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Published in final edited form as: Nat Ecol Evol. 2017 Jul 31;1(9):1348–1353. doi: 10.1038/s41559-017-0250-3

Positive selection inhibits gene mobilisation and transfer in soil bacterial communities

James P J Hall 1,2,*, David Williams 3, Steve Paterson 3, Ellie Harrison 1, Michael A Brockhurst 1,*
PMCID: PMC5584672  EMSID: EMS73212  PMID: 28890938

Horizontal gene transfer (HGT) between bacterial lineages is a fundamental evolutionary process that accelerates adaptation. Sequence analyses show conjugative plasmids are principal agents of HGT in natural communities. However, we lack understanding of how the ecology of bacterial communities and their environments affect the dynamics of plasmid-mediated gene mobilisation and transfer. Here we show, in simple experimental soil bacterial communities containing a conjugative mercury resistance plasmid, the repeated, independent mobilisation of transposon-borne genes from chromosome to plasmid, plasmid to chromosome, and, in the absence of mercury selection, interspecific gene transfers from the chromosome of one species to the other via the plasmid. By reducing conjugation, positive selection for plasmid-encoded traits, like mercury resistance, can consequently inhibit HGT. Our results suggest that interspecific plasmid-mediated gene mobilisation is most likely to occur in environments where plasmids are infectious, parasitic elements rather than those where plasmids are positively selected, beneficial elements.

Conjugative plasmids — semi-autonomous mobile genetic elements that transfer between bacteria — are key agents of horizontal gene transfer (HGT) 1,2, facilitating rapid bacterial adaptation by spreading ecologically important traits between lineages 3. The physical movement or duplication of genes (gene mobilisation) between chromosomes and plasmids, and their subsequent transfer between hosts, can be decisive in microbial evolution, and has facilitated acquisition of antimicrobial resistance 4,5 and emergence of virulent pathogens 6,7. Such mobilisation can be facilitated by transposable elements (TEs), which encode enzymes (transposases) allowing transfer of genes between replicons 2. However, the effects of ecological factors and natural selection on gene mobilisation and subsequent HGT is unclear. This is particularly the case for natural environments with a high degree of spatial structure 8, which is rarely captured by experimental studies. An outstanding question is how positive selection for plasmid-encoded traits, like resistance genes, affects the ability of that plasmid to spread genes through a community. Although positive selection can favour HGT by benefitting bacteria which have acquired the plasmid (transconjugants), it can also prevent HGT by killing or inhibiting growth of potential plasmid recipients 9,10.

Experimental evolution of bacterial communities is a powerful tool for exploring the evolutionary dynamics of plasmid-mediated HGT, bridging the gap between simplified short-term laboratory studies 11,12 and comparative genomics of natural populations 4,5,13,14. To investigate how positive selection for plasmid-borne genes and the presence of an alternative host species interact to determine plasmid dynamics, we established communities of the common soil bacteria Pseudomonas fluorescens and Pseudomonas putida, either alone or in co-culture, in sterilized soil microcosms, which offer a complex, spatially-structured, and experimentally tractable environment 15–17. Communities began with the 307-kb conjugative plasmid pQBR57 at ~50% frequency; this plasmid was isolated from agricultural soil and carries a 7-kb mercury resistance (HgR) TE, Tn5042. We also established control communities without pQBR57. Microcosms were supplemented with mercuric chloride to 16 µg/g Hg(II)), or an equivalent volume of water (0 µg/g Hg(II)). This level of mercury contamination, similar to that found in industrial or post-industrial sites 18, selects for specific HgR but does not necessitate it 16. Six replicate populations for each combination of treatments were grown for ~440 generations. Analysis of plasmid frequency dynamics showed that pQBR57 was generally maintained by P. fluorescens and lost by P. putida, but persisted in P. putida when co-cultured with P. fluorescens, due to interspecific ‘source-sink’ plasmid transfer 17. To investigate consequent effects on gene mobilisation and transfer we sequenced clones from the beginning and the end of the experiment, and used the Bacterial and Archaeal Genome Analyser (BAGA) pipeline to identify structural variations 19.

