# The Xer activation factor of TLCΦ expands the possibilities for Xer recombination

Solange Miele, James Iain Provan, Justine Vergne, Christophe Possoz, Françoise Ochsenbein, François-Xavier Barre

► To cite this version:

Solange Miele, James Iain Provan, Justine Vergne, Christophe Possoz, Françoise Ochsenbein, et al. The Xer activation factor of TLCΦ expands the possibilities for Xer recombination. Nucleic Acids Research, 2022, 50, pp.6368 - 6383. 10.1093/nar/gkac429. hal-03872179

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# The Xer activation factor of TLCΦ expands the possibilities for Xer recombination

Solange Miele, James Iain Provan, Justine Vergne, Christophe Possoz, Françoise Ochsenbein and François-Xavier Barre*

Institute for Integrative Biology of the Cell (I2BC), Université Paris-Saclay, CEA, CNRS, 1 Avenue de la Terrasse, 91198Gif-sur-Yvette, France

Received September 28, 2021; Revised May 03, 2022; Editorial Decision May 4, 2022

## ABSTRACT

The chromosome dimer resolution machinery of bacteria is generally composed of two tyrosine recombinases, XerC and XerD. They resolve chromosome dimers by adding a crossover between sister copies of a specific site, *dif*. The reaction depends on a cell division protein, FtsK, which activates XerD by protein-protein interactions. The toxin-linked cryptic satellite phage (TLCΦ) of *Vibrio cholerae*, which participates in the emergence of cholera epidemic strains, carries a *dif*-like attachment site (*attP*). TLCΦ exploits the Xer machinery to integrate into the *dif* site of its host chromosomes. The TLCΦ integration reaction escapes the control of FtsK because TLCΦ encodes for its own XerD-activation factor, XafT. Additionally, TLCΦ *attP* is a poor substrate for XerD binding, in apparent contradiction with the high integration efficiency of the phage. Here, we present a sequencing-based methodology to analyse the integration and excision efficiency of thousands of synthetic mini-TLCΦ plasmids with differing *attP* sites *in vivo*. This methodology is applicable to the fine-grained analyses of DNA transactions on a wider scale. In addition, we compared the efficiency with which XafT and the XerD-activation domain of FtsK drive recombination reactions *in vitro*. Our results suggest that XafT not only activates XerD-catalysis but also helps form and/or stabilize synaptic complexes between imperfect Xer recombination sites.

## INTRODUCTION

Repair by homologous recombination can lead to the formation of chromosome dimers when the chromosomes are circular, as is generally the case in bacteria and archaea. Chromosome dimers physically impede the segregation of genetic information. They are resolved by the addition of a crossover between the sister copies of a specific chromosomal

locus, *dif*, by a highly conserved chromosomally encoded tyrosine recombination (Xer) machinery (1).

Many mobile elements such as plasmids exploit the high conservation of the Xer machinery (1). Indeed, Xer was initially characterized as a multicopy plasmid dimer re-solvase (2,3). Integrative Mobile Elements exploiting Xer (IMEX) have also been reported (4,5). Plasmids and IMEX participate in the acquisition of antibiotic resistance and pathogenicity factors. In particular, the continuous emergence of new pathogenic forms of *Vibrio cholerae* is linked to cycles of integration and excision of several IMEX (6–10). Foremost among those are the Cholera ToXin phage, CTXΦ, which harbours the genes responsible for the deadly diarrhoea associated with cholera (11), and a Toxin-Linked Cryptic satellite phage, TLCΦ (12). It has been proposed that prior integration of TLCΦ is necessary to facilitate the integration of CTXΦ into the *dif* site of the larger of the two chromosomes of its host, chr1 (13). One or more TLCΦ-integration events preceded the integration of CTXΦ in most clinical strains (14–18). For instance, two and three TLCΦ-integration events took place before the integration of CTXΦ on the chr1 of the N16961 clinical isolate from the on-going seventh cholera pandemic and the O395 clinical isolate from the sixth cholera pandemic, respectively (Figure 1A). In both cases, a first copy of TLCΦ integrated into *difA*, the chr1 *dif* site of the clinical isolate environmental ancestor, which led to the formation of a new *dif* site, *dif1*, into which a second copy of TLCΦ integrated, prior to the integration of CTXΦ (Figure 1A and B).

In *V. cholerae*, as in most bacteria, the Xer machinery is composed of two closely related recombinases, XerC and XerD (Supplementary Figure S1A). The *dif* sites are composed of an 11 base pair (bp) XerC-binding arm and an 11 bp XerD-binding arm, which are separated by a short 6 bp central region (Figure 1B and Supplementary Figure S1B). The XerC- and XerD-arms are partial inverted repeats. XerC and XerD each catalyse the cleavage and transfer of a specific pair of DNA sequences within the tetrameric nucleoprotein complex (Figure 1B and C). To resolve chromosome dimers, XerCD follows the conventional recombination

<footnote>*To whom correspondence should be addressed. Tel: +33 169 823 224; Email: francois-xavier.barre@i2bc.paris-saclay.fr</footnote>
<figure><img src="image_2.png" /><figcaption>**Figure 1.** Xer recombination. (A) Schematic representation of the IMEX arrays found on chr1 of a *V. cholerae* strain from the 7th (N16961) and the 6th (O395) pandemics (top) and of the successive integration of TLCΦ leading to the formation of those arrays (bottom). The arrays result from successive lysogenic conversion events, a new *dif* site being generated on the right side of each newly-integrated IMEX. Pink, red, orange blocks: TLCΦ, CTXΦ and RS1 prophages; Coloured disks: *difA* (grey/black), TLCΦ *attP* (green/pink), TLCΦ *attL1* (grey/pink), *dif1* (green/black) and current *dif* site of the strains (black disk). Numbers indicate the order of integration of the phage, which can be directly deduced from their position relative to the current *dif* site. (B) *V. cholerae* *difA* and *dif1*, TLCΦ *attP* and *attL1* sequences. Green: XerC-binding site; Light grey: central region and non-canonical bp of *difA*; Blue: XerD-binding site; Pink: non-canonical bp of *attP*. The top and bottoms are cleaved by XerC and XerD, respectively. The positions of cleavage are indicated by triangles. (C) Xer recombination pathways. XerC and XerD are shown in light and dark grey, respectively. The central region, XerC and XerD-arms of the recombination sites are shaded in grey, green and blue, respectively. Black arrows: conventional recombination pathway; Red arrows: non-conventional asymmetric recombination pathway. The XerD and XerC cleavage points are depicted by empty and filled disks. (D) TLCΦ integration. Black and pink lines: *V. cholerae* chr1 and TLCΦ DNA, respectively. Question mark: the low affinity of XerD for the XerD-arm of TLCΦ *attP* poses questions about the possible formation of integration complexes.</figcaption></figure>

tion pathway of tyrosine recombinases. A first pair of DNA
strands is cleaved and transferred leading to the forma-
tion of a Holliday junction (HJ) intermediate that is subse-
quently resolved into a crossover by the exchange of the sec-
ond pair of strands (Figure 1C). During chromosome dimer
resolution, the exchange of the first pair of strands is catal-
ysed by XerD and the second by XerC (Figure 1C, (19,20)).
This process is under the control of a DNA translocase an-
chored in the cell division septum, FtsK. The C-terminal
γ-domain of FtsK activates XerD by protein-protein in-
teractions (Figure 1C, (19,21–23)). Hence, Xer recombina-
tion is normally restricted to the time of cell division and to
synaptic complexes located in the proximity of the cell divi-
sion apparatus (24–28). Many IMEX including CTXΦ in-

tegrate via a non-conventional FtsK-independent recombi-
nation pathway which is initiated by XerC-mediated strand
exchanges. The resulting HJ is resolved into fully integrated
CTXΦ by DNA replication (Figure 1C, (7,8,29)). Dimers of
the ColE1 multicopy plasmid are resolved by a similar pro-
cess (30). In contrast, TLCΦ integration follows a XerD-
first pathway like chromosome dimer resolution (9), but the
process escapes the control of FtsK control because TLCΦ
encodes for its own XerD activation factor, XafT (Figure
1C, (9,31)).

Eight bp of the XerD-arm of TLCΦ *attP* deviate from the
canonical XerD-arm of *dif* sites (Figure 1B). *In vitro* exper-
iments previously showed that XerD failed to bind TLCΦ
*attP* under conditions in which it efficiently bound the *dif1*
site (9). In addition, XerC/XerD/TLCΦ *attP* complex for-
mation was barely detectable (9). Those observations raised
questions about how XerC and XerD could promote the
previously observed highly efficient integration of TLCΦ *in
vivo* (Figure 1D, (9,13,31)).

Here, we present a methodology based on deep sequenc-
ing that permits the *in vivo* assessment of the integration
and excision frequencies of thousands of mobile elements
with differing attachment sites. This methodology was ap-
plied to investigate the XerD-arm sequence-dependence of
TLCΦ *attP* during Xer-mediated recombination reactions,
in either FtsK- or XafT-driven conditions. By *in vitro* ex-
periments we also compared the efficiency of XafT-driven
and FtsKγ-driven Xer recombination reactions between
two *dif1* sites, and between *dif1* and TLCΦ *attP*. Taken to-
gether, our results suggest that XafT is not only a XerD-
activation factor, but also a synapse formation and/or sta-
bilization factor. Our results also indicate that undesired ex-
cision events are prevented because the expression of XafT
is limited after integration.

MATERIALS AND METHODS

Strains, plasmids and oligonucleotides

Strains, plasmids and oligonucleotides are listed in Sup-
plementary Tables S1, S2 and S3, respectively. *V. cholerae*
strains were built by natural transformation using appro-
priate selection markers and/or blue/white β-galactosidase
screens (32). Plasmids pSM11 and pSM15 are derivatives of
the pTLC8 and pSM12 non-replicative mini-TLCΦ, respec-
tively (31). They were constructed by replacing the TLCΦ
*attP* locus with the BsaI-*ccdB*-BsaI cassette of pFB5 (33) us-
ing Gibson assembly (34). Pools of n<sub>8</sub>gtg, tagn<sub>8</sub> and n<sub>5a2</sub>n<sub>3g</sub>
TLCΦ *attP*-derived sites flanked by cognate BsaI sites were
generated by primer extension of oligo 4069 annealed to
degenerate 4066, 4067, 4068 oligonucleotides, respectively.
The degenerate *attP* sites flanked by BsaI sites were inserted
in place of the BsaI-*ccdB*-BsaI cassette of the XafT<sup>+</sup> pSM11
and XafT<sup>-</sup> pSM15 mini-TLCΦ plasmids by restriction di-
gest and ligation. The plasmid libraries were transformed
into an *E. coli* *xerC*<sup>-</sup> strain (FCV14). Transformation re-
actions were plated on 20 cm diameter petri dishes and re-
peated to obtain a total of about 10<sup>6</sup> colonies. Plasmid li-
braries were then extracted from ~1 × 10<sup>10</sup> FCV14 cells
using a plasmid midi-prep kit (Qiagen), and re-transformed
into the *E. coli* conjugation donor strain β2163. pSM11 and
pSM15 derivatives harbouring *dif1* in place of TLCΦ *attP*,
pSM17 and pSM18, were constructed by the same tech-
nique, respectively. The use of a Type IIS restriction digest
and ligation-based process ensured that the mini-TLCΦ
plasmids from the XafT<sup>-</sup> and XafT<sup>+</sup> libraries only differed
from pTLC8 and pSM12 by the sequence of the XerD-arm
of their attachment site.

