• horizontal gene transfer;
  • bacterial community;
  • conjugation;
  • host range;
  • soil bacteria


  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results and discussion
  6. Acknowledgements
  7. References

Horizontal gene transfer by conjugation is common among bacterial populations in soil. It is well known that the host range of plasmids depends on several factors, including the identity of the plasmid host cell. In the present study, however, we demonstrate that the composition of the recipient community is also determining for the dissemination of a conjugative plasmid. We isolated 15 different bacterial strains from soil and assessed the conjugation frequencies of the IncP1 plasmid, pKJK10, by flow cytometry, from two different donors, Escherichia coli and Pseudomonas putida, to either 15 different bacterial strains or to the mixed community composed of all the 15 strains. We detected transfer of pKJK10 from P. putida to Stenotrophomonas rhizophila in a diparental mating, but no transfer was observed to the mixed community. In contrast, for E. coli, transfer was observed only to the mixed community, where Ochrobactrum rhizosphaerae was identified as the dominating plasmid recipient. Our results indicate that the presence of a bacterial community impacts the plasmid permissiveness by affecting the ability of strains to receive the conjugative plasmid.


  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results and discussion
  6. Acknowledgements
  7. References

Horizontal gene transfer (HTG) is a driving force in bacterial evolution as it allows bacteria to rapidly acquire complex new traits. Plasmids are one of the key vectors of HTG, enabling genetic exchange between bacterial cells across species and domain barriers (Poole, 2009; Boto, 2010), and they very often encode genes that confer adaptive traits to their host, such as antibiotic resistance, biodegradation pathways and virulence (de la Cruz & Davies, 2000). Transfer of these traits by conjugation requires the donor and the recipient cells to be in direct contact.

Different abiotic and biotic factors affect the range of conjugal exchange of genetic material between environmental bacteria, such as nutrient availability, spatial architecture of the bacterial community, plasmid donor and recipient relatedness and plasmid host type (van Elsas & Bailey, 2002; De Gelder et al., 2005; Sørensen et al., 2005; Seoane et al., 2011). The fraction of the cells in a community capable of receiving and maintaining conjugative plasmids is highly dependent on several of these factors and has been described as the plasmid permissiveness (Musovic, 2010).

It has been shown that conjugative plasmids express factors that favor the establishment of planktonic bacteria in biofilm communities, thereby increasing the chances for horizontal gene transmission (Ghigo, 2001; Reisner et al., 2006; Madsen et al., 2012). Complex interspecies communities facilitate synergistic interactions between populations, affecting the function, stability and flexibility of the community (James et al., 1995; Burmølle et al., 2006).

In the present work, HTG by conjugation between single populations and microbial communities isolated from soil were investigated. The plasmid transfer frequencies and the identities of the recipients of the plasmid, when hosted by different donors, were compared. The bacterial population was analyzed based on fluorescence properties and sorted by flow cytometry (FCM) to detect and quantify the plasmid transfer to the individual isolates and the mixed community (Muller & Nebe-von-Caron, 2010). Sequencing of the 16S rRNA gene from sorted transconjugant cells was used to evaluate the host range of the plasmid when a mixed microbial community was used as recipient.

Materials and methods

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results and discussion
  6. Acknowledgements
  7. References

Soil and leaf sampling

Soil samples were collected from an agricultural field in Tåstrup, Denmark, in the late summer of 2009. Soil was sampled from the 5- to 10-cm layer. The soil water content upon sampling was 14.2%, and the water holding capacity (WHC) was 60%. The soil was classified as sandy loam with pH 7.2. Leaves of baby maize seedlings were used for bacterial isolation. The seedlings were grown for 2 weeks in Tåstrup soil before harvesting.

