The processing of repetitive extragenic palindromes: the structure of ...

The processing of repetitive extragenic palindromes:the structure of a repetitive extragenic palindromebound to its associated nucleaseSimon A. J. Messing1, Bao Ton-Hoang2, Alison B. Hickman1, Andrew J. McCubbin3,

Graham F. Peaslee3, Rodolfo Ghirlando1, Michael Chandler2 and Fred Dyda1,*

1Laboratory of Molecular Biology, National Institute of Diabetes and Digestive and Kidney Diseases, NationalInstitutes of Health, Bethesda, MD 20892, USA, 2Laboratoire de Microbiologie et Genetique MoleculairesCentre National de la Recherche Scientifique, 118 Route de Narbonne, 31062, Toulouse Cedex, France and3Chemistry Department, Hope College, 35 E. 12th Street, Holland, MI 49423, USA

Received May 1, 2012; Revised June 20, 2012; Accepted July 11, 2012

ABSTRACT

Extragenic sequences in genomes, such asmicroRNA and CRISPR, are vital players in the cell.Repetitive extragenic palindromic sequences (REPs)are a class of extragenic sequences, which form nu-cleotide stem-loop structures. REPs are found inmany bacterial species at a high copy number andare important in regulation of certain bacterial func-tions, such as Integration Host Factor recruitmentand mRNA turnover. Although a new clade ofputative transposases (RAYTs or TnpAREP) is oftenassociated with an increase in these repeats, it isnot clear how these proteins might have directedamplification of REPs. We report here the structureto 2.6 A of TnpAREP from Escherichia coli MG1655bound to a REP. Sequence analysis showed thatTnpAREP is highly related to the IS200/IS605 family,but in contrast to IS200/IS605 transposases,TnpAREP is a monomer, is auto-inhibited and isactive only in manganese. These features suggestthat, relative to IS200/IS605 transposases, it hasevolved a different mechanism for the movementof discrete segments of DNA and has beenseverely down-regulated, perhaps to prevent REPsfrom sweeping through genomes.

INTRODUCTION

For many years, a large part of the extragenic sequencein genomes was thought to be essentially silent and devoidof function, hence the popular term ‘junk DNA’. Thisparadigm changed considerably over the last two

decades, as it has become apparent that these sequencesoften encode unexpected functions as evidenced by thediscovery of microRNAs and their role in gene regulation(1,2), and by the identification of a new class of shortpalindromic repeats, known as Clustered Regularly Inter-spaced Short Palindromic Repeats (CRISPR), which arecritical in prokaryotic immunity (3–5).

Repetitive extragenic palindromic sequences (REPs) area distinct class of abundant repeats important in regula-tion of certain bacterial functions. REPs are known tointeract with several partners, by providing binding sitesfor proteins such as Integration Host Factor and DNApolymerase I, and providing the necessary cleavage sitesfor DNA gyrase to unwind DNA (6–8). REPs alsoincrease mRNA stability and can cause transcriptiontermination (9,10). These REP functions are also exhibitedto some degree in other extragenic prokaryoticDNA elements, such as the Correia elements in Neisseriagonorrhoeae, or RUP elements in Streptococcuspneumoniae (11–14). Discovered nearly 30 years ago inenteric bacteria (15,16), REPs are 35–40 nt longstem-loop structures often organized into larger unitscalled bacterial interspersed mosaic elements (BIMEs)(17–19). BIMEs comprise two REPs in inverse orientationseparated by a linker sequence (shown in Figure 1a forEscherichia coli MG1655). One is called REP and thesecond inverted sequence is designated an iREP. REPsare found dispersed throughout the chromosome inmany bacterial species, often in high copy number. Theyrepresent, for example, up to 1% of E. coli chromosomes(20–24). They are so frequent that their presence serves asthe basis for ‘REP-PCR’, a method for the rapid identifi-cation of bacterial strains (25).

Hints as to how REPs have come to populate somebacterial genomes with such high frequency were

*To whom correspondence should be addressed. Tel: +301 402 4496; Fax: +301 496 0201; Email: [email protected]

9964–9979 Nucleic Acids Research, 2012, Vol. 40, No. 19 Published online 9 August 2012doi:10.1093/nar/gks741

Published by Oxford University Press 2012.This is an Open Access article distributed under the terms of the Creative Commons Attribution Non-Commercial License (http://creativecommons.org/licenses/by-nc/3.0), which permits unrestricted non-commercial use, distribution, and reproduction in any medium, provided the original work is properly cited.

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obtained from analyses of the genetic regions aroundREPs, which revealed a new clade of putative trans-posases, termed REP-associated tyrosine transposases(RAYTs) or TnpAREP (26,27). RAYTs are found in avariety of species at a species-specific single locus, alwaysflanked by BIMEs. Phylogenetic analysis of E. coli andShigella indicated RAYTs were acquired early in speciesradiation (27). In principle, as transposases can cleave and

recombine DNA segments, this could explain the spreadof REPs in their respective genomes. Consistent with thishypothesis, RAYT presence is correlated with a generalincrease in REP copy number (26). In addition, recentin vitro studies on the TnpAREP from E. coli MG1655showed that it can cleave specific DNA segments if aREP is present and is also able to recombine BIME frag-ments (27). Furthermore, evidence that REPs may be

(a)

(b)

Figure 1. REP organization and IS200/IS605 family alignment. (a) Schematic representation of the E. coli MG1655 REPtron displaying the organ-ization of BIMEs and their respective REPs and iREPs. The y REP is in red, the y iREP in orange, z1 REP in blue and the z1 iREP in light blue.The y and z1 nomenclature preceding REP or iREP define the consensus nucleotide sequence of the hairpin (27). The 50 GTAG tetranucleotide isrepresented by a purple box at the foot of the REP, and the CT dinucleotide cleavage sites previously established in reference 27 are marked by redarrows. (b) Multiple sequence alignment of prominent members of the IS200/IS605 family with key members of the new clade. The histidines of theHUH motif are highlighted in orange, and the catalytic tyrosine in magenta. Other residues coordinating the divalent cation in the new clade are inorange script, residues involved in TnpAIS608 dimerization in cyan, C-terminal extension of TnpAREP in purple and residues sharing identitythroughout are highlighted in red boxes.

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capable of excision was obtained from Pseudomonasfluorescens (28).Sequence analysis of the RAYT clade reveals that its

members are related to the Y1 transposases of theIS200/IS605 family of ssDNA transposases. The IS200/IS605 family transposases are members of the HUHsuperfamily of ssDNA nucleases characterized by ahighly conserved His-hydrophobic-His (HUH) motif anda single catalytic tyrosine (29). Members of the HUHsuperfamily are involved in a wide variety of DNA trans-actions that use ssDNA substrates, such as replication ini-tiation in certain ssDNA viruses, plasmid conjugation,initiation of rolling circle replication and DNA transpos-ition (29–33). The HUH motif (34) is responsible forproviding two of the ligands coordinating a singledivalent metal ion cofactor, which binds and polarizesthe scissile phosphate group of the DNA, aligning it cor-rectly for nucleophilic attack by the catalytic tyrosine.Biochemical and structural data from a Y1 transposase

of the IS200/IS605 family, encoded by the insertionsequence (IS) IS608 from Helicobacter pylori, TnpAIS608,permitted a detailed description of the IS608 transpositioncycle (35–37). ISs are the simplest form of mobile DNAthat undergo transposition (38). Key to the mobility of theIS200/IS605 family is the ability of the transposase tobind, cleave and rejoin (or recombine) specific ssDNAsegments. Y1 transposases recognize and bind hairpinsformed by imperfect palindromes (IP) located very closeto the ends of the IS. Cleavage of the bound DNA strandat each IS end is mediated by an active-site tyrosineresidue, and results in formation of a covalent 50

phosphotyrosine linked intermediate on one side of thessDNA break and a free 30-OH group on the other. Oneof the most unusual features of mobile elements from thisfamily is that site-specific cleavages at the IS ends areachieved by a unique DNA/DNA recognition modeorchestrated by the transposase. A 4-nt sequence, theso-called ‘guide’ sequence just 50 of the foot of each IPhairpin, is bound near the active site, and through basepairing interactions, recognizes bases that are just 50 of thecleavage site at both left and right IS ends (35). This modeof site-specific DNA recognition also allows thetransposase to carry out strict site-specific cleavage andintegration, the hallmarks of the family, withoutencoding any sequence-specific DNA binding domains.The RAYT clade shares all the key amino acid motifs

exhibited by IS200/IS605 family TnpAs: the HUH motifand a catalytic tyrosine, as well as high sequence similarity(Figure 1b). Escherichia coli REPs also resemble IS200/IS605 family IP sequences in that they are approximatelythe same length and contain mismatched bases within thehairpin stems (and hence are ‘imperfect’ stem-loops).REPs carry a conserved tetranucleotide (GTAG) 50 tothe hairpin foot, similar to the guide sequences of theIS200/IS605 family ISs, and, like the guide sequence areproposed to be involved in cleavage specificity (26).In vitro assays show the conserved tetranucleotide in thepresence of REP IP sequences to be required for cleavageof ssDNA containing a CT dinucleotide site. In addition,TnpAREP is capable of catalyzing a strand transfer

reaction, one of the steps that would be required to recom-bine BIMEs (27).