Integration assays

To monitor integration, engineered *V. cholerae* strains with
arabinose-inducible XerCD expression and integration-
reporter *dif1*-LacZ constructs were used. CMV30 (*lacZ-dif1*
reporter at the *dif1* locus) and EPV369 (*lacZ-dif1* reporter

at the *lacZ* locus) recipient cells were cultured to an opti-
cal density (OD 600 nm) of 0.3 in L-broth supplemented
with 0.2% of L-arabinose (L-ara) to induce the expression
of XerD and XerC. The *Escherichia coli* β2163 conjuga-
tion donor cells containing the mini-TLCΦ libraries were
grown to an OD600 of 0.6 in LB supplemented with 0.3 mM
of diaminopimelic acid (*dap*). β2163 is a diaminopimelic
acid auxotroph (*dap*<sup>-</sup>) which allows for simple counters-
election after conjugation. *E. coli* donor and *V. cholerae*
recipient cells were mixed in a 1:10 ratio and incubated
for conjugation at 37°C for 3 h on LB agar plates supple-
mented with *dap* and L-ara. The conjugation mixtures were
subsequently resuspended in LB supplemented with 0.2%
of L-ara and incubated for an additional 1h of liquid cul-
ture. Serial dilutions of the conjugation mix were plated
on LB agar with X-gal to determine the total number of
*V. cholerae* conjugants (blue *dap*<sup>+</sup> clones), and on LB agar
supplemented with Chloramphenicol (Cm) and X-gal to de-
termine the number of specific integration events into *dif1*
(white *dap*<sup>+</sup> *Cm*<sup>R</sup> clones). L-ara was omitted from the in-
tegration plates to limit the possibility of excision of the
mini-TLCΦ plasmid during colony formation. The global
frequency of integration of mini-TLCΦ plasmids carrying
*dif1*, TLCΦ *attP*, and degenerate XerD-arm mini-TLCΦ
plasmid libraries was calculated as the ratio between the
number of specific integrants to the number of conjugants.
Integrant libraries were harvested for NGS by scraping the
plates after overnight growth and the recovered cells were
subsequently stored at -80°C. Over 10 conjugations were
performed for each library condition and were pooled to-
gether for sequencing or excision assays.

Excision assays

To analyse the excision of integrated *attP* library variants,
the stored integration libraries were re-inoculated into liq-
uid culture without antibiotic selection, thus the retention
of the integrated elements was not actively encouraged.
About 10<sup>9</sup> frozen cells from the integration libraries were
precultured for 2 h in 100 ml of LB. One ml of the preculture
was then used to seed 200 ml of LB supplemented with 0.2%
of arabinose to induce the expression of XerD and XerC.

NGS analysis

The *attP* sites of the donor plasmid libraries were ampli-
fied by performing 17 PCR cycles on 50–100 ng of midiprep
plasmid DNA with an equimolar mix of 4103–4105 P5
and 4174–4176/4243–4245 P7 primers (Table S4), which
bind on the TLCΦ genome and the plasmid backbone, re-
spectively. The genomic DNA from *V. cholerae* integration
and excision libraries was recovered using a gDNA extrac-
tion kit (GenElute Bacterial Genomic DNA kit, Sigma).
The *attP* sites specifically integrated into *dif1* were then
amplified by performing 17 PCR cycles on 50–100 ng of
gDNA with an equimolar mix of 4103–4105 P5 and 4178–
4180/4246–4248 P7 primers. The 4178–4180/4246–4248 P7
primers bind in *E. coli* *lacZ*, which ensures that we only
analysed the sequence of non-replicative mini-TLCΦ plas-
mids specifically integrated inside the *lacZ-dif1* reporter
cassette. The products were purified from the P5 and P7
primers by double selection with AMPure beads (Beckman). NGS was performed on an Illumina NextSeq 500 machine. NextSeq reads were trimmed with Cutadapt (version 1.17). MatLab scripts were developed to analyse results, draw 2D-maps, butterfly plots, violin plots, bar plots, and sequence frequency logos, and are available upon request. The relative integration frequency of a mini-TLCΦ plasmid was calculated as the ratio of the frequency of its *attP* motif in the integration library, *r*<sub>int</sub>(*motif*), to the frequency of the *attP* motif in the donor library, *r*<sub>lib</sub>(*motif*). The integration efficiency, *E*<sub>exc</sub>(*motif*), was then obtained by multiplying the relative integration frequencies by the global integration frequency of the corresponding library, *f*<sub>int</sub>(*library*). The retention efficiency of a motif after growth in conditions permitting its excision, *E*<sub>exc</sub>(*motif*), was calculated as the ratio of the frequency of the motif after growth in the excision conditions, *r*<sub>exc</sub>(*motif*), to its frequency in the integration library, *r*<sub>int</sub>(*motif*). It was used as an indicator of the stability of the corresponding plasmid.

### In vitro recombination assays

Short *difl* substrates were obtained by annealing oligonucleotides 3418 and 3419 (Table S4). Oligo 3418 possessed 5'-CY5 and 3'-CY3 fluorophores. The 152bp *difl* and *attP* substrates were produced by PCR using 2206/3419 and 2470/1110 primer couples on pSM17 and pTLC8, respectively. Recombination reactions were performed in 25mM Tris-HCl pH7.5, 1 mM EDTA, 10% glycerol, 100 mM NaCl and 100 ng/ml BSA. Reactions were incubated for 2 h at 37°C. To remove the reaction proteins, samples were treated with 0.5% SDS and 0.4 mg/mL proteinase K for 1 h at 37°C. Recombination reactions were separated by electrophoresis on native 5% 29:1 acrylamide gels containing TBE 0.5X buffer for a period of 80 min at 17 mA. Gels were fluorescently imaged with a Typhoon laser scanner (GE healthcare) using filters 670BP30 and 580BP30.

### XafT immunoblotting

Expression profiles of XafT from either integrated or replicative forms of pTLC plasmids were performed in *V. cholerae* EPV369 (*lacZ*-difl reporter at the *lacZ* locus) by immunoblotting against XafT with C-terminal 3xFLAG epitopes. Clones harbouring integrated or replicative forms of the pTLC variants were selected on LB-agar supplemented with Cm and X-gal, with integrated forms giving rise to white colonies, and non-integrated replicative forms giving rise to blue colonies. Total cellular protein extracts were separated on precast 8–16% gradient acrylamide TGX ‘Stain-Free’ gels (Bio-Rad) in Tris-glycine–SDS electrophoresis buffer. Gels were electroblotted by wet-transfer onto nitrocellulose. The following antibodies were used for immunoprobing: 1:10 000 anti-FLAG M2 (F1804, Sigma), and 1:10 000 anti-mouse IgG-HRP (A9044, Sigma), both diluted in 5% fat-free milk TBS-T (20 mM Tris pH 7.6, 150 mM NaCl, 0.01% Tween-20). Chemiluminescence was performed using SuperSignal™ West Pico PLUS (ThermoFisher) ECL, and captured over a 60-min exposure with a ChemiDoc imager (Bio-Rad). Supplementary Figure S6 contains uncropped source images of the original SDS-PAGE, transfer, and immunoblot. Supplementary method

contains an extended explanation of the plasmid construction and sample preparation used for this immunoblot.

### Protein purification

MBP-XafT, MBP-XerD, MBP-XerD-FtsKγ and MBP-XerC fusions were produced in *E. coli* BL21(DE3) cells from plasmids pCM153, pCM162, pSM20 and pCM157, respectively. The AA linker between the C-terminus of MBP and the N-terminus of the proteins of interest contained a 6xHIS tag and a TEV protease cleavage site. Cultures were grown at 37°C under agitation until exponential phase, then induced with 1 mM IPTG. Induction was performed at 20°C overnight under agitation. Induced cultures were pelleted, washed in LB and lysed with a French press in presence of protease inhibitors (Merck, cOmplete EDTA free), 1 M NaCl (or 200 mM for XerD), 1 mM EDTA, 50 mM Tris–HCl pH 7. The following steps were all performed at 4°C. Lysates were centrifuged at 34 000 g, then the supernatants were removed and passed though 0.45 µm and 0.2 µm filters. FPLC purification was performed using an AKTA purifier (GE Healthcare). The MBP fusion proteins were bound on 5 ml MBPTrap columns (GE Healthcare) in a solution of 50 mM Tris pH 7, 0.2 M NaCl, 1 mM EDTA. MBP fusions were eluted with in the same buffer supplemented with 10 mM maltose. Purified MBP-XerD, MBP-XerD-FtsKγ and MBP-XerC proteins were digested overnight with TEV protease to cleave their MBP-6xHis tags. Finally, MBP-XafT, XerD, XerD-FtsKγ and XerC were purified and concentrated using 5 ml HiTrap Heparin HP columns (GE Healthcare). The heparin affinity purification step removed cleaved MBP-6xHis moieties from the XerD, XerD-FtsKγ and XerC protein preparations. Protein aliquots were stored at –80°C.