Bacterial strains, plasmids, and growth media

Escherichia coli CSH26::lacIq and Pseudomonas putida KT2440::lacIq1, carrying pKJK10, a conjugative, green fluorescent protein (GFP) tagged IncP1 plasmid, originally isolated from soil (Sengeløv et al., 2001; Bahl et al., 2007) were used as donor strains. These strains were cultured in Luria Bertani (LB) broth supplemented with kanamycin monosulfate (50 mg mL−1); 1.5% (w/v) agar was added when solid medium was needed. The recipient strains (see below) were cultured in Tryptic Soy Broth medium (TSB; 17 g peptone from casein, 3 g peptone from soymeal, 2.5 g d(+)-Glucose, 5 g NaCl, 2.5 g K2HPO4 in 1 L distilled water, pH 7.3).

Isolation and identification of recipient strains from leaves incubated in soil

A 15 mg sample of a baby maize leaf was placed in 5 g Tåstrup soil adjusted to 40% WHC and incubated in triplicate at room temperature for 17 days. After 7, 12, and 17 days, the leaves were picked up from the soil, washed with PBS (8 g NaCl, 0.2 g KCl, 1.44 g Na2HPO4, and KH2PO4, adjusted to 1 L distilled water and pH 7.4), placed in a microfuge tube, added 1 mL PBS and vortexed for 1 min. DNA was extracted from the cell suspension as described below. Dilutions to 10−6 were made and 100 μL were plated in triplicate onto Tryptic Soy Agar (TSA; Difco) 10% supplemented with cycloheximide (50 mg mL−1) and incubated at 25 °C for 2–5 days. Sixteen colonies from each triplicate looking phenotypically different were isolated and purified for DNA extraction. A denaturation gradient gel electrophoresis (DGGE) analysis was performed with 16S rRNA gene PCR fragments.

From the total of 48 strains from day 7, 15 morphologically different strains were selected for the use as recipients. The strains were grown overnight (ON) in 5 mL TSB, the DNA was extracted using ‘Genomic Mini for Universal Genomic DNA Isolation Kit’ (A&A Biotechnology) and the 16S rRNA gene sequences were amplified with primers 27F and 1492R (Lane, 1991) for identification. The PCR mixture contained 0.5 μL DNA, 1XPhusion GC buffer, 0.2 mM dNTP mixture, 1 U Phusion Hot Start DNA Polymerase (FinnzymesOy, Espoo, Finland) and 0.5 μM of each primer (TAG Copenhagen A/S, Denmark). The final volume was adjusted with DNA-free water to 50 μL. Amplification was as follow: initial denaturation at 98 °C for 30 s, followed by 35 cycles at 98 °C for 10 s, at 55 °C for 30 s and at 72 °C for 45 s. A final primer extension reaction was performed at 72 °C for 6 min. The resulting sequence (1480 bp) was compared with reference sequences by BLAST search (Altschul et al., 1997) and aligned with them using clustalx 1.7 program (Thompson et al., 1997). Maximum-likelihood analyses were performed using PhyML (Guindon & Gascuel, 2003). modeltest 3.06 (Posada, 2008) was used to select appropriate models of sequence evolution by the Akaike Information Criterion. The confidence at each node was assessed by 500 bootstrap replicates. Similarities among sequences were calculated using the MatGAT v.2.01 software (Campanella et al., 2003). Taxonomic assignment was carried out based on the Roselló-Mora and Aman criteria (Rosselló-Mora & Amann, 2001).


The cells from the leaves-PBS solution and from the 48- to 15-strain pools were lysed by bead beating followed by DNA extraction as specified above. The DNA was used for a 16S rRNA gene PCR as described above and 1 μL of the product was used as a template for a new PCR using internal primers with a GC clamp 341F and 518R (Muyzer et al., 1993) and a polymerization step at 72 °C for 20 s. This PCR product was loaded onto the DGGE gel, containing a denaturation gradient of 30–70% acrylamide, and an electrophoresis was run in a Dcode system (Biorad) at 60 °C and 70 V for 17 h. The gel was stained with SYBRGold (Invitrogene) in the dark for 45 min.

Filter mating assays

Prior to filter matings, the donor strains were grown in 5 mL LB broth at 250 r.p.m. at 30 °C (P. putida) and 37 °C (E. coli) for 18 h. These ON cell cultures were then diluted 1 : 10 in fresh LB medium and grown under similar conditions for three more hours to reach exponential growth phase (OD600 ≈ 0.6). The cells were then recollected, washed twice, and resuspended in sterile PBS. The recipient strains were cultured similarly in TSB at 25 °C. The lack of background fluorescence of the donor and recipient strains was verified in the flow cytometer (see specifications below) prior to their use in the filter mating assay.