Results

We detected multiple, independent gene mobilisation events between plasmid and chromosome in each species. Strikingly and unexpectedly, we also identified numerous interspecific transfers of chromosomal genes in the co-cultured treatments, facilitated by pQBR57 (Fig. 1). Gene transfer from P. fluorescens to P. putida was exemplified by a previously-unannotated P. fluorescens TE, Tn6291, in P. putida plasmids from 2/6 co-cultured communities (Fig. 1, replicates b and f). Subsequent PCR analyses found Tn6291 in P. putida clones from 2 further co-cultured communities (replicates d and e). Tn6291, a 22 kb TE carrying 25 predicted open reading frames and located between 2060105 and 2082440 in the P. fluorescens SBW25 reference sequence (ENA identifier AM181176, part of genomic island SBW_GI-1 (ref. 20)), carries an array of cargo genes with putative cytochrome c/d oxidase functions. The presence of Tn6291 in P. putida indicates these genes mobilised from the P. fluorescens chromosome onto pQBR57, and subsequently transferred between species via conjugation. All Tn6291+ P. putida clones were also positive for pQBR57, suggesting Tn6291 remains plasmid-borne in these isolates. Indeed, we detected plasmid-borne Tn6291 in 4/6 0 µg/g Hg(II) single-species P. fluorescens populations, and BLAST analysis shows a similar transposon in another soil Pseudomonad, P. syringae pv. syringae B301D (ref. 21) , suggesting Tn6291 mobilises readily.

Figure 1.

Figure 1

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Evolved clones show extensive within- and between-species gene mobilisation. Each panel shows events detected in evolved P. putida (left, light green) and P. fluorescens (right, light blue), with changes in associated pQBR57 (if present) shown below. In each panel, six concentric lanes a–f indicate independent populations. One clone was sequenced from each population for each species, except where both plasmid-bearing and plasmid-free genotypes were detected. In this case, we sequenced one clone of each, with the plasmid-bearing clone indicated by the inner set of symbols in that lane (Supplementary Table 1). Duplicative insertions of large TEs are indicated by filled triangles coloured according to TE type (see key) and connected to ancestral positions (indicated by open triangles) by an arrow describing direction of duplication. Dotted lines indicate insertions that occurred before the evolution experiment was initiated (see Methods). Insertions of smaller insertion sequence (IS) elements are in black. Black bars indicate large deletions, and yellow bars (panel C, replicate e) indicate large tandem duplications. Scale is given in Mbp, and replicons are scaled to the same size for clarity. (A) Clones evolved in single-species populations with 0 µg/g Hg(II). (B) Clones evolved in single-species population with 16 µg/g Hg(II). In panels A and B, lines indicate the physical separation of the two species. (C) Clones evolved in co-cultured populations with 0 µg/g Hg(II). (D) Clones evolved in co-cultured populations with 16 µg/g Hg(II).

We also detected gene transfer from P. putida to P. fluorescens. P. fluorescens clones from 3/6 co-cultured populations had acquired the well-described P. putida TE Tn4652 (ref. 22) in their chromosomes, with plasmid-borne Tn4652 also present. Tn4652 is a 17 kb TE closely related to the Tn4651 toluene degradation transposon and encodes various putative enzymes including a diacylglycerol kinase and a sulfatase. Tn4652 mobilisation to the plasmid occurred readily, with events already detectible in the ancestral P. putida clones used to inoculate the soil and begin the experiment (see Materials and Methods). PCR analysis of clones obtained over the course of the evolution experiment detected Tn4652+ plasmids in P. fluorescens as early as transfer 3, however Tn4652 insertion in the P. fluorescens chromosome was only detected later, after transfer 41 (Supplementary Table 2). In all cases, Tn4652 inserted in a region of the P. fluorescens chromosome with atypical sequence composition, likely to be recently acquired DNA 20.

Importantly, interspecific transfer of chromosomal TEs via the plasmid was only detected in populations grown without positive selection for the plasmid (0 µg/g Hg(II)). The amount of plasmid conjugation occurring, and thus opportunities for interspecific gene transfer, is likely a function of the densities of plasmid bearers and recipients 23,24. By killing potential plasmid recipients, mercury selection reduces encounters between plasmid donors and recipients, and therefore conjugation 9. Indeed, short-term experiments examining pQBR57 transfer (Figure 2) showed reduced effects of conjugation on plasmid dynamics when the plasmid was under selection 10, implying limited gene exchange. Together, these data suggest that positive selection for plasmid-borne resistance genes reduced the ability of that plasmid to facilitate HGT of chromosomal genes.

Figure 2.