Despite the prevalence of REPs in bacterial chromo-somes, the mechanism through which they are propagatedremains unclear. To better understand the functional re-lationship between REPs and their associated RAYTs, wedetermined the 3D structure of a representative fromE. coli, TnpAREP, to 2.6 A in complex with a REPsequence and its conserved 50 tetranucleotide.

MATERIALS AND METHODS

Protein expression and purification

Cloning of the tnpArep gene from E. coli MG1655 wasdescribed previously (27). For expression of TnpAREP

without a C-terminal His-tag, pBAD-tnpArep wasmodified by addition of a stop codon following serine165 to recreate wild-type sequence via site-directed muta-genesis (called pBAD-tnpArep-notag). Top10 cells(Novagen) were transformed with pBAD-tnpArep-notag.An initial overnight inoculant was grown in LB brothsupplemented with 0.5% glucose, and then added to 2 lof LB broth at a 1:20 dilution and grown to an A600 nm�0.4 at 42�C. The temperature was then dropped to 18�C,and TnpAREP expression induced at A600 nm 0.6 with0.04% arabinose. Cells were harvested by centrifugationafter 18 h, and resuspended in heparin binding buffer(20mM NaH2PO4 pH 7.0, 500mM NaCl, 1mM TCEP).All subsequent steps were performed at 4�C. Lysis was bysonication. The soluble fraction was isolated by centrifu-gation at 13 000 rpm on a Beckman Coulter Avanti J-20XP, loaded onto a HiTrap Heparin HP column (GEHealthcare) equilibrated in heparin binding buffer, andeluted using a linear gradient with elution buffer (20mMNaH2PO4 pH 7.4, 2M NaCl, 1mM TCEP). The elutedprotein was loaded onto a HiTrap Chelating column (GEHealthcare) pre-equilibrated with NiSO4, and eluted usinga linear gradient with elution buffer 2 (20mM NaH2PO4

pH 7.4, 500mM NaCl, 500mM Imidazole, 1mM TCEP).TnpAREP, with DNA substrate added at a 1:1 molar ratio,was dialyzed overnight in DNA binding buffer (50mMTris pH 7.5, 50mM NaCl, 5mMMgCl2, 1mM EDTA,0.5mM TCEP). The resulting protein–DNA complexwas loaded on a HiLoad 16/60 Superdex 200 sizingcolumn (GE Healthcare) equilibrated with DNA bindingbuffer. The eluted protein–DNA complex wasconcentrated to 5mg/ml for crystallization trials.

DNA substrate preparation

All DNA oligonucleotides were from Integrated DNATechnologies, Inc. The DNA oligonucleotides were resus-pended in 10mM Tris pH 8 and annealed by heating to95�C for 15min, then rapidly cooled on ice.

Binding assay

To test binding of TnpAREP protein and various DNAsubstrates, DNA was added to the protein followingelution from the HiTrap Chelating column at a 1:1molar ratio. This was then dialyzed overnight at 4�C in

9966 Nucleic Acids Research, 2012, Vol. 40, No. 19


DNA binding buffer, or in DNA binding buffer with add-itional NaCl (Supplementary Figure S1). These mixtureswere then loaded on a Superdex 200 3.2/30 column(GE Healthcare) and eluted using the same DNAbinding buffer. The fractionated samples were analyzedby SDS–PAGE, and visualized by silver staining,followed by coomassie staining.

Cleavage assay

For the cleavage assay TnpAREP (53 mM) and DNA sub-strate were added together at a 1:1 molar ratio anddialyzed overnight at 4�C in DNA cleavage buffer(50mM Tris pH 7.5, 50mM NaCl, 1mM EDTA,0.5mM TCEP). Samples were incubated at 37�C for45min with various divalent metal ions at 5mM concen-tration. The reactions were stopped by addition of EDTA(final concentration 5mM). The products were analyzedby SDS–PAGE.

Sedimentation velocity

TnpAREP was dialyzed against 50mM Tris pH 7.5, 50mMNaCl, 5mM MgCl2, 0.5mM EDTA and 1mM TCEP andanalyzed by sedimentation velocity at 6.7 and 13.7mM.Sedimentation velocity experiments were conducted at20.0�C on a Beckman Coulter ProteomeLab XL-I analyt-ical ultracentrifuge. A total of 400ml of each sample wereloaded in two-channel centerpiece cells and analyzed at arotor speed of 50 krpm with data collected using both theabsorbance and Rayleigh interference optical detectionsystems. For the latter, data were collected as singlescans at 280 nm using a radial spacing of 0.003 cm. Bothabsorbance and interference data were individuallyanalyzed in SEDFIT12.1 b (39) in terms of a continuousc(s) distribution of Lamm equation solutions using an un-corrected s range of 0.0–5.0 S with a resolution of 100 anda confidence level of 0.68. In all cases, excellent fits wereobtained with absorbance and interference RMSD values>0.0043 A280 and 0.0062 fringes, respectively. Solutiondensities r and viscosities h, and protein partial specificvolumes v were calculated in SEDNTERP 1.09 (40).

Crystallization

Crystals of TnpAREP bound to the first REP in the 50

BIME (Figure 1a) (GTAGGACGGATAAGGCGTTTACGCCGCATCCG) were grown in hanging drops at 20�Ccontaining a mixture of 1 ml of protein–DNA complex at5mg/ml with an equal volume of a reservoir solution of4–10% PEG 5000 MME, 0.1M MES pH 6.5 and 4–12%1-propanol. TnpAREP-REP crystals had P21212 symmetryand contained one monomer in the asymmetric unit.Crystals were derivatized by soaking in 4–10% PEG5000 MME, 0.1M MES pH 6.5, 4–12% 1-propanol and1mM ethylmercury thiosalicyclic acid for 16 h.

Data collection, structure determination and refinement

TnpAREP-REP crystals were cryoprotected using 80%mother liquor/20% glycerol mixture and flash cooled inliquid nitrogen. All diffraction data were collected at 95 Kusing Cu Ka radiation from a rotating anode source with

multilayer focusing optics and an RAXIS IV image plate.The data were integrated and scaled with XDS (41). Thestructure was determined by single-wavelength anomalousdiffraction from two Hg atoms. SHELXD was used tolocate them (42), and their parameters were refinedwith SHARP (43). The map was solvent flattened withSolomon (44), and the model was built interactively withthe program O (45,46). The structure was refined usingCNS (47) with Cartesian simulated annealing, energyminimization and individual B factor refinement. Thefinal model was refined with Refmac5, using restrainedrefinement (48). Refinement was monitored by calculatingRfree using 5% of the data set aside for crossvalidation(49). The final model was refined to an R of 22% andan Rfree of 28%. Drawings were prepared with PyMOL,ESPript and Adobe Illustrator (50,51). Additional X-raydata sets were collected on a single TnpAREP-REP crystalabove the K absorption edge of Fe2+ (7200 eV) and Mn2+

(6700 eV), and below Mn2+ (6300 eV). Using anothercrystal two more data sets were collected at 8200 eV andat 8500 eV, which are below and above the K edge of Ni2+.These data were collected at the SERCAT beamline ID22of the APS at the Argonne National Laboratory on aMAR300 CCD detector.

Particle induced X-ray emission

The identity of the metal ion co-factor present in TnpAREP

was confirmed by Particle Induced X-ray Emission (PIXE)analysis at the Hope College Ion Beam AnalysisLaboratory. Targets were prepared as described previ-ously (52), where 3ml of TnpAREP protein solution aswell as 3 ml of blank buffer solution were dropcast anddried on separate thin aluminized mylar foil targets.Each of these targets was exposed to four 10-min irradi-ations with a 3.4-MeV beam of protons for a total of0.093 nC total charge on target. The resultant X-rayswere detected at 145� to the beam in a calibrated Si(Li)detector with a thin foil filter designed to shield low-energyX-rays. The measured X-ray spectra are shown inSupplementary Figure S3. The spectra are plotted on asemi-log scale and are very similar except for twoelements: sulfur that arises from the cysteine and methio-nine groups in the protein and an unambiguous nickel Ka

peak �7.5 keV. Despite an extensive contribution of Clfrom the Tris–HCl and –NaCl buffer solutions the onlymetal visible in the protein is nickel.