For the single-strain mating experiments, 10 μL of donor and recipient, respectively, were spotted onto 0.2 μm nitrocellulose filters in triplicate, mixed, placed on TSA and R2A (yeast extract 0.5 g, proteose peptone 0.5 g, casamino acids 0.5 g, glucose 0.5 g, soluble starch 0.5 g, sodium pyruvate 0.3 g, K2HPO4 0.3 g, MgSO4·7H2O 0.05 g, agar 15 g in 1 L distilled water) plates and incubated at 25 °C for 20 h. The cells were then harvested from the filter followed by resuspension in 1 mL PBS, and FCM analysis as specified below. For the microbial community, we spotted 5 μL of each isolate (OD600 ≈ 0.3–0.7) and 75 μL of donor strain (either P. putida or E. coli, prepared as described above) onto the filter, incubated and analyzed by FCM at the same conditions as for the single-strain matings. Controls with only donors or recipients were included.

Quantification and sorting by FCM

Flow cytometric enumeration of cells was carried out with a FACScalibur flow cytometer (Becton Dickinson, San Jose, CA) equipped with a 15 mW argon laser (488 nm). The following settings and voltages were used during analysis: forward scatter = E01, side scatter (SSC) = 350, and the fluorescent detectors FL1 (530/30 nm), FL2 (585/42 nm), FL3 (650/30 nm) were set at 510 V. A threshold was set on the SSC, and no compensation was used. All parameters were on logarithmic mode. Samples were run at the ‘low’ flow rate setting for 1 min.

All the samples were diluted in PBS before flow enumeration to assure optimal bacterial counts to 2000 events s−1. In part of the sample (100 μL), gfp-expression was induced by incubation in LB with 1 mM of isopropyl-b-D-1-thiogalactopyranoside (IPTG, SIGMA) for 3 h at 30 °C (P. putida) and 37 °C (E. coli) to determine the number of donor cells (Musovic et al., 2006).

To isolate and identify recipients from the E. coli-community mating, one subsample of each replicate of the cell extract was diluted to 1000 events s−1 to flow-sorted (MoFlo; DAKO) at a flow rate of 400–1000 events s−1, with an optimal setting of the sheath pressure of ca. 60 psi and drop drive frequency to ca. 95 kHz, using a 70-μm CytoNozzle tip on an enrichment sort option of single-mode per single drop envelope. Dilutions up to 10−3 were made from approximately 70 000 cells of each replicate, and 100 μL of each dilution were plated on TSA plates supplemented with kanamycin, streptomycin (100 mg mL−1) and tetracycline (20 mg mL−1) and incubated at 25 °C for 2–5 days. Four green colonies of each replicate were selected for DNA extraction and identified by sequencing after the amplification of the 16S rRNA gene as described above.

Data analysis was carried out with the cellquest software package. Two polygonal gates were defined in bivariate FL1 vs. FL2 to count for green cells and in bivariate SSC vs. FL2 density plot as a double check.

Statistical analysis

All microcosmic experiments were carried out in triplicate. Standard deviations were calculated with Excel (Microsoft®). A Student's t-test was applied and probabilities less than 0.05 were considered significant.

Results and discussion

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results and discussion
  6. Acknowledgements
  7. References

Isolation of the bacterial community

Bacterial strains established on the leaves embedded in soil were isolated to obtain a highly diverse bacterial community with the capability of attachment. We used maize leaves as the plants grow fast with no specific requirements. The diversity of the established community over time was followed by DGGE analysis. DGGE is a simple and fast method to screen and compare the diversity of a bacterial community, well suited for this study. The number of bands in a DGGE lane reflects the degree of bacterial diversity and lanes from the same gels can be compared to explore changes in diversity (Muyzer et al., 1993). Based on the number of bands associated to the sampling days 7, 12, and 17 (Fig. 1, lane 1, 4, and 7, respectively), a highly diverse community was observed from day 7 and onwards. This was confirmed by comparing colony morphology of the 48 isolated strains from the different sampling points (data not shown). Due to this, and the fact that the leaves were at this time point highly decomposed (data not shown), the day 7-samples were chosen for strain isolation.