Figure 2

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Plasmid dynamics are altered under positive selection. Top row: plasmid-bearing (‘donor’) and plasmid-free (‘recipient’) P. fluorescens were mixed in approximately equal ratios and cultured for 5 transfers in 0 µg/g Hg(II) microcosms. Densities of donors (dotted line) and recipients (solid line) and their plasmid statuses (donor, yellow fill; recipient, green fill; filled areas are overlaid) were estimated each transfer by plating onto selective media and replica plating onto Hg(II) where appropriate. Each panel represents an independent population. Bottom row: as top row except with 16 µg/g Hg(II). Conjugation makes a reduced contribution to plasmid dynamics under 16 µg/g Hg(II) (Z = 2.88, p = 0.002, n = 12, Exact General Independence Test). These results are similar to those reported by Stevenson et al.10, showing that this pattern holds in soil microcosms.

While mercury selection reduced TE transfer between species, we detected frequent mobilisation of the HgR TE Tn5042 from pQBR57 to the chromosome. Single-species P. putida populations tended to lose the plasmid 17, and sequences show that under mercury selection this was facilitated by acquisition of chromosomal Tn5042. To track the acquisition of chromosomal Tn5042 by P. putida populations, we designed PCR primers targeting ‘focal’ Tn5042 insertions (i.e. insertions detected in the end-point genome sequences) in eight different P. putida populations, and applied these to clones collected across the experiment. As with single-species P. putida populations, chromosomal Tn5042 was readily acquired by co-cultured P. putida, although these populations tended to also maintain pQBR57 (Figure 3). Similarly, chromosomal Tn5042 was detected in the P. fluorescens chromosome, which maintained the plasmid, despite its redundancy 17. These findings suggest that long-term plasmid maintenance largely depends on community context and on compatibility between plasmid and host, and, provided there are no restrictions on recombination of plasmid genes into the chromosome, is unlikely to be secured by positive selection for accessory genes alone 17,25,26.

Figure 3.

Figure 3

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Spread of chromosomally-acquired mercury resistance. (A) Frequency dynamics of focal Tn5042 insertions in P. putida chromosomes under 16 µg/g Hg(II) were tracked from transfer 9 (when insertions were first detected) to transfer 65. For each timepoint in each population, presence and frequency of the focal insertion was tested by PCR on ~30 clones using primers bridging the transposon and the chromosome (this giving a 95% chance of detecting a subpopulation comprising 10% of the total); other Tn5042 insertions were identified previously as pQBR57– merA+ clones 17. Plasmid-bearing clones were identified previously 17. Frequencies of different genotypes are indicated by filled stacked areas. Each panel represents an independent separate population: top row, single-species populations a, b, c, e; bottom row, co-cultured with P. fluorescens populations c, d, e, f. (B) Frequency dynamics of focal Tn5042 insertion in P. fluorescens chromosome under 0 µg/g Hg(II). This population was co-cultured with P. putida (replicate b).

Tn5042 also mobilised in the mercury-free treatments: we detected three instances of Tn5042 multiplying on plasmids, and one instance of Tn5042 copying to the P. fluorescens chromosome (which occurred by transfer 35, see Figure 3). Tn5042 insertions sometimes occurred multiple times in a lineage — in co-cultured P. putida with 16 µg/g Hg(II), one clone (from replicate c) ultimately carried six copies. Although Tn5042 copy number increased in some clones from the mercury-free treatment, we detected more copies in clones evolved under mercury selection (Z = –5.4404, p < 0.0001, n = 48, Exact General Independence Test). We did not detect any Tn5042 loss. For P. putida, Tn5042 tended to insert in a ~10 kb region near the origin of replication, while Tn5042 tended to insert in P. fluorescens near or inside Tn6291 (detected in 4/12 populations under mercury selection), in three cases representing the de novo formation of a composite resistance transposon. Here, Tn5042 became part of the cargo of Tn6291, broadening opportunities for spread, because subsequent events favouring Tn6291 mobilisation (perhaps different to those of Tn5042) will cause co-mobilisation of Tn5042 and its HgR genes. The pervasive mobility of Tn5042 supports a model in which TEs exploit plasmids to rapidly spread in the natural environment 5, consistent with sequence analysis suggesting Tn5042 was acquired relatively recently by pQBR57 (ref. 16) .

Surprisingly, we found plasmid size generally increased, primarily due to TE accumulation. Plasmid size in one clone evolved in 0 µg/g Hg(II) increased by over 10% compared with the ancestor (Figure 1). Increased plasmid size is expected to contribute to increased cost of plasmid carriage 27, however these results suggest such costs are negligibly small, and may be outweighed by transposition rates and/or general plasmid cost amelioration 26.