## RESULTS

### NGS-based comparison of the efficiency of integration and excision of mobile elements

A classical methodology to analyse the mechanisms of integration of a mobile element is to monitor the frequency of integration, *f*<sub>int</sub>, of wild-type or genetically modified versions of the element in genetic mutants of its host. XafT was discovered by this approach (31). In brief, a non-replicative version of TLCΦ, lacking the coding sequence for the native TLCΦ phage replication protein Cri, was cloned into a plasmid backbone (pSW23T) that has a conditional R6K origin of replication and the *cat* chloramphenicol (Cm) resistance gene (Figure 2A, (35)). pSW23T derivatives will only replicate in bacterial strains possessing a chromosomally encoded R6K-replication protein pir (π). These conditionally-replicative ‘mini-TLCΦ’ plasmids were delivered to a π⁻ Cm-sensitive N16961 *V. cholerae* strain by conjugation using a π⁺ *E. coli* donor strain which requires diaminopimelic acid (dap) for growth. Conjugation mixtures were then plated onto LB agar lacking dap (Figure 2A, (35)). As the *V. cholerae* strain lacks π, which is necessary for replication of the mini-TLCΦ plasmids, and as the *E. coli* donor strain is a dap auxotroph, any Cm-resistant colonies correspond to the integration of a mini-TLCΦ plasmid in the genome of a *V. cholerae* recipient cell. This permits the assessment
<figure><img src="image_3.png" /><figcaption>**Figure 2.** Parallel monitoring of the integration and excision of Mobile DNA elements. (A) Schematic of classical and NGS based strategies to monitor integration and excision efficiencies. 'Xer on': growth in the presence of arabinose. Pink: TLCΦ DNA; Grey: plasmid vector DNA.; 6: R6Kγ replication origin; cat: chloramphenicol resistance gene; T: transfer origin; dap: diaminopimelic acid; Cm: chloramphenicol; ara: L-arabinose; Xgal: X-gal; lacZ-dif1 reporter: E. coli lacZ gene containing an internal dif1 site. Plasmid- and lacZ-specific P5, and TLCΦ-specific P7 adaptor primers used for next generation sequencing (NGS) are indicated by black, grey and pink arrows, respectively. (B) Scheme of the *V. cholerae* reporter strains. White circle: chromosome 1 replication origin; red, blue and black crosses: *dif1* and adjoining IMEX deletion, *xerD* deletion and *V. cholerae lacZ* deletion, respectively; black rectangle: *E. coli lacZ-dif1* reporter; light and dark grey rectangles: synthetic *xerC* and *xerD* operon under the control of the arabinose promoter (Para). Yellow shading depicts the subcellular region and timing of activity of FtsK during the cell cycle. (C) Non-replicative Mini-TLCΦ plasmid libraries. Top panel: top strand of the XerD-arm of the degenerate *attP* motifs. Sequence legend as in Figure 1B. Bottom panel: scheme of XafT⁺ and XafT⁻ mini-TLCΦ plasmids. Yellow rectangle: *xafT* gene; sawed lines: stop mutation.</figcaption></figure>

of the fraction of Cm-resistant cells to the total number of recipient cells, *f*<sub>int</sub>, by spreading serial dilutions on plates lacking *dap* and supplemented or not with Cm, respectively (Figure 2A).

In our previous work we have also established a robust colorimetric assay to distinguish specific integration events from random integration events, based on the classical

Blue/White LacZ (β-galactosidase) phenotype of bacterial cells on L-agar supplemented with X-gal. The colorimetric assay is based on the insertion of the *dif1* recombination site within the coding sequence of the *E. coli lacZ* (*eclacZ*) gene in such a manner as not to perturb the β-galactosidase activity of the reporter gene product. The reporter is hereafter denoted as the ‘lacZ-dif1’ reporter. It has been utilised in several ΔlacZ *V. cholerae* strain backgrounds (Δ*vc*lacZ). These include placing the lacZ-dif1 reporter at the normal chr1 *dif1* locus (*lacZ-dif1* at *dif1*) where recombination of the site will be subject to regulation by FtsK, or at exogenous loci such as replacing *vc*lacZ in a Δdif1 background (*lacZ-dif1* at *vc*lacZ) where the recombination site reporter will be outside the domain of action of FtsK (Figure 2B, (7)). The sensitivity of this technique allows us to monitor *f*<sub>int</sub> as low as 10<sup>-6</sup>.

The classical methodology is simple and reliable, but cannot be reasonably applied to compare the efficiency of integration of thousands of different mobile elements. In addition, calculating integration rate by plating 10-fold serial dilutions has an intrinsic ten-fold precision limit. This leads to a 100-fold precision limit when estimating the fraction of integrants in the recipient population because the total number of cells is also measured using 10-fold dilutions. To solve those issues, we designed a high-throughput methodology based on Next Generation Sequencing (NGS), which we applied to libraries of conjugative non-replicative mini-TLCΦ plasmids carrying *attP* sites with different XerD-arm sequences. The relative frequency of each of the different XerD-arm sequence motifs in the plasmid libraries, *r*<sub>lib</sub>(*motif*), was determined by NGS using a primer binding in the TLCΦ genome and a primer binding in the pSW23T vector backbone (Figure 2A). The plasmid libraries were then delivered to *V. cholerae* recipient cells in which the *V. cholerae lacZ* gene, the wild-type IMEX arrays, and the original *dif1* site were deleted, and which were equipped with the *lacZ-dif1* reporter. Cm-resistant *V. cholerae* colonies were collected on large selection plates and pooled. The relative frequency of each of the different XerD-arm sequence motifs in the Cm-resistant cell pools, *r*<sub>int</sub>(*motif*) was determined by NGS (Figure 2A). PCR amplification using a primer binding in the TLCΦ genome in combination with a primer binding in *lacZ* ensured that only *dif1*-specific integration events were taken into account. The efficiency of integration of a plasmid with a given XerD-arm motif, *E*<sub>int</sub>(*motif*), was estimated as the ratio of *r*<sub>int</sub>(*motif*) to *r*<sub>lib</sub>(*motif*). It was multiplied with the global frequency of integration of the whole library, *f*<sub>int</sub>(library) to take into account the influence of the host genetic context (Figure 2A).

The NGS approach can also be used to monitor the retention efficiency of thousands of different integrated mobile elements, provided that the enzymes catalysing their excision are conditionally expressed. This is particularly interesting in the case of IMEX since they integrate into a locus controlled by the normal domain of action of FtsK, which could also lead to FtsK-driven excision events. To this end, we used *V. cholerae* reporter cells that were genetically modified to place the production of XerC and XerD under the control of the arabinose inducible promoter (Figure 2A, (28)). We then determined the relative frequency
of each of the different XerD-arm sequences in the plasmids that specifically integrated at *dif1* after growth in conditions that permitted their excision, *r*<sub>exc</sub>(*motif*), i.e. in the absence of selection and in the presence of arabinose for XerC and XerD production (Figure 2A). The influence of each XerD arm motif on the retention efficiency of the integrated mini-TLCΦ plasmids, *E*<sub>exc</sub>(*motif*), was assessed by calculating the ratio of *r*<sub>exc</sub>(*motif*) to *r*<sub>int</sub>(*motif*) (Figure 2A). As the integration of TLCΦ in *dif1* results in the formation of a prophage flanked by *attP* and *dif1*, *E*<sub>int</sub>(*motif*) and *E*<sub>exc</sub>(*motif*) reflected the efficiency of recombination reactions between the same two sites (Figure 1A).

For the purpose of this study, we generated three pools of 65 536 (4⁸) different TLCΦ *attP* sites using synthetic oligonucleotides carrying eight degenerate bases at different positions of the XerD-arm as templates (Figure 2C). Two of the pools were designed to explore the influence of the eight innermost and outermost positions of the XerD-arm. They are referred to as n₈gtg and tagn₈ on the basis of the sequence of their top strand. We added to the study a pool harbouring degenerate bases in both the inner and outer part of the XerD-arm, n₅a₂n₃g. The three pools of sequences were cloned in place of the *attP* site of XafT⁺ and XafT⁻ mini-TLCΦ plasmids (Figure 2C). NGS analysis showed that each resulting mini-TLCΦ library contained over 99.5% of the XerD-arm sequence motifs that was expected to be included by the pool from which it originated, with a median copy number in the order of 15 per million reads (Supplementary Figure S2). The XafT⁺ and XafT⁻ mini-TLCΦ libraries contained over 99.9% and 99.8% of the 190 528 XerD-arm sequence motifs included in the three combined degenerate pools, respectively.

## XafT expands the number of XerD-arm motifs that can be used for integration

We first explored the influence of the XerD-arm sequence of TLCΦ *attP* on the efficiency of integration in an FtsK-driven context. To this end, we conjugated the XafT⁻ mini-TLCΦ libraries in a *V. cholerae* recipient strain in which *lacZ-dif1* reporter was inserted at the *dif* locus of chr1 (strain CMV30, Figure 2B, (9,28)). FtsK promoted the integration of the XafT⁻ mini-TLCΦ plasmid harbouring *dif1* at a very high frequency (Figure 3A, FtsK panel). However, the integration of the XafT⁻ mini-TLCΦ plasmid harbouring the wild-type TLCΦ *attP* was below the detection level. The global integration frequency of the XafT⁻ n₈gtg, tagn₈ and n₅a₂n₃g mini-TLCΦ libraries was 1000-fold lower than that of the XafT⁻ mini-TLCΦ plasmid harbouring *dif1* (Figure 3A, FtsK panel). NGS revealed that FtsK/XerC/XerD had promoted the integration of only 2.5% of the total number of XerD-arm motifs present in the combined n₈gtg, tagn₈ and n₅a₂n₃g libraries (Figure 3A, FtsK panel). We generated the sequence logos of the integrated sites from each library (Supplementary Figure S3) and a sequence logo combining the 4729 XerD-arm sequence of the integrated mini-TLCΦ plasmids of the three n₈gtg, tagn₈ and n₅a₂n₃g libraries (Figure 3B, FtsK panel). The sequence logos correspond to the top strand of the XerD-arm. The 5' end of the sequence logos is adjacent to the central region. We refer to it as the innermost side of the

XerD-arm (Figure 3C). We refer to the 3' end of the logos as the outermost side of the XerD-arm (Figure 3C). The logos revealed that the more similar the degenerate *attP* sequence was to *dif1*, the higher the integration frequency was in the Ftsk-driven context (Figure 3B and Supplementary Figure S3, FtsK panels). It was particularly evident for the innermost bp of the XerD-arm, but the outermost bp showed the same trend (Figure 3B and Supplementary Figure S3, FtsK panels).

We next explored the influence of the XerD-arm sequence of TLCΦ *attP* on the efficiency of integration in a XafT-driven context. To this end, we conjugated the XafT⁺ mini-TLCΦ libraries in the *V. cholerae* recipient strain in which *lacZ-dif1* was inserted at the *V. cholerae* lacZ locus (strain EPV369, Figure 2B). Our previous work had shown that the choreography of chromosome segregation restricts the activity of FtsK to the terminus region of the two *V. cholerae* chromosomes (Figure 2B, (28,36–38)). Correspondingly the integration of a XafT⁻ plasmid harbouring *dif1* was found to be 100-fold less efficient when the chromosomal *dif1* site was located at its natural locus (9). Thus, the frequency of integration of a XafT⁺ mini-TLCΦ plasmid in a *V. cholerae* recipient strain harbouring *lacZ-dif1* at the *lacZ* locus can be mainly attributed to the activation of Xer recombination by XafT. The integration of the XafT⁺*dif1* and XafT⁺ TLCΦ *attP* plasmids at the *lacZ* locus was as efficient as the integration of the XafT⁻*dif1* plasmid at the *dif1* locus (Figure 3A, XafT panel). In addition, the global integration frequency of the XafT⁺ n₈gtg, tagn₈ and n₅a₂n₃g mini-TLCΦ plasmid libraries was now only 10-fold lower than that of the XafT⁺*dif1* plasmid (Figure 3A, XafT panel). NGS showed that 47579 different mini-TLCΦ plasmids had integrated, which corresponds to 25% of the total number different plasmids present in the combined n₈gtg, tagn₈ and n₅a₂n₃g mini-TLCΦ libraries (Figure 3B, XafT panel). The sequence logos of the first six positions of the sites integrated via XafT-driven recombination were statistically different from those of the sites integrated via FtsK-driven recombination (Supplementary Figure S3, XafT panels). The nature of the bases of the XerD-arm was less important for XafT-driven than for FtsK-driven integration (Figure 3B and Supplementary Figure S3, XafT panels).