Figure 1. Denaturing gradient gel electrophoresis (DGGE) analysis of the diversity of the bacterial communities isolated from maize leaves after 7, 12, and 17 days of incubation in soil. The DGGE gel shows the PCR amplified products of the 16S rRNA genes of the total bacterial consortia present on the leaves on the sampling days 7, 12, and 17 (lane 1, 4, and 7, respectively). The 16S rRNA gene profiles of a total of 48 cultured strains from each day are presented in lane 2 (day 7), lane 5 (day 12), and lane 8 (day 17). Of these 48, 15 morphologically different strains were selected to constitute representative communities (lanes 3, 6, and 9) of sampling days 7, 12, and 17, respectively. The DGGE analysis indicated that most bacterial diversity was preserved when reducing the community from 48 to 15 strains (by comparing lane 2–3, 5–6, and 8–9). The 15-strain community from sampling day 7 was used for gene transfer analysis.

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To select a manageable, yet still diverse, subcommunity from all of the isolated strains of the day 7 sampling, the colony morphology of all the 48 isolates (from the three replicates) was visually compared. Fifteen isolates appearing morphologically different were chosen, and their DGGE profile was compared with that of the 48 isolates (Fig. 1, lane 2, and 3). Based on the number of bands, a low number of strains were lost when the subcommunity was selected, but most of the initial diversity was represented in the selected community (lane 3). From the 16S rRNA gene sequencing analysis (Table 1), the isolates were identified as typical soil bacteria, mostly gram negatives, with the Pseudomonas and Chryseobacterium genera as the most abundant. Based on this, the strains of the selected subcommunity were considered as well-suited potential recipients of the pKJK10 plasmid.

Table 1. Identification of the isolated strains by 16S rRNA gene analysis
Strain nameSimilarity (%)a
  1. a

    Similarity from sequenced 16S rRNA genes calculated with the matgat v.2.01 software (see text for more details).

Flavobacterium psychrolimnae 97.8
Pseudomonas lutea 98.1
Pseudomonas brassicacearum 99.6
Pseudomonas fluorescens 99.7
Ochrobactrum rhizosphaerae 100
Chryseobacterium soldanellicol 98.2
Chryseobacterium letacus 98.5
Sphingobacteriaceae 93.7
Xanthomonas retroflexus 99.6
Micrococcaceae 94.2
Chryseobacterium ginsengisoli 99.2
Stenotrophomonas rhizophila 99.6
Microbacterium oxydans 100
Ensifer adherens 98.9
Janthinobacterium lividum 99.7

Transfer of pKJK10 to the individual soil isolates and the mixed community

The donor strains used in this study encode the lacIq1 repressor gene from the chromosome, repressing GFP expression from pKJK10 when present in these donor strains, as the lac promoter regulates GFP expression in this plasmid. Due to the lack of the lacIq1 repressor in the soil isolates, GFP will be expressed if the plasmid is transferred into these cells. This system thus allows enumeration of transconjugants and donors by direct sample analysis and after IPTG induction, respectively (Sørensen et al., 2003). Detection by FCM has several advantages in such approaches because enumeration of transconjugant cells is based solely on fluorescence markers. There is therefore no need for only including strains with specific antibiotic resistance profiles in the recipient community, and the strains do not need to be capable of expressing the resistance traits encoded by the plasmid to be characterized as transconjugants.

The transfer frequency of the conjugative plasmid from the two different donors to the soil isolates was calculated as a transconjugant/donor ratio. No green cells were observed in the negative controls with only the recipient strains present (data not shown), indicating that none of the soil isolates produced auto-fluorescence and that green cells represented plasmid transfer events.