Radiolabeled reactions in vitro

Cloning and expression of the tnpArep gene from E. coliMG1655 was described previously (27). TnpAREP�152was prepared by performing site-directed mutagenesis onpBAD-tnpArep by inverse PCR using primers His CATCATCATCATCATCATTAAGAAG and Trp3. This was ex-pressed and purified by growing Rosetta cells transformedwith mutant plasmid at 37�C in LB broth containing car-benicillin and 1% glucose overnight. The cells were thencentrifuged and diluted 50-fold into the 250ml preheatedLB medium at 37�C. Protein expression was induced atA600 nm �0.5–0.6 by addition of arabinose to 0.04%final. After 3 h, the bacteria were centrifuged and the



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pellet was washed in 10ml of cold TN solution (50mMTris 7.5 100mM NaCl) and stored at �20�C until use.The pellet then was resuspended in buffer A [50mMNa phosphate buffer 0.1MNa2HPO4 pH 8, 1M NaCl,10mM b-mercaptoethanol]+10mM Imidazole supple-mented with 1mg/ml lysozyme and EDTA-free proteaseinhibitor cocktail (Roche). After 30-min incubation on ice,the bacteria were sonicated, the lysate was cleared by cen-trifugation and the supernatant was then mixed withNi-agarose resin (Qiagen). After washes in bufferA+50mM Imidazole, TnpAREP�152 was eluted withbuffer A+200mM Imidazole. The protein was thendialyzed in 25mM HEPES pH 7.5, 400mM NaCl, 1mMEDTA, 5mM DTT and 20% glycerol and stored at�80�C.Oligonucleotides, purchased from Sigma and Euro-

gentec, were 50-end-labeled with [g-32P] ATP (PerkinElmer) using T4 polynucleotide kinase (NEB) or30-end-labeled with [a-32P] dATP Cordycepin (PerkinElmer) using Terminal Transferase (NEB), and subse-quently were purified on a G25 column (GE Healthcare).Double stranded DNA was prepared by hybridization ofcomplementary oligonucleotides. After 10min denatur-ation at 98�C, the mixture was left to slowly cool to25�C. For cleavage, 0.02mM 50-labeled oligonucleotideand 0.5 mM unlabeled oligonucleotide were incubatedwith 4 mM TnpAREP (45min, 37�C, final volume 10 ml) in12.5mM Tris pH 7.5, 120mM NaCl, 5mM MnCl2, 1mMDTT, 20 mg/ml BSA, 0.5 mg of poly-dIdC and 7% glycerol.Reactions were separated on a 9% denaturing gel andanalyzed by phosphor imaging.In reactions to detect covalent complex formation, 30

labeled substrates were incubated with TnpAREP in thereaction mixture as described above and reactionproducts were separated on a 16% SDS–PAGE gel andanalyzed by phosphor imaging.

RESULTS

Overall structure

The structure of TnpAREP fromE. coliMG1655 in complexwith a 32-mer oligonucleotide (Figure 2a) representing aREP and its associated 50 GTAG tetranucleotide (fromthe left-most BIME in Figure 1a; shown schematically inFigure 2b), was solved to 2.6 A using single-wavelengthanomalous diffraction from a mercury derivatized crystal,and refined to an R/Rfree of 22%/28% (Table 1). TheTnpAREP structure shows a monomer bound to one DNAmolecule, consistent with the behavior of the protein inanalytical ultracentrifugation analysis (Figure 2c). TheDNA is bound to the protein through two extensive setsof interactions, one involving the bases at the bottom 30

region of the IP stem, and the other the 50 GTAGsequence. We also observe a divalent metal ion (mostlikely Ni2+; see below) bound at the active site.The overall fold of TnpAREP conforms to that of the

RNA recognition motif of other HUH endonucleases(31,53,54), where a four-stranded antiparallel b-sheet issandwiched by two helices on one side (aB and aC), anda singular helix (aD) on the other side (Figure 2a). Its

topology is also very similar to those of the IS200/IS605family transposases (29,55) (Figure 1b), which is in agree-ment with their phylogenetic relationship (26,27).

TnpAREP is a monomer

Despite the clear structural and sequence similaritiesbetween TnpAREP and IS200/IS605 family transposasesat the monomer level, IS200/IS605 transposases arealways observed as interwoven obligatory dimers, bothin the unbound form and when complexed with avariety of DNA molecules (Figure 3a and b). Thecomplexes also always contained two IPs, binding oneIP per monomer (29,55).

The comparison between TnpAREP and the IS200/IS605transposases reveals that two key structural determinantsresponsible for the dimerization interface of �2500 A2 inIS200/IS605 family members are missing. As shown inFigure 3a the two b-sheets from individual IS200/IS605monomers are linked by backbone hydrogen bondingbetween each b5 (residues 111–115) and b2 (residues12–18). This interaction begins with the side chain of thehighly conserved T115 in b5 bonded to K18 in b2.TnpAREP b1, equivalent to b2 in TnpAIS608 (Figure 1b),is markedly shorter and hence unable to be sharedbetween monomers (Figure 3a). Furthermore, the highlyconserved T117 at the start of this linkage does not exist inTnpAREP, but is replaced by A102.

The second important structural element contributingto dimerization of IS200/IS605 family transposases is acomplementary hydrophobic surface at the dimer interfaceformed by residues contributed by both monomers. A keyresidue in this pocket is P73 located just after b4 inTnpAIS608 (Figure 3b) (PDB: 2A6M, 2VHG, 2VJU)(29,35). This residue is conserved among IS200/IS605family transposases, but replaced with the polar residueE68 in TnpAREP. Other residues that form the complemen-tary hydrophobic surface in TnpAIS608, such as V77 andI113, are replaced with polar residues in TnpAREP (S74and E100) (Figure 1b). Although in TnpAIS608 there aresome hydrophobic interactions between the domain-swapped aD and the protein (Figure 3a), the position ofthis helix varies in the different structures suggesting thatthese interactions are not critical for dimerization.

DNA binding interactions

REP DNA is bound to TnpAREP through a substantialinterface, which involves both the DNA hairpin and the 50

GATG tetranucleotide; the total binding interactionbetween the protein and DNA buries a surface area of�1300 A2. The hairpin consists of 10 Watson–Crick basepairs, interrupted in the middle by four bases, which forma mismatched bulge (C27:A12, A13:G26), and two unpairedTs (T19 and T20) in the hairpin loop (bases are numberedas shown in Figure 2b). Most of the interaction involves anetwork of hydrogen bonds between residues comprising aregion of strong positive electrostatic potential on theprotein surface and the negatively charged phosphateoxygens that form the backbone of the hairpin(Figure 4a). The 5 bp closest to the hairpin tip do notcontact TnpAREP nor do the two T bases of the tip itself.



The observed interactions suggest that binding of thehairpin is mediated mostly, but not exclusively, throughthe secondary structure of the DNA hairpin. This obser-vation is reinforced by hairpin binding experiments, whichshow a salt concentration dependency of DNA binding(Supplementary Figure S1). The positive surface chargeof the protein is comprised primarily of residues fromaB and aC (R46, R78, K81, K82, Q83, T85, H86, K91and R97) with aC inserted into the minor groove of thehairpin. Interestingly, there are base specific interactionsbetween K82 and the mismatched C27, R78 and T11, andR97 and G32 (Figure 2b). In TnpAREP, the K82 inter-action is critical for overall binding, since mutation ofA12 to G and A13 to C to correct the mismatch abolisheshairpin binding as judged by a loss of co-migration of

protein and DNA by size exclusion chromatography(Figure 4b, substrate designated ‘REP plus GTAG nobulge’) (27). In addition, randomizing the nucleotides inthe hairpin, while maintaining the hairpin structure andsequence of the bulge, also abrogates binding (Figure 4b:‘REP plus GTAG random’). These results confirmthe importance of these three protein–base specificinteractions.In sharp contrast to the interactions formed with the

hairpin, binding of the conserved 50 GTAG sequence ishighly base-specific and buries �590 A2 (Figure 5a). Thefour bases are splayed across the surface of the protein,pointing inward such that only the phosphate backbone isaccessible to solvent. This differs from the structure ofTnpAIS608 bound to its IP hairpin, which was extended

90°

(a) (b)

(c)

5’3’32

1

Gui

de s

eque

nce

(GTA

G)

Mismatch

Hairpin loop

C

3’

5’

αB

αC

αD 5’

1

Figure 2. TnpAREP structure. (a) Ribbon diagrams of TnpAREP bound to hairpin, with one orientation rotated 90� around the y-axis relative to theother. DNA is colored green, with the bases of the mismatched bulge in magenta. The C-terminus is highlighted in purple, the catalytic tyrosine as astick model in magenta, resides involved in the divalent metal coordination as stick models in orange and the divalent metal as a black sphere.(b) Schematic of the binding hairpin with important binding residues, and interactions between the DNA and protein by red arrows. (c) Graphicalrepresentation of the analytical ultracentrifugation by sedimentation velocity results. Sedimentation velocity experiments carried out at two loadingconcentrations indicated the presence of a major species at 2.35±0.01 S, representing >98% of the loading signal. The best-fit molar mass for thisspecies is 20.3±0.8 kDa demonstrating that TnpAREP is a monomer at these concentrations.