Discussion

The central role of HGT in adaptation is increasingly apparent, as ever-wider sequencing of isolates reveals the dynamic nature of microbial genomes 4,5,28. Between-species transfer of chromosomal genes occurred only where plasmid-encoded mercury resistance was not under positive selection and the plasmid persisted instead as an infectious element. Bacterial genome evolution is determined by the interaction between selection and recombination 29 — here, we observed that recombination indeed makes an increased contribution to genome evolution when selection is relaxed. The transferred genes were part of the ‘accessory’ genome, which can vary even between closely-related strains and is often more strongly associated with ecological niche than phylogenetic lineage 30. In this case, the transferred genes were located on TEs and putative transposases could be identified. This is relevant because TEs can transfer between replicons at a high rate 31, providing an efficient platform for the movement of genes between chromosomes and conjugative elements 2,3. Plasmids and TEs have a close — even symbiotic — relationship. TEs can comprise a substantial fraction of a plasmid genome 32,33 and where their genes are under positive selection they can boost the fitness of the plasmids that carry them due to genetic linkage. Similarly, unless they encode their own conjugative machinery, TEs must collaborate with elements such as conjugative plasmids to access new hosts 34; indeed models suggest that conjugative plasmids are required for TE survival and spread 35. Transposase activity can be affected by stress, for example nutritional deprivation or oxidative damage 36, and one intriguing possibility is that stresses caused by plasmid acquisition 37 could signal to a TE that a vehicle had arrived, triggering transposition and thus increasing rates of exchange from the chromosome to that plasmid. At least one Pseudomonad TE has been shown to increase activity following conjugation 38.

Our results provide rare direct experimental evidence of pervasive plasmid-mediated gene mobilization, transfer and acquisition in a simple soil microcosm community. This has profound implications for the spread of accessory genes in natural communities. Consistent with our findings, two recent studies of resistance plasmids in hospital outbreaks 5,39 indicate that TE mobilisation dominates plasmid evolution. Furthermore, both studies suggest that plasmids may have acquired TEs outside of patients, i.e. in the environment, where they are less likely to experience direct antibiotic selection. HGT vastly expands the evolutionary opportunities available to bacteria, allowing species to draw upon a collective mobile gene pool: our data indicate that environmental and ecological factors will be key modulators of the rate and extent of HGT in natural communities. HGT, particularly of antibiotic resistance and virulence genes, poses a major health concern 40 and understanding the ecology of HGT-mediated bacterial evolution will be crucial to predicting and designing interventions to prevent and mitigate such threats.

Materials and methods

Experimental design

The evolution experiment, described previously 17, was designed to understand the effect of an alternative host species on plasmid population dynamics and evolution. The experiment used Pseudomonas fluorescens SBW25 and P. putida KT2440 — representative soil Pseudomonas species, a widespread and naturally co-occurring genus 41 — and the 307-kb HgR plasmid pQBR57 which was isolated from the same geographic site as P. fluorescens SBW25 (ref. 16). Cultures were grown at 28°C and 80% relative humidity in soil microcosms consisting of 10 g twice-autoclaved John Innes No. 2 potting soil, supplemented with 900 µl sterile H2O or 900 µl HgCl2 solution. We used a fully-factoral design with two levels of mercury treatment (0 µg/g, or 16 µg/g); two levels of plasmid treatment (pQBR57+ starting with pQBR57-bearers at 50% frequency, or plasmid-free starting without plasmid); and three levels of culture treatment (single-species P. fluorescens, single-species P. putida, or co-culture with each species starting at 50% frequency). Six independent biological replicates (‘populations’) were initiated for each treatment, consistent with previous evolution experiments 42,43 and sufficient to detect differences in population dynamics between the treatments 17. Each replicate was initiated from independent single colonies, and populations were blocked by replicate to minimize confounding effects. The experiment was not blinded. To control for marker effects, replicates a–c used gentamicin-labelled (GmR) P. fluorescens and streptomycin-labelled (SmR) P. putida, whereas replicates d–f used SmR P. fluorescens and GmR P. putida. Samples of culture (100 µl soil wash) were serially transferred into fresh soil microcosms containing either H2O or HgCl2 every four days for 65 transfers (estimated as ~440 generations 17); this was decided before the experiment to be broadly consistent with other plasmid experimental evolution studies 42,43. At 16 points during and at the end of the experiment samples were spread on selective media to isolate clones, which were archived for subsequent analysis. After 65 transfers, a random number generator was used to select one plasmid-bearing and one plasmid-free clone (where present) from each pQBR57+ population for DNA sequencing. If plasmid-free or plasmid-bearing clones were present throughout the experiment but not at transfer 65, a clone from transfer 59 were used (this was the case for plasmid-bearing P. putida from single-species 16 µg/g replicate f, and plasmid-bearing P. putida from co-cultured 16 µg/g replicates a and c). We also sequenced ancestral clones, and three (single-species) or two (co-culture) clones from plasmid-free treatments to test for mutations occurring in the absence of plasmid. No Tn6291, Tn4652, or Tn6290 activity was detected in the plasmid-free treatments.