Next, we explored the integration of the XafT⁺ mini-TLCΦ libraries in the *V. cholerae* recipient strain equipped with *lacZ-dif1* at the natural *dif1* locus, i.e. in a context in which both XafT and FtsK can act (strain CMV30, Figure 2B). The results were remarkably similar to those obtained for the integration of the XafT⁺ mini-TLCΦ plasmids in the strain carrying *lacZ-dif1* at the *lacZ* locus (Figure 3 and Supplementary Figure S3). The only notable difference was a slightly higher global integration frequency (Figure 3A, FtsK & XafT panel) and greater number of different integrated XafT⁺ mini-TLCΦ plasmids (Figure 3B, FtsK & XafT panel). This could be explained by the growth advantage of the reporter strain carrying *lacZ-dif1* at its natural locus over the reporter strain carrying it at the *lacZ* locus, since the latter is *Δdif1* and therefore cannot resolve chr1 dimers (20). Nevertheless, the XerD-arm sequence logos of the XafT⁺ mini-TLCΦ plasmids integrated at the *dif1* locus were similar to those of the XafT⁺ mini-TLCΦ plasmids
<figure><img src="image_4.png" /><figcaption>**Figure 3.** Non-replicative mini-TLCΦ plasmid integration. (A) Integration frequencies of mini-TLCΦ plasmids harbouring *dif1* or TLCΦ *attP*, and global integration frequencies of n<sub>8</sub>gtg, n<sub>5a2n3g</sub> and tagn<sub>8</sub> plasmid libraries (*f*<sub>int</sub>). FtsK panel: XafT<sup>-</sup> plasmids conjugated in a strain harbouring *dif1* at its natural locus; XafT panel: XafT<sup>+</sup> plasmids conjugated in a strain harbouring *dif1* at the *lacZ* locus; FtsK & XafT panel: XafT<sup>+</sup> plasmids conjugated in a strain harbouring *dif1* at its natural locus. Mean and standard deviations of at least 3 independent assays. (B) Combined sequence of the XerD-arm from the n<sub>8</sub>gtg, n<sub>5a2n3g</sub> and tagn<sub>8</sub> integration libraries. The logos show the frequency of each base at the degenerate positions. The number of integrated motifs top which each logo corresponds is indicated. Individual logos are shown in Supplementary Figure S3. (C) Illustration of our naming scheme for bases of the XerD-binding arm.</figcaption></figure>

integrated at the *lacZ* locus (Figure 3B and Supplementary Figure S3).

Overall, FtsK-driven integration seemed restricted to the *attP* sites that most resembled *dif1*, with the inner positions of the XerD-arm playing a more important role than the outer positions, whereas XafT seemed to allow the integration of plasmids harbouring *attP* sites that significantly deviated from *dif1* at any given position of the XerD-arm.

**The first and fifth innermost bp of the XerD-arm are crucial for integration**

Sequence logos allow a quick interpretation of pools of motifs, but the information that can be derived from them is limited. In particular, it is not possible to determine whether a logo results from the combination of motifs deviating from a single specific consensus sequence, or from the overlap of two or more distinct consensus sequences. For instance, the highly degenerate sequence logos of the XafT and FtsK & XafT integration libraries could be due to the addition of motifs slightly deviating from *dif1* along with motifs slightly deviating from TLCΦ *attP* (Figure 3B). In addition, the relative influence of each particular bp on the efficiency of a process cannot be assessed with a sequence logo. The higher frequency of integration obtained with XafT<sup>+</sup> mini-TLCΦ plasmids compared to XafT<sup>-</sup> mini-TLCΦ plasmids could not be simply explained by the higher number of different XerD-arm sequences that could be integrated (Figure 3B).

We developed a new two-dimensional map (2D-map) graphical representation to avoid such limitations. In these maps, each recovered XerD-arm motif is represented by a small coloured square at specific x and y coordinates based on its sequence (Figure 4A). Each square is coloured on the basis of the integration efficiency of the plasmids harbouring

the corresponding motif. Colours range from black to bright yellow, with black, dark blue and bright yellow corresponding to non-recovered motifs, motifs with the poorer integration efficiency, and motifs with higher integration efficiency, respectively.

The low density of coloured positions highlighted the limited number of XerD-arm sequences that could be integrated by FtsK (Figure 4B, Supplementary Figures S4A and S4B, FtsK 2D-maps). A single small green tile was visible on the n<sub>8</sub>gtg 2D-map, which corresponded to the 64 TTATGNNNNGTG top strand sequences. It demonstrated the importance of first 5 inner positions of the XerD-arm for FtsK-driven integration (Figure 4A, zoom on the ttatn<sub>4</sub>gtg region). Results obtained with the n<sub>5a2n3g</sub> and tagn<sub>8</sub> libraries confirmed this observation (Supplementary Figure S4A and S4B, FtsK panels). The n<sub>8</sub>gtg, n<sub>5a2n3g</sub> and tagn<sub>8</sub> 2D-maps also revealed that FtsK could promote the integration of a plasmid harbouring TLCΦ *attP*, albeit at a low integration frequency, which demonstrates the utility of combining the NGS approach with a 2D-map graphical representation (Figure 4B, Supplementary Figures S4A and B, zooms).

The 2D-maps of the n<sub>8</sub>gtg motifs recovered in the XafT and FtsK & XafT integration libraries presented a very striking checkered pattern (Figure 4B, Supplementary Figures S4A and B, XafT and FtsK & XafT panels). The black stripes (indicating sequences that were not recovered) of the n<sub>8</sub>gtg and n<sub>5a2n3g</sub> 2D-maps correspond to DNA motifs that lack a G at the fifth position of the top strand of the XerD-arm (Figure 4B and Supplementary Figure S4A, XafT and FtsK & XafT 2D-maps). The pattern of the tagn<sub>8</sub> 2D map fitted with the importance of the base at this position (Supplementary Figure S4B, XafT and FtsK & XafT panels). To confirm the importance of the fifth bp
<figure><img src="image_5.png" /><figcaption>**Figure 4.** 2D-maps representation of the integration results. (A) 2D-map representation methodology: Different [x, y] coordinates are assigned to each nucleotide for the 2⁸ possible motifs as follows: the [x, y] coordinates are initially set to [1, 1]. Then, [±1/2n, ±1/2n], [−1/2n, ±1/2n], [−1/2n, −1/2n] or [±1/2n, −1/2n] are added to [x, y] for each n position of the degenerate motif if the nth base is A, T, G or C, respectively. Relative integration efficiencies are represented with a dark-blue to bright yellow colour scale. Positions that were not found in the integration libraries are shown in black. (B) 2D-maps of the integrated n<sub>8</sub>gtg motifs. FtsK: results from the conjugation of the n<sub>8</sub>gtg XafT<sup>−</sup> mini-TLCΦ library in the strain harbouring *lacZ-dif1* at the natural *dif1* locus; XafT: results from the conjugation of the n<sub>8</sub>gtg XafT<sup>+</sup> mini-TLCΦ library in the strain harbouring *lacZ-dif1* at *lacZ* locus; FtsK & XafT: results from the conjugation of the n<sub>8</sub>gtg XafT<sup>+</sup> mini-TLCΦ library in the strain harbouring *lacZ-dif1* at the natural *dif1* locus. Below each 2D-maps are shown zooms of the regions corresponding to the 2<sup>4</sup> motifs that carry the same first innermost bases as the XerD arm of TLCΦ *attP* (*tagan*<sub>4gtg</sub>) and *dif1* (*ttatn*<sub>4gtg</sub>). The position of the TLCΦ *attP* XerD motif and of the motif most similar to the XerD arm of *dif1* are highlighted by white and red squares, respectively. Their sequences are indicated below the panels. Bases identical to the XerD-arms of TLCΦ *attP* and *dif1* are written in blue and red, respectively. 2D-maps of the integrated n<sub>5a2n3g</sub> and n<sub>8</sub> motifs are shown in Supplementary Figure S4. (C) Sequence logo of the XerD-arm of the *attP* sites of the n<sub>8</sub>gtg XafT<sup>+</sup> mini-TLCΦ plasmids that were found integrated in *lacZ-dif1* at *lacZ* locus. The total number of n<sub>8</sub>gtg integrated motifs is indicated. 2D-maps of the integrated n<sub>4tn3gtg</sub>, n<sub>4an3gtg</sub>, n<sub>4gn3gtg</sub> and n<sub>4cn3gtg</sub> motifs, as indicated. The number of integrated motifs from each category is indicated. The sequence logo the XerD-arm of the integrated sites from each category is shown below each 2D-map. Positions in which the bases of the integrated n<sub>4tn3gtg</sub>, n<sub>4an3gtg</sub>, and n<sub>4cn3gtg</sub> XerD-arm motifs are significantly different from the integrated n<sub>4gn3gtg</sub> XerD-arm motif are indicated by a star (chi²-test, P-value < 0.001).</figcaption></figure>

of the XerD-arm for XafT-driven integration, we grouped
the n<sub>8</sub>gtg motifs of the XafT<sup>+</sup> n<sub>8</sub>gtg integration library
on the basis of their fifth base (Figure 4C). The 2D-maps
showed that most of the different possible n<sub>4</sub>gn<sub>3</sub>gtg mo-
tifs were recovered, while the recovered n<sub>4</sub>tn<sub>3</sub>gtg, n<sub>8</sub>an<sub>3</sub>gtg
and n<sub>4</sub>cn<sub>3</sub>gtg motifs were mainly limited to those that car-
ried a T at the first and second innermost positions of the
XerD-arm. It further suggested that C was the most detri-

mental nucleic acid at the fifth position, which needed to
be compensated for by the presence of the same bases as
dif1 at the four innermost positions. Correspondingly, base
frequencies at each of the seven positions of the n4gn3gtg
and n4cn3gtg motif lists were statistically different (Figure
4C). The presence of A or T at the fifth position was less
detrimental. Yet, the n4an3gt and n4tn3gt motifs statistically
differed from the n4gn3g motifs at all positions except the
third and the eighth innermost positions, respectively (Figure 4C).

The 2D-maps of the XafT and FtsK & XafT integration libraries also showed the influence of the first innermost bp of the XerD-arm: yellow-coloured motifs were for the most part restricted to the top left quarter of the n<sub>8</sub>gtg and n<sub>5a2n3g</sub> 2D-maps, which indicates the presence of a T at the first position of the top strand of the XerD-arm (Figure 4B and Supplementary Figure S4A, XafT and FtsK & XafT panels). In addition, half of the plasmids harbouring an *attP* site with a tagn<sub>8</sub> XerD-arm motif were recovered in the XafT and FtsK & XafT libraries, further highlighting the importance of a T at the first innermost bp of the XerD-arm for efficient integration (Supplementary Figure S4B).