to include its 50 tetranucleotide guide sequence. In thatstructure, the four 50 nucleotides at the base of the hairpincurl away from the surface of the protein (Figures 5b and6b), apparently poised to recognize and bind the cleavagesite. In particular, two bases of the GTAG sequence inTnpAREP, G4 and A3, point toward the active site andform a number of H-bonds with residues of theC-terminus around E161 (Figure 5a).In vitro binding experiments showed that the four 50

tetranucleotide bases are crucial for REP binding. Intheir absence, binding to the isolated hairpin was notdetected (Figure 4b: ‘REP’) (27). On the other hand,while this sequence is necessary for hairpin binding, it isnot sufficient, as addition of the conserved sequence to aniREP does not confer detectable binding (Figure 4b:‘iREP plus GTAG’).It was also shown that the four bases comprising the 50

GTAG sequence are crucial for cleavage activity, aschanging the sequence from GTAG to ACGA results inthe loss of cleavage (27). To further understand the role ofthe 50 GTAG, we individually changed each of the fourbases and then tested for cleavage and binding of thesemodified substrates. Each base change was chosen topreserve as much as possible the observed DNA proteininteractions while disrupting optimal base pairing. Asshown in Figure 5c, when cleavage was monitored by fol-lowing the formation of a covalent intermediate, mutationof any of the four bases resulted in the loss of cleavageactivity (compare lane 4 with lanes 5–8). Also, DNAbinding could no longer be detected (SupplementaryFigure S2). The reciprocal substitution, changing G4 toC and the C of the cleavage site to G, did not rescuecleavage (Figure 5c; lane 13).

Active site

As shown in Figure 6a, the active site of TnpAREP bringstogether the catalytic tyrosine Y115 and the HUH motif(H59, M60, H61), which together with two residues fromaD (N119, H123) and E161 of the C-terminus,octahedrally coordinate a co-purifying metal ion. Whencompared with the active sites of Y1 transposases, theactive site of TnpAREP most closely resembles that ofTnpAISDra2(Y132F) bound to the right end of ISDra2from Deinococcus radiodurans (PDB: 2XO6) (55). Theinactivating Y132F mutation of the active site tyrosineallowed the crystallographic capture of the fully assembledactive site, including the scissile phosphate (55).Superposition of 2XO6 with TnpAREP shows that theaD helices align, and are positioned directly over b2 andb3, and in both cases the catalytic tyrosines face into theactive site (Figure 6b).

The position of the co-purifying metal ion at theTnpAREP active site is the same as the catalytically essentialdivalent cations seen in otherHUHnucleases (29,35,55,56).To determine the metal in the active site, we collected X-raydiffraction data at several energies near the K absorptionedges of several metal ions, including 8200 eV (below theNi2+ K edge), and at 8500 eV (above the Ni2+ K edge).Comparisons of the peak heights in the anomalous differ-ence Fourier maps using these data suggest the metal ismost likely Ni2+. The identity of the metal ion wasfurther confirmed via Particle-induced X-ray emission(PIXE) that again indicated the presence of Ni2+ in asample that was prepared identically to the one used forcrystallization (Supplementary Figure S3). Ni2+is bound tothe HiTrap Chelating affinity column (GE Healthcare)

Table 1. Data collection and refinement statistics

TnpAREP at 8027 eV TnpAREP at 8200 eV TnpAREP at 8500 eV

Data collectionSpace group P21212 P21 P21

Cell dimensionsa, b, c (A) 100.83, 60.30, 70.78 39.44, 61.50, 70.20 39.52, 61.66, 70.34a, b, g (�) 90.00, 90.00, 90.00 90.00, 95.12, 90.00 90.00, 95.09, 90.00Resolution (A) 30–2.6 (2.67–2.6)* 60–2.5 (2.57–2.6)* 60–2.5 (2.57–2.5)*Rsym or Rmerge 4.0 % (56.5%) 3.1 % (51.5%) 3.7 % (57.2%)I/sI 23.46 (2.23) 21.28 (2.23) 18.66 (2.06)Completeness (%) 99.7% (100.0%) 99.6 (99.9%) 99.6% (99.8%)Redundancy 3.5 (3.5) 3.7 (3.7) 3.7 (3.7)

RefinementResolution (A) 2.6No. reflections 12 056Rwork/Rfree 22%/28%No. atoms

Protein 2080Ligand/ion 1Water 22

B-factorsProtein/DNA 54.11Ligand/ion 38.93Water 43.71

RMS deviationsBond lengths (A) 0.016Bond angles (�) 1.930

*Values in parentheses are for highest-resolution shell.








used during purification, and thus is the likely source of themetal observed in the structure. Significantly, we foundthat only Mn2+ supports cleavage and the formation of acovalent intermediate, while no activity was observed in thepresence of other divalent cations (Figure 6c).

The most surprising aspect of the active site organizationis that the carboxylate group of the metal-coordinatingresidue E161 occupies the same position as the scissile phos-phate in the TnpADra2(Y132F) transposase crystal structure(55,57). E161 is conserved among many of the RAYTproteins, and is part of an �25 amino acid C-terminalextension not found in the IS200/IS605 transposases

(Figure 1b and Figure 7b). As shown in Figure 7a, thisC-terminal extension (shown in purple) packs intimatelyinto a long surface groove that wends its way in a semicir-cular path across the surface of TnpAREP.The location of the C-terminal extension suggests that

TnpAREP is in an auto-inhibited state in the crystal, as itappears to be physically blocking access of a ssDNAcleavage substrate to the active site. We reasoned if theC-terminus of TnpAREP were indeed acting as an inhibitor,one way to relieve this inhibition would be to delete it.We therefore expressed and purified a deletion mutant ofTnpAREP where all residues after G152 had been deleted

(a)

(b)

β sheet dimerization

Complementary hydrophobic surface

TnpAIS608

TnpAREP

β5β2

β3β4

β3 β2

β1

β4

β5

β2

β3β4

T117

αD

αD

αD

Figure 3. TnpAREP versus TnpAIS608 and dimerization. (a) (Left) TnpAIS608 as a ribbon diagram with one molecule colored cyan and the othermolecule in magenta, with the DNA in gray. Note that aD containing the catalytic tyrosine is domain swapped. (Right) TnpAREP is aligned to thecyan monomer of TnpAIS608, and is also colored in cyan, with DNA in gray. Note the differences in the positioning of aD. In each molecule theb-strands are labeled (b2, b3, b4, b5 of TnpAIS608 are equivalent to b1, b2, b3, b4 of TnpAREP). (b) One monomer of the TnpAIS608 dimer isrepresented as a surface diagram in cyan and the second molecule as a ribbon diagram in magenta, with all DNA in gray. The two majorcomponents of dimerization are highlighted, with the key surface area involved in the b sheet interaction shown in red and the surface areainvolved in hydrophobic interaction in gray.



(designated TnpAREP�152), and tested it for cleavageactivity. As shown in Figure 7c, C-terminal truncation ofTnpAREP indeed resulted in a increase of cleavage activityrelative to the full-length protein. TnpAREP�152 was alsocompetent for strand recombination (SupplementaryFigure S4), indicating that the C-terminus is not neededfor strand transfer.

Cleavage site recognition

It was previously reported that TnpAREP is specificfor cleavage at CT dinucleotide sequences, and thatthe CT can be located on either side of the REP hairpinand at a considerable distance from the REP hairpin (27).In addition, the inhibitory C-terminal extension of

(a)

REP plus GTAGGTAGGACGGATAAGGCGTTTACGCCGCATCCG

REPCGGATAAGGCGTTTACGCCGCATCCG

REP plus GTAG no bulgeGTAGGACGGATGCGGCGTTTACGCCGCATCCG

REP plus GTAG randomGTAGGACCAAGAACAGATTTATCTGGCCTTGG

iREP plus GTAGTGATGCGACGCTGGCGCGTCTTATCATGGATG

(b)

G1

T2

A3

G4

36 kDa31

2114

63.5

Protein

DNA

Figure 4. TnpAREP DNA hairpin binding. (a) View of the TnpAREP molecule showing electrostatic charge surface in a vacuum bound to its hairpin.Shades of blue represent positive charge, and red negative charge. The DNA is colored green with the individual bases of the 50 GTAG labeled.(b) SDS–PAGE analysis of TnpAREP binding by size exclusion chromatography of modified REP and iREP substrates. The lanes in each gelrepresent the same elution volume off of a Superdex 200 3.2/30 column (GE Healthcare). The protein and DNA were visualized by successivestaining with silver and coomassie blue. The top gel represents the wild-type binding between protein and DNA substrate (i.e. co-migration of proteinand DNA), while the four subsequent gels show a lack of DNA binding as evidenced by lack of co-migration.