DNA sequencing and analysis

DNA was extracted using the QIAGEN DNeasy kit, prepared using the TruSeq Nano DNA Library Preparation Kit (350 bp insert size), and sequenced on the Illumina HiSeq platform. Reads trimmed using Cutadapt (version 1.2.1) and Sickle (1.200) were analysed using the Bacterial and Archaeal Genome Analyser pipeline 19, which uses the bwa short read aligner 44 and calls variants using the Genome Analysis Toolkit HaplotypeCaller 45. To identify structural variation (deletions, duplications and TE insertions) in the re-sequenced clones we used the BAGA module Structure, which uses a threshold ratio of non-proper to proper paired reads to identify putative genome disruptions. Reads mapping to putative disruptions were re-assembled using SPAdes 46, and contigs were aligned with the reference to identify structural variants. We also used two complementary approaches to identify structural variation: Breakdancer 47, and custom scripts that examined coverage for characteristic direct repeats introduced by TE insertion (increase in coverage of ≥25% over a <30 bp region, compared with neighbouring positions). These different approaches were broadly consistent, and all putative structural variants were examined using the Integrated Genome Viewer (IGV) 48. Because of differences between ancestral clones and the sequenced reference genome, variation appearing in all samples (including the ancestor) was removed from the analysis. In addition, apparent variation in hard-to-map regions (identified in an examination of parallel mutations in IGV) was considered unreliable and excluded (Supplementary Table 3). We also examined putative SNV called near TE insertions, and removed these manually if miscalled. Representative TE insertions were tested by PCR on clones, and in all cases yielded products of the anticipated size.

Sequence analysis of ancestral clones revealed that in three cases, pQBR57 had acquired a TE before the experiment was initiated, indicated by dotted lines in Figure 1. In the SmR P. fluorescens ancestor, pQBR57 had acquired Tn6290 at position 164349–164354. This event likely occurred in P. putida UWC1 during preparation as a donor for transfer of pQBR57 into P. fluorescens 16 because Tn6290 is present in P. putida UWC1 and not in P. fluorescens SBW25. In SmR P. putida, pQBR57 had acquired Tn4652 at 152552–152558 while in GmR P. putida pQBR57 had acquired Tn4652 at 162797–162802. These events may have either occurred in the donor P. putida UWC1 strain or in the recipient P. putida KT2440 strain, as both contain identical copies of Tn4652. In any case, TE insertion must have occurred rapidly as our stocks were all prepared from single colonies and ancestral pQBR57 contains neither Tn6290 nor Tn4652 (ref. 16) Tn4652 insertion into resident plasmids is consistent with previous work which found Tn4652 in pQBR plasmids pQBR55 and pQBR44 (ref. 16) presumably after acquisition by P. putida UWC1 (ref. 49).

PCR analysis of clones

We tested archived clones for TE insertions by PCR. Standard reactions were performed using GoTaq Green Master Mix (Promega), 0.4 µM each primer (Supplementary Table 4), and 0.2 µl archived culture on a program of 95°C for 5 minutes, followed by 30 cycles of 95°C for 30 seconds, 58°C for 30 seconds and 72°C for 1 minute, followed by a final extension of 72°C for 5 minutes. Tn6291 was detected in reisolated P. putida clones, and parallel reactions using primers targeting the P. fluorescens 16S rDNA locus were performed to rule out the presence of contaminating P. fluorescens.

Statistical analyses

To analyse the number of Tn5042 insertions in 0 µg/g and 16 µg/g Hg(II) we used the R package ‘coin’ to perform an Exact General Independence Test. To avoid pseudoreplication with populations from which >1 sample was sequenced, we analysed the mean number of Tn5042 insertions per species per population. To analyse the effect of mercury on conjugation dynamics we performed an Exact General Independence Test on plasmid distribution between donor and recipient after 5 transfers.