Taken together, those results suggest that the highly degenerate XerD-arm sequence logo of the XafT and FtsK & XafT integration libraries derives from the superposition of two categories of motifs, those harbouring a G at the fifth position and those harbouring a T at the first position (Figure 3B).

### XafT permits the highly efficient integration of sites deviating from *difl*

XafT allows the integration of plasmids harbouring *attP* sites deviating from *difl* (Figure 4 and Supplementary Figure S4). Yet, the efficiency of integration of the sites that most resembled *difl* was higher than the efficiency of integration of the sites that most resembled TLCΦ *attP* (Figure 4B, Supplementary Figures S4A and 4B zooms). To further highlight differences of integration efficiency, we developed a second graphical representation of the data obtained by the NGS approach, which we refer to as butterfly plots. In this representation, recovered sites are separated in different bins as a function of the number of bases that differentiate them from TLCΦ *attP* (Figure 5A). For the n<sub>8</sub>gtg, n<sub>5a2n3g</sub> and tagn<sub>8</sub> degenerate sites we studied, it corresponds to eight bins. Each bin was further divided into two (+) and (−) bins, corresponding to the sites that were more similar to *difl* than TLCΦ *attP* or not, respectively (Figure 5A). We then plotted the integration efficiency of each motif from the different bins.

Only a third (21 070/ 65 536) of the possible n<sub>8</sub>gtg motifs are closer to *difl* than to TLCΦ *attP*. Nevertheless, over 99% of the plasmids whose integration could be driven by FtsK harboured an n<sub>8</sub>gtg XerD-arm that was closer to *difl* than TLCΦ *attP* (Figure 5B, FtsK panel). In addition, those sites accounted for over 99.4% of the integration events (Figure 5B, FtsK panel). The same observations could be made on the n<sub>5a2n3g</sub> and tagn<sub>8</sub> butterfly plots (Supplementary Figure S5, FtsK panels). We attributed red, orange, cyan and blue colours to the XerD-arm motifs of the plasmids that integrated at *difl* at frequencies higher than 10<sup>-6</sup>, 10<sup>-7</sup>, 5 × 10<sup>-8</sup> or lower than 5 × 10<sup>-8</sup>, respectively. The red sites comprised 53 n<sub>8</sub>gtg, 61 n<sub>5a2n3g</sub> and 61 tagn<sub>8</sub> sequences (Supplementary Figure S5, FtsK panels). Together, those sites represented a total of 174 different sequences (Figure 5C, FtsK panel). The corresponding sequence logo was similar to *difl*, highlighting that FtsK-driven integration was

restricted to the *attP* sites that most resemble *difl* (Figure 5C, FtsK panel).

The butterfly representation further highlighted that the FtsK-driven integration frequency of a plasmid harbouring TLCΦ *attP* was over a 1000-fold lower than the mean integration frequency of the n<sub>8</sub>gtg and n<sub>5a2n3g</sub> red *attP* sites, and over a 100-fold lower than the mean integration frequency of the red tagn<sub>8</sub>*attP* sites (Figure 5 and Supplementary Figure S5, FtsK panels). In contrast, the efficiency of integration of TLCΦ *attP* jumped from about 10<sup>-9</sup> to about 10<sup>-6</sup> when it was driven by XafT (Figure 5 and Supplementary Figure S5, XafT and FtsK & XafT panels). Differences between the integration efficiency of the different *attP* sites were alleviated by the presence of XafT. In particular, the mean frequency of the red n<sub>8</sub>gtg *attP* sites was now only 30-fold higher than that of TLCΦ *attP* (Figure 5B, XafT and FtsK & XafT panels). Furthermore, the proportion of sites with an integration efficiency higher than 10<sup>-6</sup> dramatically increased. High integration efficiencies were not anymore limited to the (+) wing of the butterfly plots: XafT drove the integration of about 30% of the n<sub>8</sub>gtg mini-TLCΦ plasmids harbouring a site from the (−) bins (Figure 5B, XafT and FtsK & XafT panels). Results obtained with the n<sub>5a2n3g</sub> and tagn<sub>8</sub> plasmid libraries confirmed those observations (Supplementary Figure S5, XafT and FtsK & XafT panels).

As a result, XafT promoted the integration of over 6000 different *attP* sites with a frequency higher than 10<sup>-6</sup> in the recipient strain harbouring *lacZ-difl* at the *lacZ* locus and over 20000 different *attP* sites in the recipient strain harbouring *lacZ-difl* at the *difl* locus (Figure 5C, XafT and FtsK & XafT panels). The logo of those sites emphasized that efficient integration was no longer limited to sites that carried the same sequences as the XerD-arm of *difl* (Figure 5C and Supplementary Figure S5, XafT and FtsK & XafT panels).

### Mini-TLCΦ plasmids with *attP* sites deviating from *difl* escape FtsK-driven excision

An unusually long IMEX, the gonococcal genomic island (GGI), is integrated at the *dif* site of the chromosome of most pathogenic Neisseria species (39,40). It is flanked by a *dif*-like site of varying sequence between the species (Supplementary Figure S1B). The FtsK DNA translocase was shown to strip XerC and XerD bound to one of those sites. GGI1 *attP*, thereby preventing excision events (41). This is in contrast to the activity of FtsK at canonical *dif* sites, where the translocase stalls upon interaction with *dif*-bound XerD, prior to the activation of recombination. As GGI1 *attP* only deviates from the canonical *dif* sequence by 4 bp within the 6 outermost bp of the XerD arm, this suggests that sequence alterations in XerD-arm of IMEX *attP* sites could have evolved to prevent FtsK-driven excision events. To explore this hypothesis in the context of TLCΦ *attP*, we monitored the excision of XafT<sup>+</sup> n<sub>8</sub>gtg, tagn<sub>8</sub> and n<sub>5a2n3g</sub> mini-TLCΦ plasmids that had integrated at the *difl* locus. Inducing the expression of XerC and XerD did not reduce the total diversity of different *attP* sites in the cell population after 19h of growth in the absence of selective pressure (Figure 6A and Supplementary Figure S6, Xer<sup>+</sup>
<figure><img src="image_6.png" /><figcaption>**Figure 5.** Butterfly plots showing the relative integration efficiency of *attP* sites harbouring the different possible n<sub>8</sub>gtg XerD arm motifs. (A) Butterfly plot representation methodology. Δn: number of bases by which an XerD-arm motif deviates from TLCΦ *attP*. If the motif is more similar to *dif1* than TLCΦ *attP*, its integration frequency is plotted at the +Δn position of the X-axis. It is not closer to *dif1* than TLCΦ *attP* its integration frequency is plotted at the -Δn position of the X-axis. (B) Butterfly plots of the integrated n<sub>8</sub>gtg motifs. The proportion of integrated motifs falling in the (-) group is indicated. Their contribution to the global frequency of integration of the library is indicated between brackets. We coloured the various *attP* sites leading to significant integration driven by FtsK (FtsK panel) with respect to their integration frequency: from higher than 10<sup>-6</sup>, 10<sup>-7</sup>, 5 × 10<sup>-8</sup> or lower than 5 × 10<sup>-8</sup> in red, orange, cyan and blue, respectively. The corresponding sites in the XafT and FtsK & XafT panels are shown with the same colour code. Sites which were absent in the FtsK panel (motifs which were not capable of being integrated by FtsK) are shown in grey. The difference between the mean frequency of integration of red motifs and the integration frequency of TLCΦ *attP* (located in the middle of the graph) is indicated in red. The positions of TLCΦ *attP* and of the *attP* sites most similar to *dif1* are indicated by squares. (C) Combined sequence logos of the n<sub>8</sub>gtg, n<sub>5</sub>a<sub>2</sub>n<sub>3</sub>g and tagn<sub>8</sub> motifs with an integration frequency higher than 10<sup>-6</sup>. The logos show the frequency of each base at the degenerate positions. The number of motifs from which the logo was derived is indicated. Individual logos and Butterfly plots of the integrated n<sub>5</sub>a<sub>2</sub>n<sub>3</sub>g and tagn<sub>8</sub> motifs are shown in Supplementary Figure S5.</figcaption></figure>

butterfly plots). However, the proportion of the *attP* sites from the red and orange categories decreased (Figure 6A and Supplementary Figure S6, Xer⁺ bar plots). No excision was observed when the expression of XerC and XerD was not induced, demonstrating that excision depended on Xer recombination (Figure 6A and Supplementary Figure S6, Xer⁻ butterfly and bar plots). The efficiency of excision of the *attP* sites from the red and orange categories, i.e. those that most resembled *dif1*, was higher than the excision efficiency of TLCΦ *attP* (Figure 6A and Supplementary Figure S6, Xer⁺ butterfly plots). However, the efficiency of excision of all the sites, including those from the red and orange categories, was far lower than the 20% per generation excision frequency of a DNA cassette flanked by two *dif1* sites (28).

To determine the relative contribution of FtsK and XafT to the observed excision events, we analysed the excision of XafT⁺ n₈gtg plasmids integrated at the *lacZ* locus, where the action of FtsK is very limited. Even after 42 h of growth, there was little change in the relative frequency of the different *attP* sites (Figure 6B, Xer⁺ butterfly plots) and in the relative proportion of the sites from the five colour code categories (Figure 6B, Xer⁺ bar plots). These results suggested that under laboratory growth conditions, FtsK drove most if not all of the excision events of mini-TLCΦ plasmids integrated at the *dif* locus of chr1.