TnpAREP makes a number of hydrogen bonds with twobases of the 50 GTAG sequence, G4 and A3, which pointtoward the active site (Figure 5a). In the IS200/IS605transposase family, the guide sequence bases recognizenucleotides just 50 of the cleavage site and therefore

determine cleavage specificity. Assuming a similar rolefor G4 and A3 in TnpAREP would also imply cleavagesite specificity at a CT.The observed conformation of the 50 GTAG sequence is

remarkable. A3 and G4 are stacked on each other and in

(a)

(c)

(b)

Figure 5. Analysis of the 50 GTAG sequence. (a) Ribbon diagram of TnpAREP in cyan with key residues and bases involved in guide sequencebinding shown as stick models. Hydrogen bonds are shown as dashed lines. (b) Schematic display of the binding hairpin from TnpAREP withimportant binding residues as in Figure 2b, with the corresponding schematic display of the binding hairpin with binding residues from TnpAIS608.(c) In the top part is displayed a typical SDS–PAGE analysis of TnpAREP cleavage in which the DNA and protein are denatured. The DNA isvisualized by silver stain, and the protein is visualized by coomassie stain. To the right of the gel are labels defining the bands, with Protein plusDNA label emphasizing protein–DNA covalent complex in the higher bands of lanes 4 and 11. A schematic of substrate used is to the right. Below isa table of substrates tested for cleavage by TnpAREP, where the red arrow/asterisk indicates the cleavage site, the purple lettering the guide sequence,the red lettering the hairpin, the blue lettering iREP sequence and magenta lettering nucleotides that have been mutated.



principle can base pair with the CT of a cleavage sub-strate. However, in the crystal structure, the Watson–Crick face of A3 is pointing away from the active siteand the nucleotide is in the anti conformation. G4 is alsoin the anti conformation but its Watson–Crick face pointstoward the active site and therefore is available forbase-pairing. For A3 to become available for base-pairing,it would either have to flip into the syn conformation, or,more likely, some DNA backbone movement would benecessary to turn the base so it can recognize the T ofthe cleavage site sequence. Notably, none of the nucleo-tides flanking A3 and G4 are available for base-pairingneither with cleavage site bases nor with any of theadjacent bases. W99 is positioned between A3 and T2,moving T2� 9 A away from the active site. Neither T2

nor G1 is available for base-pairing, because of extensivehydrogen bonding with the protein. Similarly, due to theinsertion of R21 and Q95, G5 is �12 A away, and while itsWatson–Crick face is partially accessible, it is also held in

syn due to a hydrogen bond to a non-bridging oxygen ofits own phosphate. Both its distance from G4 and thegeometry of the backbone makes it very unlikely that G5

could participate in cleavage site recognition. Takentogether, it appears that the observed mode of 50 GTAGbinding assures that only A3 and G4 are available for sub-strate recognition, consistent with the CT dinucleotidesequence requirement for cleavage.

As shown in Figure 5c (lanes 9, 10), mutation of the Cor T at the cleavage site to G and A, respectively,abolishes activity. In an artificial substrate in which wemoved the CT dinucleotide upstream, closer to thehairpin by eight bases, cleavage was moved to the newlocation of the CT (Figure 5c; lane 11). This result con-firmed that the CT is necessary and sufficient to determinethe cleavage site. Interestingly, if the enzyme and substratewere working in trans, as was the case for both TnpAIS608

and TnpADra2, then the combination of substrates 5 and 9would provide one correct cleavage site in substrate 5 and

H59H61

E161

N119

H123

G4

A3

Y115

(b)(a)

(c)Metal: -- -- Mn2+ Mg2+ Cd2+ Ca2+ Zn2+ Ni2+

B268i: -- + + + + + + +

Protein

Protein + DNA

B268i:

REP iREP

5’5’ 5’

CT5T5

TnpATnpADra2Dra2

TnpAREP

31 kDa

21

14

6

3.5

Figure 6. TnpAREP active site. (a) The active site of TnpAREP with critical residues highlighted. Protein/DNA is displayed as a ribbon diagram, withthe protein in cyan and DNA in green. Key ligands to the cation, colored dark gray, are displayed in orange (H59, H61, N119, H123) and purple(E161). The C-terminus is highlighted in purple. (b) TnpAREP and DNA in blue and TnpADra2 and DNA in yellow shown as ribbon diagrams tohighlight the T5 substrate of TnpADra2 and the C-terminus of TnpAREP, and differences in 50 GTAG conformation. Key residues in the active site ofTnpAREP and TnpADra2 are colored blue and yellow, respectively, with their oxygens colored red and nitrogens blue. (c) A typical SDS-PAGEanalysis of TnpAREP cleavage in which the DNA and protein are denatured. The DNA is visualized by silver stain, and the protein is visualized bycoomassie stain. To the right of the gel are labels defining the bands, with Protein plus DNA label emphasizing protein-DNA covalent complex in thehigher bands of lanes 4. A schematic of substrate used is to the right.



one correct 50 GTAG in substrate 9 from each protein/DNA complex (Figure 5c; lane 14). However, this did notrescue activity, suggesting no in trans cooperation betweencomplexes.

DISCUSSION

Repetitive extragenic repeat sequences (REPs) are foundin many bacterial species in high copy numbers and theycan have a variety of functions (21–24). These widespreadREPs are associated with TnpAREP, closely related to Y1transposases of the IS200/IS605 family (26,27), and thereis strong evidence suggesting that they may be capable ofacting upon or perhaps even mobilizing REPs. Excision ofREPs was reported in P. fluorescens (28), and it wasrecently demonstrated that the TnpAREP from E. colistrain MG1655 is capable of ssDNA cleavage and recom-bination (27). The structure of TnpAREP suggests thatalthough many of the hallmarks of Y1 transposases areretained, the enzyme is highly regulated and exhibits struc-tural features important for keeping its activity in check.

The structure of TnpAREP bound to a REP hairpinincluding its conserved 50 tetranucleotide extension weobserved is evocative of the structures of the IS200/IS605 transposases bound to their DNA intermediates.Together with the previously determined structures ofIS608 and ISDra2 transposases it provides an elegant ex-planation for the CT specificity of cleavage by TnpAREP.In particular, the pattern of base recognition first observedfor IS608 (Figure 8a) (35) and later echoed in ISDra2

(Figure 8b) (55) follows the same rules of base–baseinteraction. Thus G4 of TnpAREP dictates C on the 50

side of the cleavage site, while A3 dictates T on the 30

side (Figure 8c). While this explains the role of A3 andG4 within the proposed guide sequence, the role ofother two conserved nucleotides, G1 and T2, is less clear.This raises the question of whether the conserved 50

GTAG guide sequence is dictating cleavage specificity inthe same fashion as the IS200/IS605 transposases. Asshown in the schematic in Figure 8, in IS200/IS605transposases, three of the four guide sequence nucleotidesinteract with nucleotides 50 of the cleavage sequence todictate cleavage after specific tetra- or pentanucleotide se-quences (35). In contrast, only two bases constitute thecleavage site of TnpAREP with a C on the 50 side and Ton the 30 site of the strand break.Our results are consistent with this different mode of

base recognition, and the crystal structure indicates thatthe 50 GTAG sequence is bound in such a way that only2 nt, A3 and G4, are available for base-pairing with thecleavage substrate sequence. Specific protein residues(e.g. W99, R21, Q95) assure that flanking nucleotidesare sequestered and unavailable, thereby preventingthem from contributing as specificity determinants. Thisis consistent with analysis of cleavage in E. coli MG1655and P. fluorescens (27,28) (Figure 5c) where cleavageoccurs at CT sites irrespective of the flanking nucleotides.It is intriguing that there is considerable flexibility in the

location of the CT dinucleotide sequence relative to theREP hairpin (27). The reliance only on 2 nt for specificity

(a) (b)

(c)

1 5 15 60 1 5 15 60 1 5 15 60 1 5 15 60 0 min

- metal + Mn2+

WT Δ152 Δ152WT

S165

D153

5’

T5

5’

3’

B268i:

REP iREP

TnpADra2TnpAREP

Figure 7. TnpAREP�152 and the role of the C-terminus. (a) TnpAREP protein shown with molecule surface modeled in cyan, DNA in light gray as aribbon diagram and the C-terminus as a stick model colored in purple. (b) TnpADra2 also as a surface model in cyan, with DNA in light gray inthe same orientation as TnpAREP in panel a to highlight the similar role and position of the TnpAREP C-terminus to the T5 DNA substrate ofTnpADra2. (c) SDS–PAGE analysis of DNA cleavage by TnpAREP and TnpAREP�152.



and binding may imply that the exact geometry of scissilephosphate presentation for cleavage is less precise thanseen in the IS200/IS605 transposases. Correspondingly,we observe an active site in TnpAREP that prefers Mn2+

rather than Mg2+, most likely due to three coordinatingimidazoles (H59, H61, H123). A search of the MIPSdatabase (58) reveals that there are only eight structurescurrently in the PDB where Mg2+ is coordinated by threeimidazoles (and, in some cases, close inspection ofthese coordinate sets raises suspicions about the assignedmetal’s identity) but 109 structures where Mn2+ is coor-dinated by three histidine ligands. As the geometry of theoctahedral coordination around a Mn2+ ion can be lessstrict than that of Mg2+(59), Mn2+might be more suitablefor TnpAREP as it has to deal with a range of cleavagesubstrates which may not be precisely positioned withinthe active site. Interestingly, conjugative relaxases, whichalso bind their ssDNA cleavage substrates withoutstabilizing base pairing interactions, similarly use threehistidines to coordinate a metal ion cofactor, although a

preference for Mn2+ has not been demonstrated for thisclass of enzyme (32,60–62).