Supplementary Material

Supplementary Information

Acknowledgements

We thank Pia Koldkjaer and others at the Liverpool Centre for Genomic Research for assistance with sample preparation and sequencing. This work was supported by ERC Grant Agreement no. 311490-COEVOCON to MAB and a Philip Leverhulme Prize from Leverhulme Trust to MAB.

Footnotes

Author contributions: JPJH, EH, and MAB designed the study; JPJH collected data; JPJH, DW and SP analysed the data. JPJH and MAB drafted the manuscript. All authors discussed results and commented on the manuscript.

Author information: The authors declare no competing financial interests.

Data availability

Short read data is available at the European Nucleotide, Archive Project Accession PRJEB15009. Data presented in Figures 1–3 are on Dryad Digital Archive doi:10.5061/dryad.6gf28.

Code availability

The Bacterial and Archaeal Genome Analyser (BAGA) is available online at https://github.com/daveuu/baga . Representative scripts used to analyse our data are on Dryad Digital Archive doi:10.5061/dryad.6gf28.

References

  • 1.Halary S, Leigh JW, Cheaib B, Lopez P, Bapteste E. Network analyses structure genetic diversity in independent genetic worlds. Proceedings of the National Academy of Sciences. 2010;107:127–132. doi: 10.1073/pnas.0908978107. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 2.Norman A, Hansen LH, Sorensen SJ. Conjugative plasmids: vessels of the communal gene pool. Philosophical Transactions of the Royal Society B: Biological Sciences. 2009;364:2275–2289. doi: 10.1098/rstb.2009.0037. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 3.Frost LS, Leplae R, Summers AO, Toussaint A. Mobile genetic elements: the agents of open source evolution. Nat Rev Micro. 2005;3:722–732. doi: 10.1038/nrmicro1235. [DOI] [PubMed] [Google Scholar]
  • 4.Conlan S, et al. Plasmid Dynamics in KPC-Positive Klebsiella pneumoniae during Long-Term Patient Colonization. mBio. 2016;7:e00742–16. doi: 10.1128/mBio.00742-16. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 5.Sheppard AE, et al. Nested Russian Doll-Like Genetic Mobility Drives Rapid Dissemination of the Carbapenem Resistance Gene blaKPC. Antimicrobial Agents and Chemotherapy. 2016;60:3767–3778. doi: 10.1128/AAC.00464-16. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 6.Rasmussen S, et al. Early Divergent Strains of Yersinia pestis in Eurasia 5,000 Years Ago. Cell. 2015;163:571–582. doi: 10.1016/j.cell.2015.10.009. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 7.Johnson TJ, Nolan LK. Pathogenomics of the virulence plasmids of Escherichia coli. Microbiol Mol Biol Rev. 2009;73:750–774. doi: 10.1128/MMBR.00015-09. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 8.Stoodley P, Sauer K, Davies DG, Costerton JW. Biofilms as Complex Differentiated Communities. Annu Rev Microbiol. 2002;56:187–209. doi: 10.1146/annurev.micro.56.012302.160705. [DOI] [PubMed] [Google Scholar]
  • 9.Lopatkin AJ, et al. Antibiotics as a selective driver for conjugation dynamics. Nature Microbiology. 2016;1:16044. doi: 10.1038/nmicrobiol.2016.44. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 10.Stevenson C, Hall JP, Harrison E, Wood AJ, Brockhurst MA. Gene mobility promotes the spread of resistance in bacterial populations. The ISME Journal. 2017;63:1577. doi: 10.1038/ismej.2017.42. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 11.Jacoby GA, Rogers JE, Jacob AE, Hedges RW. Transposition of Pseudomonas toluene-degrading genes and expression in Escherichia coli. Nature. 1978;274:179–180. doi: 10.1038/274179a0. [DOI] [PubMed] [Google Scholar]
  • 12.Hedges RW, Jacob AE. In vivo translocation of genes of Pseudomonas aeruginosa onto a promiscuously transmissible plasmid. FEMS Microbiology Letters. 1977;2:15–19. [Google Scholar]
  • 13.Hemme CL, et al. Lateral Gene Transfer in a Heavy Metal-Contaminated-Groundwater Microbial Community. mBio. 2016;7:e02234–15. doi: 10.1128/mBio.02234-15. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 14.Xue H, et al. Eco-Evolutionary Dynamics of Episomes among Ecologically Cohesive Bacterial Populations. mBio. 2015;6:e00552–15. doi: 10.1128/mBio.00552-15. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 15.Gomez P, Buckling A. Bacteria-Phage Antagonistic Coevolution in Soil. Science. 2011;332:106–109. doi: 10.1126/science.1198767. [DOI] [PubMed] [Google Scholar]