When E. coli was used as donor, no transfer of pKJK10 was detected to any of the individual 15 soil isolates, but P. putida was observed to transfer pKJK10 to Stenotrophomonas rhizophila. The plasmid transfer frequency from P. putida to S. rhizophila was higher when the filters were placed on TSA medium (1.07 ± 3.05 × 10−1) compared with R2A medium (0.33 ± 2.32 × 10−2, Table 2), supporting the fact that the metabolic state of the cells may in some cases influence conjugation frequencies (van Elsas & Bailey, 2002). These results reflect the fact that the host range of plasmids depends on the identity of the donor strain (De Gelder et al., 2005).

Table 2. Transfer frequencies of pKJK10 by different donors to individual recipients and mixed communitiesa
  1. a

    All values are transconjugants/donor ratios ± standard deviation of triplicate experiments.

  2. b

    No transconjugants were obtained on the filter (below detection limit).

  3. c

    Number of transconjugants obtained when placing the filters on R2A plates.

E. coli CSH26::lacIq S. rhizophila b
Mixed community1.56 ± 3.43 × 10−1
P. putida KT2440::lacIq S. rhizophila

1.07 ± 3.05 × 10−1

0.33 ± 2.32 × 10−2c

Mixed communityb

In contrast to the results observed when transferring pKJK10 to individual isolates, no plasmid transfer events were observed from P. putida to the mixed community consisting of the same 15 strains applied individually above. Transconjugants were, however, obtained when applying E. coli as donor of pKJK10. The green fluorescent transconjugant cells were sorted by FACS and cultured on TSA agar plates. By sequence analysis of the 16S rRNA gene from four colonies from each replicate, the selected transconjugants were shown all to be identical and identified as Ochrobactrum rhizosphaerae. This does not exclude the possibility that other isolates may also have received the plasmid, but it does show that O. rhizosphaerae in fact did so and that it was the most dominant strain among the plasmid recipients. Interestingly, O. rhizosphaerae was not able to receive the plasmid in the individual mating experiment, indicating that the plasmid permissibility does not only depend on the abilities of the plasmid, host and recipient strains, but also on the surrounding microbial community, which may reduce or enhance plasmid transfer. Both of these scenarios were observed in this study; transfer of pKJK10 from P. putida to S. rhizophila was observed in diparental mating experiments, but not in a mixed community, possibly caused by reduced survival/competition ability of the strains or by the fact that the donor and this specific recipient populations had less opportunity for interaction in the mixed community. In contrast, the presence of a mixed community induced pKJK10 transfer from E. coli to O. rhizosphaerae, which may be due to altered physical cell–cell interaction or the presence of one or several intermediate plasmid host(s). These ‘plasmid step-stones’ may facilitate plasmid transfer from E. coli to O. rhizosphaerae, but are unable to establish and stabilize the plasmid in their own population. Because it was not possible to isolate the strains individually after growth in the community, the fraction of O. rhizosphaerae herein could not be determined; It is possible that O. rhizosphaerae is the dominating strain in the consortium or the most metabolically active, explaining its enhanced abilities as plasmid recipient. Regardless of this strain being dominant or representing a minor population of the community, it is still intriguing that no plasmid transfer was observed in the dual-strains mating from E. coli to O. rhizosphaerae.

The results of this study indicate that the surrounding bacterial community strongly impacts the plasmid host range, which needs to be considered when analyzing potential plasmid dissemination in natural environments in association to risk assessment. Plasmid mediated traits, including antibiotic resistance and virulence, may spread to natural bacterial populations in situ, in spite of an apparent narrow host range detected in simple, dual-strain-mating experiments.


  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results and discussion
  6. Acknowledgements
  7. References

This research was supported by funding to Søren Sørensen by The Danish Council for Independent Research (Natural Sciences), The Danish Council for Independent Research (Technology and Production) (ref no: 09-090701, Mette Burmølle) and the Department of Biotechnology and Bioengineering (Cinvestav, Mexico). Claudia I. de La Cruz-Perera received grant-aided support from ‘ConsejoNacional de Ciencia y Tecnologia’ (CONACyT, Mexico) scholarship 166878.


  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results and discussion
  6. Acknowledgements
  7. References
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