Molecular machines carrying out transposition reac-tions are often found in forms that have suboptimalactivity. This is understandable as highly active run-away transposition may cause genomic damage andcould result in the loss of viability. In turn, this givesrise to the possibility of engineering hyperactive versionsof both prokaryotic and eukaryotic transposases for use ina variety of genomic applications (63–65). It is now alsoclear that stress conditions such ionizing radiation cansometimes adventitiously relieve transposon inhibition(66–68), perhaps as a last resort in an attempt togenerate genetic diversity and speed up adaptation. Inthe structure of TnpAREP bound to REP DNA, theC-terminal tail is bound within the active site suggestingthat this conserved sequence feature of TnpAREP proteinsis a mechanism to inhibit or down-regulate the potentialactivity of the protein. Consistent with this notion,C-terminal truncation results in dramatically increasedcleavage activity, while retaining competence for strandrecombination (Figure 7c). Thus, this C-terminal regionis not important for catalysis but instead appears toserve a regulatory role.

The TnpAREP structure shows a novel way in whichdown-regulation of a DNA rearranging system can beachieved by using a part of the transposase to act asan inhibitor of its own active site. While autoinhibitionis a regulatory feature of many enzyme systems, andautoregulation has been observed through in vitro studyof the DNA transposases Tn5, Tn10 and IS911 (69–72), toour knowledge this is the first time that it has been seen ina structure of a DNA transposase working as competitiveautoinhibitor. It would be interesting to establish if thereis a signal or particular growth condition in E. coli thatactivates TnpAREP either by interaction with other cellularproteins, by proteolytic removal of the inhibitingC-terminal tail or perhaps by the production of atruncated and hence hyperactive form of TnpAREP suchas the deletion version we characterized.

Another consequence of removal of the C-terminal tailfrom the active site is that this would necessarily destroythe interactions observed between G4 and A3 of theconserved 50 tetranucleotide and the residues aroundE161 (Figure 5a). This could provide yet another level ofregulation as, at least prior to binding a cleavage sub-strate, the REP 50 GTAG sequence is bound toTnpAREP through a dense network of interactions thatrenders it apparently unavailable to recognize a suitablecleavage site. The C-terminal tail binding appears to bevery tight as, despite extensive efforts, we have not beenable to detect binding of DNA containing a CT cleavagesite (added in the form of an oligonucleotide) to apre-formed TnpAREP–REP complex. This has, to date,prevented us from structurally characterizing a cleavagesite complex using the type of experimental approachesthat proved successful for the IS608 and ISDra2transposases.

One of the most intriguing aspects of the known bio-chemical activities of TnpAREP is its apparent ability tocarry out strand transfer in vitro. Transposases of the

T T A C

TnpAIS608

LE Cleavage site

A A A G

T T G A T G

TnpADra2

LE Cleavage site

C A C A

T T C A A

TnpADra2

RE Cleavage site

G A A T

T C A A

TnpAIS608

RE Cleavage site

G A A T

? ? ? ? C T

TnpAREP

CT Cleavage site

G T A G

(a)

(b)

(c)

Figure 8. Models of guide sequence cleavage site recognition.(a) Scheme of TnpAIS608 guide sequence cleavage site recognition.(b) Scheme of TnpADra2 guide sequence cleavage site recognition.(c) Scheme of TnpAREP guide sequence cleavage site recognition.



IS200/IS605 family can do this straightforwardly as theyare obligatory dimers with two active sites and can bindsimultaneously both left and right ends of the mobileelement (73). Following end cleavages, the 30 end of thecleaved strands stays in the active site, bound there bybase-pairing interactions with the protein-bound guidesequences. The other end becomes covalently attached tothe transposase through a 50-phosphotyrosine linkage tothe nucleophilic tyrosine located on a mobile helix. A con-formational change, in which the two helices carrying thenucleophilic tyrosine swap places within the dimer,delivers the attached 50 ends to the other active site ofthe dimer where the parked 30-OH of the cleaved strandcan then attack the 50-phosphotyrosine. This reestablishesthe phosphodiester backbone with a different connectivityand hence strands are transferred. It is not obvious howthis could happen in the context of the monomericTnpAREP.

The organization of the TnpAREP active site when boundto the GTAG sequence suggests one possible mechanismfor strand transfer. It seems likely that after cleavage at aCT dinucleotide, the ssDNA 50 of the cleavage site wouldreadily dissociate. As the covalent 50-phosphotyrosinelinkage remains, in principle any other ssDNA possessinga 30-OH end could enter the active site and chemistry couldproceed, thereby resolving the 50-phosphotyrosine linkageand accomplishing strand transfer. The notion of a mono-meric HUH nuclease catalyzing a strand transfer reaction,while not common, is not novel. The conjugative relaxaseTrwC of plasmid R388 (but interestingly not the relatedTraI of the F plasmid) can catalyze strand transfer (74) ascan a deletion mutant of TrwC that is monomeric.Furthermore, strand transfer occurs even with the Y26Fmutant of TrwC, in which only one of the two catalytictyrosines (Y18) is functional. Interestingly, similar toTnpAREP the 30 end nick site is not bound by base pairinginteractions giving rise to the possibility that it can bereplaced by an 30-OH end resulting in strand transfer (32).

The key observation that TnpAREP is a monomer—incontrast to the characterized IS200/IS605 transposases,which are obligate dimers—imposes several constraintson any model of its activities. For example, there is noready mechanism to explain the possibility of coordinatedcleavage events on the two ends of a single BIME unlessthese are mediated through the DNA rather than by theprotein. Furthermore, it is not clear how a circular BIMEintermediate could be excised from a DNA strand in theabsence of the type of reciprocal strand exchange stepsbetween two active sites that have been invoked for theIS200/IS605 family transposases (35). Any proposedmechanism for propagation must also take into accountthat, whereas TnpAREP binds REPs, binding to iREPs hasnot been detected (27) (Figure 4b). Although we cannotrule out that TnpAREP undergoes a major conformationalrearrangement during the reaction that could result in di-merization, we have no evidence that it does so. Indeed,our structure demonstrates that even upon REP binding,TnpAREP remains monomeric.

While sequence similarities and many of our structuralobservations point to the close relationship betweenTnpAREP and transposases from the IS200/IS605 family,

the overall evidence here puts into question whether or notTnpAREP uses an IS200/IS605 transposition mechanism.In particular, the loss of the ability to dimerize, as well asthe differing role for the conserved 50 tetranucleotidesequence in the REPs, point to crucial differences inprotein activities. Furthermore, the enzyme appearshighly regulated through auto-inhibition by theC-terminus and use of manganese, suggesting an evolvedmechanism to limit REP populations in the cell. In allprobability, TnpAREP started as an IS200/IS605 trans-posase, but subsequently evolved into a REP propagationenzyme developing its own distinct ‘transposition’ mech-anism that is kept under tight check.

ACCESSION NUMBERS

Coordinates have been deposited in the Protein DataBank with accession code 4ER8.

SUPPLEMENTARY DATA

Supplementary Data are available at NAR Online:Supplementary Figures 1–4.

ACKNOWLEDGEMENTS

We would especially like to thank John Chrzas at theSERCAT 22-ID beamline for collecting the data at8200 eV and 8500 eV, and Megan M. Sibley, and Paul A.DeYoung, for help with the PIXE measurements, thatidentified the bound metal ion species. We thank B.Marty for expert technical assistance. Data were also col-lected at the SER-CAT 22-ID beamline at the AdvancePhoton Source, Argonne National Laboratory.