  • 16.Hall JPJ, et al. Environmentally co-occurring mercury resistance plasmids are genetically and phenotypically diverse and confer variable context-dependent fitness effects. Environmental Microbiology. 2015;17:5008–5022. doi: 10.1111/1462-2920.12901. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 17.Hall JPJ, Wood AJ, Harrison E, Brockhurst MA. Source–sink plasmid transfer dynamics maintain gene mobility in soil bacterial communities. Proc Natl Acad Sci USA. 2016;113:8260–8265. doi: 10.1073/pnas.1600974113. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 18.Li P, Feng XB, Qiu GL, Shang LH, Li ZG. Mercury pollution in Asia: A review of the contaminated sites. Journal of Hazardous Materials. 2009;168:591–601. doi: 10.1016/j.jhazmat.2009.03.031. [DOI] [PubMed] [Google Scholar]
  • 19.Williams D, Paterson S, Brockhurst MA, Winstanley C. Refined analyses suggest that recombination is a minor source of genomic diversity in Pseudomonas aeruginosa chronic cystic fibrosis infections. Microbial Genomics. 2016;2 doi: 10.1099/mgen.0.000051. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 20.Silby MW, et al. Genomic and genetic analyses of diversity and plant interactions of Pseudomonas fluorescens. Genome Biol. 2009;10:R51. doi: 10.1186/gb-2009-10-5-r51. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 21.Ravindran A, Jalan N, Yuan JS, Wang N, Gross DC. Comparative genomics of Pseudomonas syringae pv. syringae strains B301D and HS191 and insights into intrapathovar traits associated with plant pathogenesis. Microbiologyopen. 2015;4:553–573. doi: 10.1002/mbo3.261. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 22.Kivistik PA, Kivisaar M, Horak R. Target Site Selection of Pseudomonas putida Transposon Tn4652. Journal of Bacteriology. 2007;189:3918–3921. doi: 10.1128/JB.01863-06. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 23.Levin BR, Stewart FM, Rice VA. The kinetics of conjugative plasmid transmission: fit of a simple mass action model. Plasmid. 1979;2:247–260. doi: 10.1016/0147-619x(79)90043-x. [DOI] [PubMed] [Google Scholar]
  • 24.Hausner M, Wuertz S. High rates of conjugation in bacterial biofilms as determined by quantitative in situ analysis. Applied and Environmental Microbiology. 1999;65:3710–3713. doi: 10.1128/aem.65.8.3710-3713.1999. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 25.Bergstrom CT, Lipsitch M, Levin BR. Natural selection, infectious transfer and the existence conditions for bacterial plasmids. Genetics. 2000;155:1505–1519. doi: 10.1093/genetics/155.4.1505. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 26.Harrison E, et al. Rapid compensatory evolution promotes the survival of conjugative plasmids. Mob Genet Elements. 2016;6:e1179074–7. doi: 10.1080/2159256X.2016.1179074. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 27.Baltrus DA. Exploring the costs of horizontal gene transfer. Trends in Ecology & Evolution. 2013;28:489–495. doi: 10.1016/j.tree.2013.04.002. [DOI] [PubMed] [Google Scholar]
  • 28.McCarthy AJ, et al. Extensive horizontal gene transfer during Staphylococcus aureus co-colonization in vivo. Genome Biol Evol. 2014;6:2697–2708. doi: 10.1093/gbe/evu214. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 29.Shapiro BJ. How clonal are bacteria over time? Current Opinion in Microbiology. 2016;31:116–123. doi: 10.1016/j.mib.2016.03.013. [DOI] [PubMed] [Google Scholar]
  • 30.Polz MF, Alm EJ, Hanage WP. Horizontal gene transfer and the evolution of bacterial and archaeal population structure. Trends Genet. 2013;29:170–175. doi: 10.1016/j.tig.2012.12.006. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 31.Sousa A, Bourgard C, Wahl LM, Gordo I. Rates of transposition in Escherichia coli. Biology Letters. 2013;9:20130838–20130838. doi: 10.1098/rsbl.2013.0838. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 32.Dennis JJ. The evolution of IncP catabolic plasmids. Current Opinion in Biotechnology. 2005;16:291–298. doi: 10.1016/j.copbio.2005.04.002. [DOI] [PubMed] [Google Scholar]