As *dif1* was used as the bacterial attachment site in the integration assays, excision was due to the reverse recombination reaction between the same initial sites (Figure 1). Therefore, we expected XafT to promote the excision of the plasmids for which it permitted integration. Two hypotheses could explain the inefficiency of XafT-driven excision: (i) FtsK could dismantle the formation of intramolecular synapses between *dif1* and non-canonical Xer recombination sites as proposed for the GGI1 (41); (ii) XafT expression from an integrated copy of TLCΦ could be insufficient for its activity. The inefficiency of XafT-driven excision events at the *lacZ* locus ruled out the first hypothesis since this locus is outside of the normal FtsK activity zone. To test the second hypothesis, we directly compared the expression of XafT in a strain carrying an integrated copy of TLCΦ and a strain carrying a replicative non-integrated copy of TLCΦ. We utilised 'pTLC' plasmids, which contain the entire genome of TLCΦ (except for its capsid-encoding genes) harboured on a conjugatable plasmid with a conditional R6K origin of replication and a chloramphenicol resistance marker. To observe expression by immunoblotting, we inserted a triple FLAG epitope (3xDYKDDDDDK) at the C-terminus of XafT (XafT-3FLAG), generating pTLC-3FLAG (Figure 6C). As controls, we constructed a non-integrative version of pTLC-3FLAG by precisely deleting its *attP* site (pTLC-3FLAG, Δ*attP*), and a non-replicative
<figure><img src="image_7.png" /><figcaption>**Figure 6.** Retention efficiency of the different possible n<sub>8</sub>gtg XafT<sup>+</sup> mini-TLCΦ libraries. (A) Retention efficiency from the natural *dif1* locus after growth in LB in conditions of production of XerCD (Xer<sup>+</sup>) or not (Xer<sup>-</sup>). Bar plots show the proportion of n<sub>8</sub>gtg motifs belonging to the red, orange, cyan, blue and grey categories within the integrated motifs. Retention efficiencies of the n<sub>5</sub>a<sub>2</sub>n<sub>3</sub>g and tagn<sub>8</sub> XafT<sup>+</sup> mini-TLCΦ libraries are shown in Supplementary Figure S6. (B) Excision from the *lacZ* locus after growth in LB, in conditions of production of XerCD (Xer<sup>+</sup>) or not (Xer<sup>-</sup>). (C) Anti-FLAG immunoblot of XafT-3xFLAG expression in integrated vs replicative pTLC conditions. The predicted molecular weight of XafT-3xFLAG C-terminal fusion protein (220 aa) is 25.5 kDa (EMBOSS Pepstats). Immunoblot contains normalized whole-cell samples of *V. cholerae* strain EPV369 over several plasmid conditions. Negative control is EPV369. Positive control is EPV369 containing an (induced) arabinose-inducible XafT-3xFLAG expression plasmid. Subsequent lanes contained cell extracts from either integrated or replicative versions of pTLC synthetic phage plasmids. White and blue illustrated colonies indicate samples derived from integrated (into the *lacZ-dif1* reporter cassette) or replicative *V. cholerae* clones, respectively. Cri<sup>-</sup> and Δ*attP* denote pTLC variants which are unable to replicate, or unable to integrate, respectively. Supplementary Figure S6C contains uncropped source images of the original SDS-PAGE, transfer, and immunoblot.</figcaption></figure>

version of pTLC-3FLAG by mutating its HUH endonuclease (pTLC-3FLAG, *cri*<sup>-</sup>). As the R6K origin of replication of the plasmid vector is non-functional in *V. cholerae*, replicative forms of pTLC are only driven by the rolling-circle replication activity of Cri. The three constructs were conjugated in the *V. cholerae* recipient strain harbouring the *lacZ-dif1* reporter at the *lacZ* locus. Conjugation of the non-integrative pTLC-3FLAG led to the formation of blue Cm-resistant colonies, indicating its efficient replication. Conjugation of the non-replicative pTLC-3FLAG led to the formation of white Cm-resistant colonies, indicating that it integrated at *lacZ-dif1*. Conjugation of the replica-

tive and integrative XafT-3FLAG pTLC led to the formation of both white and blue Cm-resistant colonies. We could detect XafT-3FLAG in protein extracts from the blue Cm-resistant cells harbouring either the TLCΦ plasmid or its integration-defective version. In contrast, we could not detect XafT-3FLAG in protein extracts from white Cm-resistant cells harbouring an integrated copy of TLCΦ, whether it could replicate or not, which was in agreement with the idea that XafT was not sufficiently expressed to drive Xer recombination after integration (Figure 6C).
<figure><img src="image_8.png" /><figcaption>**Figure 7.** Relative efficiency of XafT- and FtsKγ-driven Xer recombination reactions. (A) Synapse formation between *dif1* and TLCΦ *attP*. Scheme depicting that the non-canonical XerD-arm of TLCΦ *attP* should limit synapse formation and that FtsK translocation could dismantle and/or inhibit the formation of synapses, thereby preventing recombination. (B) Scheme of the *in-vitro* recombination substrates, the HJ recombination intermediate and the crossover products. Green ball: 3' Cy3 label; Red ball: 5' Cy5 label; Grey, Green and Blue rectangles: Central region, XerC-binding and XerD-binding arm, respectively. The DNA strands exchange by XerC and XerD are depicted as thin and fat lines, respectively. (C) XafT- and FtsKγ-promoted recombination reactions. Left panel: recombination of a short labelled *dif1* substrate (S1) and a long non-labelled *dif1* substrate (S2); Right panel: recombination of a short labelled *dif1* substrate (S1) and a long non-labelled TLCΦ *attP* substrate (S2). The S2 substrates are depicted above each gel images. +: *V. cholerae* XerC, *V. cholerae* XerD or MBP-XafT, as indicated; Dγ: *V. cholerae* XerD-FtsKγ fusion; -: mock buffer of the corresponding purified proteins. A white star indicates the presence of a faint HJ band.</figcaption></figure>

### TLCΦ *attP*/*dif1* synaptic complexes are rare and/or transient

FtsK only efficiently activates recombination between *dif* sites when it is bound on the XerD side of one or the other of the two DNA duplexes engaged in the recombination complex (42,43). It suggested the possibility that FtsK translocation could dismantle weakly assembled TLCΦ *attP*/*dif1* synaptic complexes before the γ-domain of FtsK could activate XerD, thereby preventing FtsK-driven integration events (Figure 7A, (41)). C-terminal fusions of FtsKγ to XerD have been previously shown to maximize the efficiency of reactions mediated by the *E. coli* and *N. gonorrhoea* Xer recombinases. Therefore, if the XerD-FtsKγ interaction was the limiting factor for Xer recombination of *dif1* vs TLCΦ *attP*, we expected to be able to promote the recombination of *dif1* with TLCΦ *attP* *in vitro* using a *V. cholerae* XerD-FtsKγ fusion protein (22,41). As a point of comparison, we reconstituted XafT-mediated reactions using XerC and XerD proteins and an N-terminal fusion of XafT to the maltose binding protein, MBP-XafT (31).

We used a short 34-bp synthetic double-stranded DNA (dsDNA) *dif1* fragment and longer dsDNA fragments containing either *dif1* or TLCΦ *attP* as substrates. To facilitate the differentiation of the HJ intermediate and the crossover products from the substrates, the 5' and 3' ends of the bottom strand of the short *dif1* substrate were labelled with cy5 and cy3 fluorescent dyes, respectively (Figure 7B). Incubation of the two *dif1* substrates with XerC and XerD-FtsKγ or with XerC, XerD and MBP-XafT yielded similar amounts of HJs and crossover products, suggesting that FtsKγ promoted recombination as efficiently as XafT when it was fused to XerD (Figure 7C, left panel). Yet, incubation of the short labelled *dif1* substrate and the TLCΦ *attP* substrate with XerC and XerD-FtsKγ produced barely detectable amounts of HJs and crossover products, suggesting that FtsKγ does not greatly stimulate the reaction between *attP* and *dif*, and that this poor activation is not a direct result of the FtsK translocase activity (Figure 5C, right panel). In contrast, XerC, XerD and MBP-XafT mediated the formation of similarly high amounts of HJs and crossover products between the short labelled *dif1* substrate and the long *dif1* or TLCΦ *attP* substrates (Figure 7B). We conclude that TLCΦ *attP*/*dif1* synaptic complexes are normally too rare and/or too transient to allow for FtsKγ-driven activation of XerD.

## DISCUSSION

### An NGS-based methodology to analyse the sequence requirements of a DNA recombination reaction

Studies on the sequence specificity of DNA transactions are generally limited to the consensus sequence targeted by DNA binding proteins. When there are multiple copies of a binding motif in the genome, it can be achieved with Chromatin Immuno-Precipitation (ChIP) or Whole-Genome Affinity Determination of protein-DNA interactions (WGADseq) methodologies (44). When the binding motif has low sequence variability, additional investigations may include electromobility gel assays, DNA footprinting, and/or systematic evolution of ligands by exponential enrichment (SELEX). The resulting consensus or sequence logo can be refined by replacing each base of the target motif one at time and monitoring how it affects the binding of the protein. However, sequence logos can mask the ability of proteins to bind to several distinct consensus sequences. In addition, they do not permit the determination of the relative influence of each particular position of a sequence on the base requirements at the other positions. To bypass those limitations, we designed a methodology based on NGS to monitor the integration and excision efficiency of thousands of different mobile elements *in vivo* (Figure 2). We also developed two new graphical representations that facilitate the analysis of the results, 2D-maps (Figure 4) and butterfly plots (Figure 5). We applied the methodology to explore the influence of the XerD-arm of TLCΦ *attP* on the efficiency of Xer recombination reactions (Figures 1 and 2). The extreme sensitivity of the methodology permitted the evaluation of the integration efficiency of events that were too rare to be detected by the classical methodology, such as the integration of TLCΦ *attP* by a FtsK-driven Xer
recombination reaction (Figures 3–5). 2D-map and butterfly plot graphical representations highlighted that efficient FtsK-driven Xer recombination reactions were limited to the *attP* sites whose XerD-arm only very slightly deviated from the consensus XerD-arm of *dif* sites (Figures 3–6). In contrast, they revealed that XafT promoted recombination reactions between *difl* and almost all of the different possible *attP* sites provided that the first or fifth innermost bases of their XerD-arm was identical to the corresponding *dif* bases (Figures 3–5). Our methodology and graphical representations could be easily adapted to explore the sequence specificity of any DNA binding protein using the relative retention of DNA motifs by the protein *in vitro*, or using the relative activity of an artificial promoter carrying the DNA motifs *in vivo*.

### XafT allows Xer recombination between *dif* and *attP* sites with non-canonical XerD-arms

Xer recombination takes place in a nucleoprotein complex consisting of two recombinating sites and a pair of each of the two XerC and XerD recombinases (Figure 1). The complex is held together by the affinity of the recombinases to their DNA binding sites and cyclic interactions between their C-terminal domains. However, binding of XerC to *dif* is relatively weaker than binding of XerD (2,20,45). Those observations suggested that contacts between XerD and *dif* played a primary role in the assembly and stability of the recombination synapses (2,20,45). In this regard, the efficiency with which TLCΦ was integrated into the primary chromosome of clinical and environmental clones of *V. cholerae* was surprising since the XerD-arm of TLCΦ *attP* was a poor substrate for XerD-binding whether its XerC protein partner was present or not *in vitro* (9,13). TLCΦ relies on its own XerD-activation factor for integration, XafT (9,31). No other *V. cholerae* or TLCΦ protein or sequence factors (in combination with XerCD) are required to promote a complete Xer recombination reaction between TLCΦ *attP* and *difl* *in vitro* (31).