FUNDING

Intramural Program of the National Institute of Diabetesand Digestive and Kidney Diseases of the NationalInstitutes of Health (F.D., in part); Nancy NossalFellowship award from NIDDK (to S.M.); PostdoctoralIntramural Research Training Award from NIDDK(to S.M.); CNRS (France, the work in Toulouse was sup-ported by intramural funding); ANR [285051 to M.C.]and US Department of Energy, Basic Energy Sciences,Office of Science [Contract No. W-31-109-Eng-38, use ofAPS]. Funding for open access charge: IntramuralProgram of the National Institute of Diabetes andDigestive and Kidney Diseases of the National Institutesof Health.

Conflict of interest statement. None declared.

REFERENCES

1. Lee,R., Feinbaum,R. and Ambros,V. (2004) A short history of ashort RNA. Cell, 116, S89–92.

2. Lee,R.C., Feinbaum,R.L. and Ambros,V. (1993) The C. elegansheterochronic gene lin-4 encodes small RNAs with antisensecomplementarity to lin-14. Cell, 75, 843–854.




3. Garneau,J.E., Dupuis,M.E., Villion,M., Romero,D.A.,Barrangou,R., Boyaval,P., Fremaux,C., Horvath,P.,Magadan,A.H. and Moineau,S. (2010) The CRISPR/Cas bacterialimmune system cleaves bacteriophage and plasmid DNA. Nature,468, 67–71.

4. Horvath,P. and Barrangou,R. (2010) CRISPR/Cas, the immunesystem of bacteria and archaea. Science, 327, 167–170.

5. Barrangou,R., Fremaux,C., Deveau,H., Richards,M., Boyaval,P.,Moineau,S., Romero,D.A. and Horvath,P. (2007) CRISPRprovides acquired resistance against viruses in prokaryotes.Science, 315, 1709–1712.

6. Boccard,F. and Prentki,P. (1993) Specific interaction of IHF withRIBs, a class of bacterial repetitive DNA elements located at the30 end of transcription units. EMBO J., 12, 5019–5027.

7. Espeli,O. and Boccard,F. (1997) In vivo cleavage of Escherichiacoli BIME-2 repeats by DNA gyrase: genetic characterization ofthe target and identification of the cut site. Mol. Microbiol., 26,767–777.

8. Gilson,E., Perrin,D. and Hofnung,M. (1990) DNA polymerase Iand a protein complex bind specifically to E. coli palindromicunit highly repetitive DNA: implications for bacterialchromosome organization. Nucleic Acids Res., 18, 3941–3952.

9. Gilson,E., Rousset,J.P., Clement,J.M. and Hofnung,M. (1986) Asubfamily of E coli palindromic units implicated in transcriptiontermination? Ann. Inst. Pasteur Microbiol., 137B, 259–270.

10. Higgins,C.F., McLaren,R.S. and Newbury,S.F. (1988) Repetitiveextragenic palindromic sequences, mRNA stability and geneexpression: evolution by gene conversion? A review. Gene, 72,3–14.

11. Oggioni,M.R. and Claverys,J.P. (1999) Repeated extragenicsequences in prokaryotic genomes: a proposal for the origin anddynamics of the RUP element in Streptococcus pneumoniae.Microbiology, 145, 2647–2653.

12. Black,C.G., Fyfe,J.A. and Davies,J.K. (1995) A promoterassociated with the neisserial repeat can be used to transcribe theuvrB gene from Neisseria gonorrhoeae. J. Bacteriol., 177,1952–1958.

13. De Gregorio,E., Abrescia,C., Carlomagno,M.S. and DiNocera,P.P. (2003) Ribonuclease III-mediated processing ofspecific Neisseria meningitidis mRNAs. Biochem. J., 374, 799–805.

14. Rouquette-Loughlin,C.E., Balthazar,J.T., Hill,S.A. andShafer,W.M. (2004) Modulation of the mtrCDE-encoded effluxpump gene complex of Neisseria meningitidis due to a Correiaelement insertion sequence. Mol. Microbiol., 54, 731–741.

15. Gilson,E., Clement,J.M., Brutlag,D. and Hofnung,M. (1984) Afamily of dispersed repetitive extragenic palindromic DNAsequences in E. coli. EMBO J., 3, 1417–1421.

16. Higgins,C.F., Ames,G.F., Barnes,W.M., Clement,J.M. andHofnung,M. (1982) A novel intercistronic regulatory element ofprokaryotic operons. Nature, 298, 760–762.

17. Gilson,E., Saurin,W., Perrin,D., Bachellier,S. and Hofnung,M.(1991) Palindromic units are part of a new bacterial interspersedmosaic element (BIME). Nucleic Acids Res., 19, 1375–1383.

18. Gilson,E., Saurin,W., Perrin,D., Bachellier,S. and Hofnung,M.(1991) The BIME family of bacterial highly repetitive sequences.Res. Microbiol., 142, 217–222.

19. Bachellier,S., Saurin,W., Perrin,D., Hofnung,M. and Gilson,E.(1994) Structural and functional diversity among bacterialinterspersed mosaic elements (BIMEs). Mol. Microbiol., 12, 61–70.

20. Bachellier,S., Clement,J.M. and Hofnung,M. (1999) Shortpalindromic repetitive DNA elements in enterobacteria: a survey.Res. Microbiol., 150, 627–639.

21. Stern,M.J., Ames,G.F., Smith,N.H., Robinson,E.C. andHiggins,C.F. (1984) Repetitive extragenic palindromicsequences: a major component of the bacterial genome. Cell, 37,1015–1026.

22. Gilson,E., Bachellier,S., Perrin,S., Perrin,D., Grimont,P.A.D.,Grimont,F. and Hofnung,M. (1990) Palindromic unit highlyrepetitive DNA sequences exhibit species specificity withinEnterobacteriaceae. Res. Microbiol., 141, 1103–1116.

23. Tobes,R. and Ramos,J.L. (2005) REP code: defining bacterialidentity in extragenic space. Environ. Microbiol., 7, 225–228.

24. Aranda-Olmedo,I., Tobes,R., Manzanera,M., Ramos,J.L. andMarques,S. (2002) Species-specific repetitive extragenic

palindromic (REP) sequences in Pseudomonas putida. NucleicAcids Res., 30, 1826–1833.

25. Louws,F.J., Bell,J., Medina-Mora,C.M., Smart,C.D.,Opgenorth,D., Ishimaru,C.A., Hausbeck,M.K., de Bruijn,F.J. andFulbright,D.W. (1998) rep-PCR-mediated genomic fingerprinting:a rapid and effective method to identify Clavibactermichiganensis. Phytopathology, 88, 862–868.

26. Nunvar,J., Huckova,T. and Licha,I. (2010) Identification andcharacterization of repetitive extragenic palindromes (REP)-associated tyrosine transposases: implications for REP evolutionand dynamics in bacterial genomes. BMC Genomics, 11, 44.

27. Ton-Hoang,B., Siguier,P., Quentin,Y., Onillon,S., Marty,B.,Fichant,G. and Chandler,M. (2011) Structuring the bacterialgenome: Y1-transposases associated with REP-BIME sequences.Nucleic Acids Res, 40, 3596–3609.

28. Bertels,F. and Rainey,P.B. (2011) Within-genome evolution ofREPINs: a new family of miniature mobile DNA in bacteria.PLoS Genet., 7, e1002132.

29. Ronning,D.R., Guynet,C., Ton-Hoang,B., Perez,Z.N.,Ghirlando,R., Chandler,M. and Dyda,F. (2005) Active sitesharing and subterminal hairpin recognition in a new class ofDNA transposases. Mol. Cell, 20, 143–154.

30. Garcillan-Barcia,M.P. and de la Cruz,F. (2002) Distribution ofIS91 family insertion sequences in bacterial genomes: evolutionaryimplications. FEMS Microbiol. Ecol., 42, 303–313.

31. Hickman,A.B., Ronning,D.R., Kotin,R.M. and Dyda,F. (2002)Structural unity among viral origin binding proteins: crystalstructure of the nuclease domain of adeno-associated virus Rep.Mol. Cell, 10, 327–337.

32. Guasch,A., Lucas,M., Moncalian,G., Cabezas,M., Perez-Luque,R.,Gomis-Ruth,F.X., de la Cruz,F. and Coll,M. (2003) Recognitionand processing of the origin of transfer DNA by conjugativerelaxase TrwC. Nat. Struct. Biol., 10, 1002–1010.

33. Datta,S., Larkin,C. and Schildbach,J.F. (2003) Structural insightsinto single-stranded DNA binding and cleavage by F factor TraI.Structure, 11, 1369–1379.

34. Koonin,E.V. and Ilyina,T.V. (1993) Computer-assisted dissectionof rolling circle DNA replication. Biosystems, 30, 241–268.

35. Barabas,O., Ronning,D.R., Guynet,C., Hickman,A.B., Ton-Hoang,B., Chandler,M. and Dyda,F. (2008) Mechanism of IS200/IS605 family DNA transposases: activation andtransposon-directed target site selection. Cell, 132, 208–220.

