for distin-guishing general from specific features of the gyrasemechanochemical cycle.A strength of the RBT studies is that they provide
kinetic as well as structural information, yielding a
quantitative dynamic description of the motor cyclein which structural transitions are coupled to specificsubsteps in fuel consumption. However, work to datehas left this mechanochemical description of themotor mechanism substantially incomplete: the νstate has not been characterized in enough detail toeither define its geometry or determine whether itsdynamics depend on the nucleotide cycle, andglobal conformational changes coupled to hydrolysisand product release have not been directly probed.Future RBT experiments conducted with varyingnucleotides, including non-hydrolysable analogs,may address these questions and provide furthervaluable points of comparison with bulk solutionmeasurements. Hydrolysis has previously beenstudied using bulk single-turnover kinetics in therelated enzyme yeast topoisomerase II, and it wasfound that ATP hydrolysis dramatically acceleratesstrand passage [43], although it has long beenknown that hydrolysis is not strictly required for thisstep [44]. If this turns out to be true for gyrase as well,it will complete a repeating pattern of loosely coupledstructural transitions, in which the Ω-to-α chiralwrapping transition can happen slowly without ATPor quickly when ATP is bound, presenting aT-segment that can then be transferred slowly withATP binding alone or quickly when ATP is hydro-lyzed. Chemical substeps thus modulate the confor-mational energy landscape to guide the motortoward productive forward progress, without requir-ing a one-to-one correspondence between chemicaland conformational states.
Mechanics of gyrase and its interactionsin the chromosome
Mechanical perturbations applied in single-mole-cule experiments are valuable for probing the energylandscapes of molecular motors [45] and for testingresponses to stresses that may be experienced incells. The first RBTmeasurements of DNAgyrase [35]found that the processivity of the motor is exquisitelysensitive to tension in the DNA molecule, while thesupercoiling velocity of the motor is relatively insen-sitive to this parameter. In light of subsequent work[36], this behavior may be understood since dissoci-ation involves a large change in DNA extension whenthe enzyme releases sequestered contour length,while the rate-limiting step in the supercoiling cycleinvolves a transition between two states (Ω and α) thatboth sequester extensive contour length and thushave similar extensions. Transitions to and from thetransient ν state [37] are expected to be highlytension-sensitive due to the large changes in exten-sion relative to Ω and α, but this perturbation has notbeen characterized directly and would not be expect-ed to affect the supercoiling velocity under moderatetensions. The major influence of even sub-pN forces
1841Review: Dynamics and Coupling in DNA Gyrase
on processivity, and potentially on the dynamics oftransient states, could be a control mechanism in vivoand should also be accounted for when comparingsingle-molecule measurements under tension to bulksolution measurements.Torque may be a more important parameter than
tension in cellular contexts, where the enzyme mustwork against accumulated negative supercoiling ormay act on transiently positively supercoiled do-mains. In the RBT assays used for gyrase, super-coils do not accumulate, but alternative assays havebeen used to probe this condition. Nollmann et al.measured gyrase activity on both positively andnegatively supercoiled DNA molecules [46], using amagnetic tweezers assay in which changes in linkingnumber are reflected in changes in DNA extensiondue to the accumulation of plectonemic structures.They noted robust relaxation of positive supercoilseven under elevated tensions, and also observedmechanically induced switching between introduc-tion and relaxation of negative supercoils. Similarresults were obtained by Fernandez-Sierra et al.[47], who also studied the activity of gyrase ondiaminopurine-substituted tethers, which have ahigher bending stiffness than unsubstituted DNA.E. coli gyrase essentially fails to supercoil diamino-purine-substituted DNA, which was ascribed to theadditional energy required to achieve the very tightlybent conformations seen in both the Ω and α states.Sequence-dependent modulation of DNA bendingstiffness was proposed as a potential mechanism oflocalized biological control over gyrase activity [47].The mechanics of DNA gyrase must be under-
stood in a larger context: gyrase communicates withother cellular machinery through torsional strain inthe DNA. It has long been appreciated that torsiongenerated by gyrase is used to transmit informationthrough the genome and exert sophisticated controlover biological processes such as replication initia-tion and transcription of specific genes, includinghomeostatic control of gyrase itself [48] and tran-scriptional responses to metabolic changes that maybe sensed directly by the DNA gyrase motor viathe cellular ATP energy stores [49,50]. In a recentstriking example [51], oscillating DNA supercoilinglevels act asaglobal regulator of shifting transcriptionalprograms during the circadian rhythms of cyanobac-teria: distinct promoters are simultaneously up- anddown-regulated by torsional changes, and inhibition ofDNA gyrase is sufficient to induce a transcriptionalresponse that mimics a change in the time of day.Recent single-molecule investigations have investi-gated the transcriptional side of this mechanicalinteraction, by measuring how RNA polymerasegenerates and responds to torque, and by directlyobserving the impact of gyrase on transcription.In seminal work, Liu and Wang noted that progres-
sion of an elongating transcription complex cangenerate positive supercoils ahead of and negative