Our analysis of the XerD-arm of TLCΦ *attP* on the integration efficiency and stability of mini-TLCΦ plasmids showed that FtsK only promotes the integration into *difl* of a very small subset of the different 190 528 TLCΦ *attP* derived sites we studied, restricted mostly to those that were the most similar to *difl* (Figures 3–5). In contrast, XafT promoted the recombination of *difl* with almost any of the 190 528 *attP* sites that carried the same residues as *difl* at the first or fifth position of the XerD-arm (Figures 3–5). In addition, it alleviated differences in the integration frequency of *attP* sites harbouring more or less canonical XerD-arms (Figure 5). The 2D map and butterfly plot representations of the integration of XafT⁺ mini-TLCΦ plasmid libraries in the *V. cholerae* recipient strain harbouring *lacZ-difl* at the native *difl* locus were similar to those obtained for the integration of XafT⁺ mini-TLCΦ plasmid libraries in the *V. cholerae* recipient strain harbouring *lacZ-difl* at the *lacZ* locus indicating that FtsK did not prevent XafT-driven integration of XafT⁺ mini-TLCΦ plasmids (Figures 4 and 5). The slightly higher integration efficiencies observed for the reporter strain carrying *lacZ-difl* at the native *difl* locus is probably explained by the growth advantage of this

strain over the reporter strain carrying *difl* at the *lacZ* locus (Figure 3). Finally, *in vitro* comparison of the efficiency with which XafT and FtsKγ drove recombination reactions between two *difl* sites or between *difl* and TLCΦ *attP* showed that TLCΦ *attP/difl* synaptic complexes are normally too rare and/or too transient to allow for FtsKγ-driven activation of XerD (Figure 7). Taken together, those results suggest that XafT can promote the efficient recombination of *difl* with TLCΦ *attP*-derived sites harbouring a non-canonical XerD-arm because XafT helps assemble and/or stabilize Xer synaptic complexes. As XafT contains a dimerization domain and directly interacts with XerD, it is tempting to propose that it recruits a second XerD recombinase at the *difl* locus by forming a proteinaceous bridge with an XerD recombinase bound to *difl* (31). The resulting *difl/XerC/XerD/XafT/XerD* complex could then be engaged in a larger nucleoprotein complex with an *attP* site solely bound by XerC via the interactions of recombinases. Thus, a limited amount of homology between the XerD-arm of the *attP* site and the canonical XerD binding site would be required to ensure the formation of a productive recombination synapse. Future work will need to confirm this hypothesis by directly analysing synaptic complex assembly with or without XafT using biophysical methods.

This scenario is reminiscent of the control of Xer recombination on multicopy plasmids. The dimer resolution sites of multicopy plasmids exploiting Xer and the attachment sites of IMEX relying on the non-canonical XerC-first recombination pathway contain a few degenerate positions in the XerC and XerD arms (Supplementary Figure S1B). In particular, there are two mutations in the inner region of the XerC-arm of the ColE1 dimer resolution site, *cer*, including one at the 5th position, and there is a mutation at the sixth position of the XerD-arm, which together would be expected to limit the formation of and/or the stability of synaptic complexes (Supplementary Figure S1B). The efficiency and directionality of Xer recombination then relies on accessory proteins that bind to accessory sequences flanking the core *cer* site, which bring together the core *cer* sites in a synaptic complex with a specific topological configuration (30,46). Future studies will also need to address the evolutionary pressures that seems to have set the non-canonical bases of each plasmid dimer resolution and IMEX attachment sites (Supplementary Figure S1B). The NGS methodology we present will be a great asset for future work on this subject.

### Importance of the fifth innermost bp of the XerD-arm for recombination

The NGS data allowed us to revisit the importance of the different bases within the XerD arm of *dif* sites. Whether integration was driven by FtsK or XafT, the mini-TLCΦ plasmids that integrated the most efficiently were those that harboured the most *dif*-like motifs (Figure 5). It further indicated that the five outermost positions of the XerD-arm were less constrained than the six innermost positions (Figure 5). This result could seem surprising since previous work showed that the specificity of binding of the *E. coli* Xer recombinases was ensured by the base composition of the sixth, seventh and ninth to eleventh non-palindromic posi-
tions of their binding arms (47,48). However, contacts be-
tween the *E. coli* recombinases and *dif* were shown to be
mainly limited to the seven innermost bases of their bind-
ing sites, and in particular to the first five palindromic bases
(Supplementary Figure S1, (45)). Correspondingly, the in-
nermost bases of the XerC and XerD binding sites are more
conserved than the outermost bases through evolution (49).
Interestingly, it was recently reported that *Bacillus subtilis*
and *Staphylococcus aureus* XerD serves to unload bacte-
rial SMC complexes (50). This activity relies on the bind-
ing of XerD to additional chromosomal loci other than *dif*,
independently from XerC. Those loci harbour a *dif*-like site
composed of a bona fide XerD binding site, a 5-6 bp degen-
erate central region and the five innermost bp of the XerD
binding site (50). Our results raise the possibility that those
sites evolved to allow the cooperative binding of two XerD
molecules.

The nature of the fifth bp of the XerD-arm of the *attP*
sites seems the most important for XafT-driven recombi na-
tion (Figures 3–5). Correspondingly, it was previously ob-
served in *E. coli* that a *dif* site carrying a single mutation at
the fifth bp of the XerD-arm was less proficient than wild-
type *dif* for Xer recombination (45). In addition, methyla-
tion interference analysis showed that *E. coli* XerD inter-
acted with the guanine at the fifth position of the top strand
of its binding site (45). A model based on the crystal struc-
ture of *E. coli* XerD and the Catabolite Activator Protein-
DNA complex further suggested that XerD contacts the
guanine of the top strand of its binding site with a con-
served arginine of its C-terminal domain (R220 in *E. coli*
XerD and R224 in *V. cholerae* XerD, (51)). A cytosine base
is present at the 5th innermost position of top strand of the
*dif* site of *Helicobacter pylori* and *Lactococcus lactis*, which
harbour alternative Xer machineries composed of a single
Xer recombinase (Supplementary Figure S1). This position
was shown to be important for the binding and the activity
of *H. pylori* Xer recombinase (52). In effect, this basepair is
inverted with respect to the GC bp of canonical XerC and
XerD-arms (Supplementary Figure S1B). X-ray structure
analysis revealed that the *H. pylori* recombinase contacts
the bottom-strand guanine of this bp with an arginine of
its N-terminal domain, R65 (52). These observations high-
light the evolutionary distance between conventional and
alternative Xer machineries.

Preventing undesired Xer-mediated excision

IMEX such as CTXΦ and VGJΦ exploit the low basal
ability of XerC to catalyse the formation of HJs, which
are subsequently resolved by DNA replication (Figure 1C,
(7,8,29)). This strategy of integration is advantageous be-
cause it escapes the requirement for FtsK and because the
reverse reaction necessarily leaves an intact chromosomal
copy of the phage (8). The stability of CTXΦ is further en-
sured because its *attP* site is formed by a hairpin of its sin-
gle stranded DNA genome, which is masked in the double
stranded integrated prophage (Supplementary Figure S1,
(7,29)). In contrast, TLCΦ and the GGIs rely on an intrin-
sically fully reversible reaction (Figure 1D, (9,31)). It was
proposed that FtsK translocation could prevent excision of
the GGIs because it strips XerD from non-canonical XerD-

binding sites (41). Our results lend further support to this
hypothesis since all of the 190 528 mini-TLCΦ plasmids we
studied, including those that harboured the *attP* sites that
most resembled *dif1*, were far more stable than a DNA cas-
sette flanked by two *dif1* sites (Figure 6 and Supplementary
Figure S6, (28,37)). However, this hypothesis does not ex-
plain the excision inefficiency of mini-TLCΦ plasmids that
are integrated at the *lacZ* locus, which is outside the FtsK
activity zone (Figure 6, (28,36–38)). It suggested that in-
tegrated mini-TLCΦ plasmids were stable because the ex-
pression of XafT was insufficient after integration. In sup-
port of this hypothesis, we could detect a C-terminal tagged
3XFLAG tag fusion of XafT, which is active in recombi na-
tion, in cells harbouring a non-integrated replicative copy
of TLCΦ but did not detect it in cells harbouring an in-
tegrated copy of TLCΦ (Figure 6). Future work will need
to investigate the regulation of XafT expression and under
which conditions it can be increased to promote the excision
of integrated copies of TLCΦ.

DATA AVAILABILITY

NGS plasmid library verification, integration and excision
data have been deposited in the ArrayExpress database at
EMBL-EBI (www.ebi.ac.uk/arrayexpress) under accession
number E-MTAB-10951.

SUPPLEMENTARY DATA

Supplementary Data are available at NAR Online.

ACKNOWLEDGEMENTS

We thank Raphaël Guérois and Virginia Lioy for helpful
discussions, and Mhairi Davidson for critical reading.

FUNDING

European Research Council [FP7/2007-2013/28159 to F.-X.B.]; Agence Nationale pour la Recherche [2016-CE12-0030-0 and 2021-CE35-0013-01 to F.-X.B., F.O., 2018-CE12-0012-03 to F.-X.B.]; Fondation pour la Recherche Médicale [EQU202003010328 to F.-X.B.]. Funding for open access charge: Fondation pour la Recherche Médicale [EQU202003010328 to F.-X.B.].

*Conflict of interest statement.* None declared.

REFERENCES

1. Midonet, C. and Barre, F.-X. (2014) Xer site-specific recombination: promoting vertical and horizontal transmission of genetic information. *Microbiol. Spectr.*, **2**.

2. Blakely, G., May, G., McCulloch, R., Arciszewska, L. K., Burke, M., Lovett, S. T. and Sherratt, D. J. (1993) Two related recombinases are required for site-specific recombination at *dif* and *cer* in *E. coli* K12. *Cell*, **75**, 351–361.

3. Colloms, S.D., Sykora, P., Szatmari, G. and Sherratt, D.J. (1990) Recombination at *cole1* cer requires the *Escherichia coli* xerC gene product, a member of the lambda integrase family of site-specific recombinases. *J. Bacteriol.*, **172**, 6973–6980.

4. Das, B., Martínez, E., Midonet, C. and Barre, F.-X. (2013) Integrative mobile elements exploiting xer recombination. *Trends Microbiol.*, **21**, 23–30.
5. Balalovski,P. and Grainge,I. (2020) Mobilization of pdfif modules in acinetobacter: a novel mechanism for antibiotic resistance gene shuffling? *Mol. Microbiol.*, 10, 1111/mmi.14563.

6. Huber,K.E. and Waldor,M.K. (2002) Filamentous phage integration requires the host recombinases XerC and xerD. *Nature*, **417**, 656–659.

7. Das,B., Bischerour,J., Val,M.-E. and Barre,F.-X. (2010) Molecular keys of the tropism of integration of the cholera toxin phage. *Proc. Natl. Acad. Sci. U.S.A.*, **107**, 4377–4382.

8. Das,B., Bischerour,J. and Barre,F.-X. (2011) VGJphi integration and excision mechanisms contribute to the genetic diversity of vibrio cholerae epidemic strains. *Proc. Natl. Acad. Sci. U.S.A.*, **108**, 2516–2521.

9. Midonet,C., Das,B., Paly,E. and Barre,F.-X. (2014) XerD-mediated ftsk-independent integration of TLCφ into the vibrio cholerae genome. *Proc. Natl. Acad. Sci. U.S.A.*, **111**, 16848–16853.