36. Guynet,C., Hickman,A.B., Barabas,O., Dyda,F., Chandler,M. andTon-Hoang,B. (2008) In vitro reconstitution of a single-strandedtransposition mechanism of IS608. Mol. Cell, 29, 302–312.

37. Ton-Hoang,B., Guynet,C., Ronning,D.R., Cointin-Marty,B.,Dyda,F. and Chandler,M. (2005) Transposition of ISHp608,member of an unusual family of bacterial insertion sequences.EMBO J., 24, 3325–3338.

38. Mahillon,J. and Chandler,M. (1998) Insertion sequences.Microbiol. Mol. Biol. Rev., 62, 725–774.

39. Schuck,P. and Rossmanith,P. (2000) Determination of thesedimentation coefficient distribution by least-squares boundarymodeling. Biopolymers, 54, 328–341.

40. Cole,J.L., Lary,J.W., Moody,T.P. and Laue,T.M. (2008)Analytical ultracentrifugation: sedimentation velocity andsedimentation equilibrium. Methods Cell Biol., 84, 143–179.

41. Kabsch,W. (2010) Integration, scaling, space-group assignmentand post-refinement. Acta Crystallogr. D Biol. Crystallogr., 66,133–144.

42. Sheldrick,G.M. (2008) A short history of SHELX. ActaCrystallogr. A, 64, 112–122.

43. Vonrhein,C., Blanc,E., Roversi,P. and Bricogne,G. (2007)Automated structure solution with autoSHARP. Methods Mol.Biol., 364, 215–230.

44. Abrahams,J.P. and Leslie,A.G.W. (1996) Methods used in thestructure determination of bovine mitochondrial F1 ATPase. ActaCrystallogr. D Biol. Crystallogr., 52, 30–42.

45. Jones,T.A. and Kjeldgaard,M. (1997) Electron-density mapinterpretation. Methods Enzymol., 277, 173–208.

46. Jones,T.A., Zou,J.Y., Cowan,S.W. and Kjeldgaard,M. (1991)Improved methods for building protein models in electron densitymaps and the location of errors in these models. Acta CrystallogrA., 47, 110–119.



47. Brunger,A.T., Adams,P.D., Clore,G.M., DeLano,W.L., Gros,P.,Grosse-Kunstleve,R.W., Jiang,J., Kuszewski,J., Nilges,M.,Pannu,N.S. et al. (1998) Crystallography & NMR system: a newsoftware suite for macromolecular structure determination. ActaCrystallogr. D Biol. Crystallogr., 54, 905–921.

48. Murshudov,G.N., Vagin,A.A. and Dodson,E.J. (1997) Refinementof macromolecular structures by the maximum-likelihood method.Acta Crystallogr. D Biol. Crystallogr., 53, 240–255.

49. Brunger,A.T. (1997) Free R value: cross-validation incrystallography. Methods Enzymol., 277, 366–396.

50. Gouet,P., Courcelle,E., Stuart,D.I. and Metoz,F. (1999) ESPript:analysis of multiple sequence alignments in PostScript.Bioinformatics (Oxford, England), 15, 305–308.

51. DeLano,W.L. (2002). The PyMOL Molecular Graphics System.DeLano Scientific, San Carlos, CA, USA.

52. Warner,J.D., DeYoung,P.A., Ellsworth,L.A., Kiessel,L.M.,Rycenga,M.J. and Peaslee,G.F. (2010) Quantitative analysis of ametalloprotein compositional stoichiometry with PIXE and PESA.Nucl. Instrum. Meth. B, 268, 1671–1675.

53. Burd,C.G. and Dreyfuss,G. (1994) Conserved structures anddiversity of functions of RNA-binding proteins. Science, 265,615–621.

54. Clery,A., Blatter,M. and Allain,F.H. (2008) RNA recognitionmotifs: boring? Not quite. Curr. Opin. Struct. Biol., 18, 290–298.

55. Hickman,A.B., James,J.A., Barabas,O., Pasternak,C., Ton-Hoang,B., Chandler,M., Sommer,S. and Dyda,F. (2010) DNArecognition and the precleavage state during single-stranded DNAtransposition in D. radiodurans. EMBO J., 29, 3840–3852.

56. Larkin,C., Datta,S., Nezami,A., Dohm,J.A. and Schildbach,J.F.(2003) Crystallization and preliminary X-ray characterization ofthe relaxase domain of F factor TraI. Acta Crystallogr. D Biol.Crystallogr., 59, 1514–1516.

57. Larkin,C., Datta,S., Harley,M.J., Anderson,B.J., Ebie,A.,Hargreaves,V. and Schildbach,J.F. (2005) Inter- andintramolecular determinants of the specificity of single-strandedDNA binding and cleavage by the F factor relaxase. Structure,13, 1533–1544.

58. Hemavathi,K., Kalaivani,M., Udayakumar,A., Sowmiya,G.,Jeyakanthan,J. and Sekar,K. (2010) MIPS: metal interactions inprotein structures. J. Appl. Crystallogr., 43, 196–199.

59. Harding,M.M. (1999) The geometry of metal-ligand interactionsrelevant to proteins. Acta Crystallogr. D Biol. Crystallogr., 55,1432–1443.

60. Boer,R., Russi,S., Guasch,A., Lucas,M., Blanco,A.G., Perez-Luque,R., Coll,M. and de la Cruz,F. (2006) Unveiling themolecular mechanism of a conjugative relaxase: the structure ofTrwC complexed with a 27-mer DNA comprising the recognitionhairpin and the cleavage site. J. Mol. Biol., 358, 857–869.

61. Lucas,M., Gonzalez-Perez,B., Cabezas,M., Moncalian,G.,Rivas,G. and de la Cruz,F. (2010) Relaxase DNA binding andcleavage are two distinguishable steps in conjugative DNAprocessing that involve different sequence elements of the nic site.J. Biol. Chem., 285, 8918–8926.

62. Monzingo,A.F., Ozburn,A., Xia,S., Meyer,R.J. and Robertus,J.D.(2007) The structure of the minimal relaxase domain of MobA at2.1 A resolution. J. Mol. Biol., 366, 165–178.

63. Goryshin,I.Y. and Reznikoff,W.S. (1998) Tn5 in vitrotransposition. J. Biol. Chem., 273, 7367–7374.

64. Mates,L., Chuah,M.K.L., Belay,E., Jerchow,B., Manoj,N.,Acosta-Sanchez,A., Grzela,D.P., Schmitt,A., Becker,K., Matrai,J.et al. (2009) Molecular evolution of a novel hyperactive SleepingBeauty transposase enables robust stable gene transfer invertebrates. Nat. Genet., 41, 753–761.

65. Yusa,K., Zhou,L., Li,M.A., Bradley,A. and Craig,N.L. (2011) Ahyperactive piggyBac transposase for mammalian applications.Proc. Natl Acad. Sci. USA, 108, 1531–1536.

66. Twiss,E., Coros,A.M., Tavakoli,N.P. and Derbyshire,K.M. (2005)Transposition is modulated by a diverse set of host factors inEscherichia coli and is stimulated by nutritional stress. Mol.Microbiol., 57, 1593–1607.

67. Pasternak,C., Ton-Hoang,B., Coste,G., Bailone,A., Chandler,M.and Sommer,S. (2010) Irradiation-induced Deinococcusradiodurans genome fragmentation triggers transposition of asingle resident insertion sequence. PLoS Genet., 6, e1000799.

68. Dai,J., Xie,W., Brady,T.L., Gao,J. and Voytas,D.F. (2007)Phosphorylation regulates integration of the yeast Ty5retrotransposon into heterochromatin. Mol. Cell, 27, 289–299.

69. Mahnke Braam,L.A., Goryshin,I.Y. and Reznikoff,W.S. (1999) Amechanism for Tn5 inhibition. Carboxyl-terminal dimerization. J.Biol. Chem., 274, 86–92.

70. Allingham,J.S. and Haniford,D.B. (2002) Mechanisms of metalion action in Tn10 transposition. J. Mol. Biol., 319, 53–65.

71. Allingham,J.S., Wardle,S.J. and Haniford,D.B. (2001)Determinants for hairpin formation in Tn10 transposition. EMBOJ., 20, 2931–2942.

72. Duval-Valentin,G. and Chandler,M. (2011) Cotranslationalcontrol of DNA transposition: a window of opportunity. Mol.Cell, 44, 989–996.

73. Montano,S.P. and Rice,P.A. (2011) Moving DNA around: DNAtransposition and retroviral integration. Curr. Opin. Struct. Biol.,21, 370–378.

74. Cesar,C.E., Machon,C., de la Cruz,F. and Llosa,M. (2006) A newdomain of conjugative relaxase TrwC responsible for efficientoriT-specific recombination on minimal target sequences. Mol.Microbiol., 62, 984–996.



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