supercoils behind the polymerase [52] due to helicaltracking on a constrained DNA duplex. This phenom-enon may be expected to occur in anchored ~10 kbsupercoiling domains [53] in the bacterial chromo-some (Fig. 5a). To directly measure the effect ofaccumulated torque on transcription elongation, Maet al. used an angular optical trap (Fig. 5b) to followtranscription against a torsional load. Among theirobservations, they found that RNAP generatespositive supercoiling until it stalls at a characteristictorque of ~10 pNnm. In this experiment, negativesupercoils do not accumulate behind the polymerasedue to the presence of a free end. In a bacterial cell,positive andnegative supercoiling domains generatedby transcription may be relaxed by gyrase andtopoisomerase I, respectively, and a local imbalancebetween these topoisomerases could lead to netsupercoilingwithin a chromosomal loop. This scenariowas directly investigated [54] using a single-moleculeassay for transcription in which the growing nascentRNA produces an increasing fluorescence signal dueto binding of a dye (Fig. 5c). With this assay, Chong etal. were able to measure transcriptional activity ontethered DNA circles that mimic chromosomal loops,and observe the effect of including topoisomerase Iand gyrase in the system.Results on the mechanical interplay of gyrase,
topoisomerase I, and RNA polymerase supported thefeasibility of a model that may explain the phenome-non of transcriptional bursting in E. coli [54]. In thismodel, excess topoisomerase I continually relaxesnegative supercoils generated behind transcribingcomplexes, while the positive supercoils generatedahead of RNA polymerase are only relaxed whengyrase is present in the supercoiling domain (Fig. 5d).Noting that the number of gyrase holoenzymes in thecell [55] is of the same order as the number ofconstrained supercoiling domains [53], the modelpredicts that transcription within a ~ 10 kb domain willswitch bimodally between (i) active gene expressionwhen gyrase is present, and (ii) arrest when gyrase isabsent and positive supercoils accumulate to inhibittranscription. This study presents an example of acomplex emergent phenomenon that arises fromsimple interactions between molecular machines inthe chromosome, and hints at parallels with thecytoskeletal motor field, where theoretical and exper-imental studies have shown how collections of motorsand filaments may display emergent behaviorsdependent on microscopic properties of individualmotors such as force-velocity relationships andforce-dependent off rates [56].In order to further relate biophysical measurements
to in vivo functions, it will be critical to measure variedproperties of gyrases found in different organisms,where they may be adapted for a range of cellularrequirements. For example, M. tuberculosis gyrase,which lacks the C-terminal tail required for coordina-tion of chiral wrapping and ATP binding in E. coli [42],
Fig. 5. Mechanical interplay of gyrase, transcription, and DNA supercoiling investigated using single-moleculemethods. (a) Helical tracking of the advancing transcription complex leads to twin supercoiling domains in a constrainedDNA duplex [54,63]. (b) An optical torque wrench assay [64] showed that RNA polymerase stalls due to positive supercoilsthat accumulate ahead of the enzyme, with a measured stall torque of ~10 pN nm. (c) Single-molecule assay fortranscription on tethered constrained circular templates [54]. Fluorescence accumulates during transcription due to anRNA-binding dye. Dynamics can be investigated in the presence of gyrase and/or topoisomerase I. (d) Model fortranscriptional bursting based on single-molecule measurements [54]. Topoisomerase I constitutively relieves (−)supercoils behind the transcription complex, leading to the accumulation of (+) supercoils in a constrained chromosomalloop. When gyrase is bound, (+) supercoils are relaxed and transcription can proceed. When gyrase dissociates,accumulated (+) supercoils inhibit transcription, intermittently shutting off gene expression.
1842 Review: Dynamics and Coupling in DNA Gyrase
acts slowly and stalls at much lower supercoilingdensities than the E. coli enzyme [57]. Even closelyrelated bacterial species can have differing super-coiling requirements: Salmonella enterica serovarTyphimurium supercoils its genome to a lower densitythan E. coli [58]. Few single-molecule measurementshave been repeated on more than one species — acomparison between E. coli and Salmonella reportedby Fernandez-Sierra et al. is a rare exception [47] —andmeasurements that control for species differencesare needed for making consistent biophysical modelsthat integrate data from structural, biochemical, fluo-rescence, and mechanical experiments. As we havenoted, the cryoEM, smFRET, and RBT experimentsreviewedherewereperformedusing gyrase from threedivergent organisms (T. thermophilus, B. subtilis, andE. coli, respectively), complicating comparisons.An additional source of varied gyrase behavior is
the DNA binding site. Biophysical measurementshave exploited sequences that form unusually tight
complexes with DNA gyrase. RBT measurements,for example, have made use of a variant of the stronggyrase site from Mu phage [59], in order to increasethe processivity of the enzyme [60] and counteractthe destabilizing effect of tension. Comparativemeasurements on diverse sequences, includinggyrase binding sites of biological interest such asREP sequences identified in the E. coli chromosome[61,62], will be valuable for generalizing results andrelating mechanochemistry to biology.
Toward a mechanochemical descriptionof gyrase motor function
Recent biophysical studies havebuilt upondecadesof biochemical and structural investigations to showhow coordinated conformational changes in thegyrase nucleoprotein complex lead to motor function.Single-molecule measurements have begun to reveal
1843Review: Dynamics and Coupling in DNA Gyrase
the complexity of a branched kinetic pathway in whichstructural transitions are loosely coupled to chemicalsubsteps, and more work is needed to fully define themechanochemical cycle. Models that relate proteinconformational changes to the dynamics of DNAgeometry must be tested, and a major challenge forstructural biology is to establish the detailed three-di-mensional architectures of conformational statesidentified in single-molecule studies, including the Ω,α, and ν states. Finally, the mechanical capabilitiesand responses of gyrase and other DNA-associatedmachines must be understood and may be tested incombinations [54] in order to build a quantitativeunderstanding of an emerging mechanobiology of thechromosome.
Acknowledgements
This work was supported by NIH grants GM106159and 5U19AI10966202.
Received 12 November 2015;Received in revised form 16 February 2016;
Accepted 15 March 2016Available online 22 March 2016
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