  • 33.Revilla C, et al. Different pathways to acquiring resistance genes illustrated by the recent evolution of IncW plasmids. Antimicrobial Agents and Chemotherapy. 2008;52:1472–1480. doi: 10.1128/AAC.00982-07. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 34.Carraro N, Rivard N, Burrus V, Ceccarelli D. Mobilizable genomic islands, different strategies for the dissemination of multidrug resistance and other adaptive traits. Mob Genet Elements. 2017;7:1–6. doi: 10.1080/2159256X.2017.1304193. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 35.Condit R, Stewart FM, Levin BR. The population biology of bacterial transposons: a priori conditions for maintenance as parasitic DNA. Am Nat. 1988;132:129–147. [Google Scholar]
  • 36.Twiss E, Coros AM, Tavakoli NP, Derbyshire KM. Transposition is modulated by a diverse set of host factors in Escherichia coliand is stimulated by nutritional stress. Mol Microbiol. 2005;57:1593–1607. doi: 10.1111/j.1365-2958.2005.04794.x. [DOI] [PubMed] [Google Scholar]
  • 37.Baharoglu Z, Bikard D, Mazel D. Conjugative DNA Transfer Induces the Bacterial SOS Response and Promotes Antibiotic Resistance Development through Integron Activation. PLoS Genet. 2010;6:e1001165–10. doi: 10.1371/journal.pgen.1001165. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 38.Christie-Oleza JA, Lanfranconi MP, Nogales B, Lalucat J, Bosch R. Conjugative interaction induces transposition of ISPst9 in Pseudomonas stutzeri AN10. Journal of Bacteriology. 2009;191:1239–1247. doi: 10.1128/JB.01071-08. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 39.He S, et al. Mechanisms of Evolution in High-Consequence Drug Resistance Plasmids. mBio. 2016;7:e01987–16–11. doi: 10.1128/mBio.01987-16. 11. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 40.O'Neill J. Tackling drug-resistant infections globally: final report and recommendations. London: Wellcome Trust & HM Government; 2016. [Google Scholar]
  • 41.Cho JC, Tiedje JM. Biogeography and degree of endemicity of fluorescent Pseudomonas strains in soil. Applied and Environmental Microbiology. 2000;66:5448–5456. doi: 10.1128/aem.66.12.5448-5456.2000. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 42.De Gelder L, Williams JJ, Ponciano JM, Sota M, Top EM. Adaptive Plasmid Evolution Results in Host-Range Expansion of a Broad-Host-Range Plasmid. Genetics. 2008;178:2179–2190. doi: 10.1534/genetics.107.084475. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 43.Harrison E, Guymer D, Spiers AJ, Paterson S, Brockhurst MA. Parallel compensatory evolution stabilizes plasmids across the parasitism-mutualism continuum. Curr Biol. 2015;25:2034–2039. doi: 10.1016/j.cub.2015.06.024. [DOI] [PubMed] [Google Scholar]
  • 44.Li H, Durbin R. Fast and accurate short read alignment with Burrows-Wheeler transform. Bioinformatics. 2009;25:1754–1760. doi: 10.1093/bioinformatics/btp324. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 45.McKenna A, et al. The Genome Analysis Toolkit: A MapReduce framework for analyzing next-generation DNA sequencing data. Genome Res. 2010;20:1297–1303. doi: 10.1101/gr.107524.110. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 46.Bankevich A, et al. SPAdes: A New Genome Assembly Algorithm and Its Applications to Single-Cell Sequencing. Journal of Computational Biology. 2012;19:455–477. doi: 10.1089/cmb.2012.0021. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 47.Chen K, et al. BreakDancer: an algorithm for high-resolution mapping of genomic structural variation. Nature Methods. 2009;6:677–681. doi: 10.1038/nmeth.1363. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 48.Robinson JT, et al. Integrative genomics viewer. Nature Biotechnology. 2011;29:24–26. doi: 10.1038/nbt.1754. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 49.Lilley AK, Bailey MJ, Day MJ, Fry JC. Diversity of mercury resistance plasmids obtained by exogenous isolation from the bacteria of sugar beet in three successive years. FEMS Microbiology Ecology. 1996;20:211–227. [Google Scholar]

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