10. Ramamurthy,T., Mutreja,A., Weill,F.-X., Das,B., Ghosh,A. and Nair,G.B. (2019) Revisiting the global epidemiology of cholera in conjunction with the genomics of vibrio cholerae. *Front. Public Health*, **7**

11. Waldor,M.K. and Mekalanos,J.J. (1996) Lysogenic conversion by a filamentous phage encoding cholera toxin. *Science*, **272**, 1910–1914.

12. Rubin,E.J., Lin,W., Mekalanos,J.J. and Waldor,M.K. (1998) Replication and integration of a vibrio cholerae cryptic plasmid linked to the CTX prophage. *Mol. Microbiol.*, **28**, 1247–1254.

13. Hassan,F., Kamruzzaman,M., Mekalanos,J.J. and Faruque,S.M. (2010) Satellite phage TLCφ enables toxigenic conversion by CTX phage through dif site alteration. *Nature*, **467**, 982–985.

14. Chun,J., Grim,C.J., Hasan,N.A., Lee,J.H., Choi,S.Y., Haley,B.J., Taviani,E., Jeon,Y.-S., Kim,D.W., Lee,J.-H. et al. (2009) Comparative genomics reveals mechanism for short-term and long-term clonal transitions in pandemic vibrio cholerae. *Proc. Natl. Acad. Sci. U.S.A.*, **106**, 15442–15447.

15. Mutreja,A., Kim,D.W., Thomson,N.R., Connor,T.R., Lee,J.H., Kariuki,S., Croucher,N.J., Choi,S.Y., Harris,S.R., Lebens,M. et al. (2011) Evidence for several waves of global transmission in the seventh cholera pandemic. *Nature*, **477**, 462–465.

16. Heidelberg,J.F., Eisen,J.A., Nelson,W.C., Clayton,R.A., Gwinn,M.L., Dodson,R.J., Haft,D.H., Hickey,E.K., Peterson,J.D., Umayam,L. et al. (2000) DNA sequence of both chromosomes of the cholera pathogen vibrio cholerae. *Nature*, **406**, 477–483.

17. Weill,F.-X., Domman,D., Njamkepo,E., Tarr,C., Rauzier,J., Fawal,N., Keddy,K.H., Salje,H., Moore,S., Mukhopadhyay,A.K. et al. (2017) Genomic history of the seventh pandemic of cholera in Africa. *Science*, **358**, 785–789.

18. Domman,D., Quilici,M.-L., Dorman,M.J., Njamkepo,E., Mutreja,A., Mather,A.E., Delgado,G., Morales-Espinosa,R., Grimont,P.A.D., Lizárraga-Partida,M.L. et al. (2017) Integrated view of vibrio cholerae in the Americas. *Science*, **358**, 789–793.

19. Aussel,L., Barre,F.X., Aroyo,M., Stasiak,A., Shasiak,A.Z. and Sherratt,D. (2002) FtsK is a DNA motor protein that activates chromosome dimer resolution by switching the catalytic state of the XerC and XerD recombinases. *Cell*, **108**, 195–205.

20. Val,M.-E., Kennedy,S.P., El Karoui,M., Bonné,L., Chevalier,F. and Barre,F.-X. (2008) FtsK-dependent dimer resolution on multiple chromosomes in the pathogen vibrio cholerae. *PLoS Genet.*, **4**, e1000201.

21. Yates,J., Aroyo,M., Sherratt,D.J. and Barre,F.-X. (2003) Species specificity in the activation of xer recombination at dif by ftsK. *Mol. Microbiol.*, **49**, 241–249.

22. Grainge,I., Lesterlin,C. and Sherratt,D.J. (2011) Activation of xercd-dif recombination by the FtsK DNA translocase. *Nucleic Acids Res.*, **39**, 5140–5148.

23. Yates,J., Zhekov,I., Baker,R., Eklund,B., Sherratt,D.J. and Arciszewska,L.K. (2006) Dissection of a functional interaction between the DNA translocase, FtsK, and the XerD recombinase. *Mol. Microbiol.*, **59**, 1754–1766.

24. Bigot,S., Saleh,O.A., Lesterlin,C., Pages,C., El Karoui,M., Dennis,C., Grigoriev,M., Allemand,J.-F., Barre,F.-X. and Cornet,F. (2005) KOPS: DNA motifs that control e. coli chromosome segregation by orienting the FtsK translocase. *J. Biol.*, **24**, 3770–3780.

25. Kennedy,S.P., Chevalier,F. and Barre,F.-X. (2008) Delayed activation of xer recombination at dif by FtsK during septum assembly in escherichia coli. *Mol. Microbiol.*, **68**, 1018–1028.

26. Barre,F.X., Aroyo,M., Colloms,S.D., Helfrich,A., Cornet,F. and Sherratt,D.J. (2000) FtsK functions in the processing of a holliday junction intermediate during bacterial chromosome segregation. *Genes Dev.*, **14**, 2976–2988.

27. Cornet,F., Louarn,J., Patte,J. and Louarn,J.M. (1996) Restriction of the activity of the recombination site dif to a small zone of the *Escherichia coli* chromosome. *Genes Dev.*, **10**, 1152–1161.

28. Galli,E., Midonet,C., Paly,E. and Barre,F.-X. (2017) Fast growth conditions uncouple the final stages of chromosome segregation and cell division in escherichia coli. *PLoS Genet.*, **13**, e1006702.

29. Val,M.-E., Bouvier,M., Campos,J., Sherratt,D., Cornet,F., Mazel,D. and Barre,F.-X. (2005) The single-stranded genome of phage CTX is the form used for integration into the genome of vibrio cholerae. *Mol. Cell*, **19**, 559–566.

30. Colloms,S.D., Bath,J. and Sherratt,D.J. (1997) Topological selectivity in xer site-specific recombination. *Cell*, **88**, 855–864.

31. Midonet,C., Miele,S., Paly,E., Guerois,R. and Barre,F.-X. (2019) The tlcφ satellite phage harbors a xer recombination activation factor. *Proc. Natl. Acad. Sci. U.S.A.*, **116**, 18391–18396.

32. Meibom,K.L., Blokesch,M., Dolganov,N.A., Wu,C.Y. and Schoolnik,G.K. (2005) Chitin induces natural competence in vibrio cholerae. *Science*, **310**, 1824–1827.

33. Engler,C., Kandzia,R. and Marillonnet,S. (2008) A one pot, one step, precision cloning method with high throughput capability. *PLoS One*, **3**, e3647.

34. Gibson,D.G., Young,L., Chuang,R.-Y., Venter,J.C., Hutchison,C.A. and Smith,H.O. (2009) Enzymatic assembly of DNA molecules up to several hundred kilobases. *Nat. Methods*, **6**, 343–345.

35. Demarre,G., Guérout,A.-M., Matsumoto-Mashimo,C., Rowe-Magnus,D.A., Marlière,P. and Mazel,D. (2005) A new family of mobilizable suicide plasmids based on broad host range R388 plasmid (IncW) and RP4 plasmid (IncPα) conjugative machineries and their cognate escherichia coli host strains. *Res. Microbiol.*, **156**, 245–255.

36. Espinosa,E., Paly,E. and Barre,F.-X. (2020) High-resolution whole-genome analysis of sister-chromatid contacts. *Mol. Cell*, **79**, 857–869.e3.

37. Demarre,G., Galli,E., Muresan,L., Paly,E., David,A., Possoz,C. and Barre,F.-X. (2014) Differential management of the replication terminus regions of the two vibrio cholerae chromosomes during cell division. *PLoS Genet.*, **10**, e1004557.

38. David,A., Demarre,G., Muresan,L., Paly,E., Barre,F.-X. and Possoz,C. (2014) The two Cis-acting sites, parS1 and oriC1, contribute to the longitudinal organisation of vibrio cholerae chromosome I. *PLoS Genet.*, **10**, e1004448.

39. Dillard,J.P. and Seifert,H.S. (2001) A variable genetic island specific for neisseria gonorrhoeae is involved in providing DNA for natural transformation and is found more often in disseminated infection isolates. *Mol. Microbiol.*, **41**, 263–277.

40. Domínguez,N.M., Hackett,K.T. and Dillard,J.P. (2011) XerCD-mediated site-specific recombination leads to loss of the 57-kilobase gonococcal genetic island. *J. Bacteriol.*, **193**, 377–388.

41. Fournes,F., Crozat,E., Pages,C., Tardin,C., Salomé,L., Cornet,F. and Rousseau,P. (2016) FtsK translocation permits discrimination between an endogenous and an imported Xer/dif recombination complex. *Proc. Natl. Acad. Sci. U.S.A.*, **113**, 7882–7887.

42. Massey,T.H., Aussel,L., Barre,F.-X. and Sherratt,D.J. (2004) Asymmetric activation of xer site-specific recombination by ftsK. *EMBO Rep.*, **5**, 399–404.

43. Bonné,L., Bigot,S., Chevalier,F., Allemand,J.-F. and Barre,F.-X. (2009) Asymmetric DNA requirements in xer recombination activation by ftsK. *Nucleic Acids Res.*, **37**, 2371–2380.

44. Poidevin,M., Galli,E., Yamaichi,Y. and Barre,F.-X. (2017) WGADseq: whole genome affinity determination of protein-dna binding sites. In: *The Bacterial Nucleoid: Methods and Protocols, Methods in Molecular Biology*. Humana Press, New York, NY.

45. Blakely,G.W. and Sherratt,D.J. (1994) Interactions of the site-specific recombinases XerC and XerD with the recombination site dif. *Nucleic Acids Res.*, **22**, 5613–5620.

46. Bregu,M., Sherratt,D.J. and Colloms,S.D. (2002) Accessory factors determine the order of strand exchange in xer recombination at psi. *EMBO J.*, **21**, 3888–3897.

47. Blakely,G. and Sherratt,D. (1996) Determinants of selectivity in xer site-specific recombination. *Genes Dev.*, **10**, 762–773.
48. Hayes, F. and Sherratt, D.J. (1997) Recombinase binding specificity at the chromosome dimer resolution site dif of escherichia coli. *J. Mol. Biol.*, **266**, 525–537.

49. Kono,N., Arakawa,K. and Tomita,M. (2011) Comprehensive prediction of chromosome dimer resolution sites in bacterial genomes. *BMC Genomics*, **12**, 19.

50. Karaboja,X., Ren,Z., Brandão,H.B., Paul,P., Rudner,D.Z. and Wang,X. (2021) XerD unloads bacterial SMC complexes at the replication terminus. *Mol. Cell*, **81**, 756–766.e8.

51. Subramanya,H.S., Arciszewska,L.K., Baker,R.A., Bird,L.E., Sherratt,D.J. and Wigley,D.B. (1997) Crystal structure of the site-specific recombinase, xerD. *EMBO J.*, **16**, 5178–5187.

52. Bebel,A., Karaca,E., Kumar,B., Stark,W.M. and Barabas,O. (2016) Structural snapshots of xer recombination reveal activation by synaptic complex remodeling and DNA bending. *Elife*, **5**.