Root growth and exudate production define the frequency of horizontal plasmid transfer in the Rhizosphere (original) (raw)

Abstract

To identify the main drivers of plasmid transfer in the rhizosphere, conjugal transfer was studied in the rhizospheres of pea and barley. The donor Pseudomonas putida KT2442, containing plasmid pKJK5::gfp, was coated onto the seeds, while the recipient P. putida LM24, having a chromosomal insertion of dsRed, was inoculated into the growth medium. Mean transconjugant-to-donor ratios in vermiculite were 4.0±0.8 × 10−2 in the pea and 5.9±1.4 × 10−3 in the barley rhizospheres. In soil, transfer ratios were about 10 times lower. As a result of a 2-times higher root exudation rate in pea, donor densities in pea (1 × 106–2 × 109 CFU g−1 root) were about 10 times higher than in barley. No difference in recipient densities was observed. In situ visualization of single cells on the rhizoplane and macroscopic visualization of the colonization pattern showed that donors and transconjugants were ubiquitously distributed in the pea rhizosphere, while they were only located on the upper parts of the barley roots. Because the barley root elongated about 10 times faster than the pea root, donors were probably outgrown by the elongating barley root. Thus by affecting the cell density and distribution, exudation and root growth appear to be key parameters controlling plasmid transfer in the rhizosphere.

Introduction

Whole-genome sequence comparisons have provided retrospective evidence suggesting that extensive horizontal exchange of genes has taken place between bacterial species (van Elsas et al., 2003). Hence, horizontal gene transfer (HGT) is believed to be important for bacterial evolution (Ochman et al., 2000; Sørensen et al., 2005). Retrospective evidence of HGT, however, does not explain the dynamics of the transfer process (Rensing et al., 2002), and studies are therefore needed to obtain actual measurements of transfer frequencies, and to provide an understanding of which environmental conditions govern HGT in the environment.

Horizontal plasmid transfer is one of the mechanisms by which bacteria may exchange DNA. Several studies of plasmid transfer have been published (see Davidson, 1999 and Dröge et al., 1999 for reviews), and a number of environmental ‘hot spots’ have been identified. Among these are the rhizospheres of plants (Lilley et al., 1994; Dröge et al., 1999). The elevated plasmid transfer in the rhizosphere compared with that in bulk soil has been attributed to relatively high cell densities and/or accessible root exudates that stimulate the bacterial activity (van Elsas et al., 1988, 2003; Kroer et al., 1998). Recent data have indicated that some rhizosphere-associated hot spots are ‘hotter’ than others (Schwaner & Kroer, 2001; Sengeløv et al., 2001), but a relationship between conjugation, root exudate production, metabolic activity and cell density has not been established to explain the difference between the rhizosphere hot spots.

The aim of the present study was to identify the main drivers of horizontal plasmid transfer in the rhizosphere. We compared transfer in the rhizospheres of barley and pea, as models of plants having different root types and representing dicots and monocots, respectively. The two plant species were ideal for the investigation, because it has been demonstrated that the pea rhizosphere is 10 times more conducive to plasmid transfer than barley (Schwaner & Kroer, 2001). Using a combination of in situ visualization of single cells on the rhizoplane, macroscopic visualization of the colonization pattern of donor and transconjugant cells in the rhizosphere, quantification of the amount and quality of the root exudates, and measurements of the root growth, we were able to establish a relationship between production of root exudate, root growth, spatial distribution of the bacterial cells and horizontal plasmid transfer.

Materials and methods

Bacterial strains, plasmids, and growth media

Conjugal gene transfer was studied in biparental mating experiments. Pseudomonas putida KT2442, containing plasmid pKJK5::gfp (Sengeløv et al., 2001), served as the donor. pKJK5 is a natural rhizosphere plasmid belonging to an outbranch of the IncP1- incompatibility group (L. Mølbak & N. Kroer, unpublished data). The recipient, P. putida LM24, had a chromosomal insertion of the red fluorescent protein gene, dsRed. The donor was constructed by inserting pKJK5::gfp into P. putida KT2442 from P. putida KT2440/pKJK5::gfp (Sengeløv et al., 2001) by biparental mating as previously described (Mølbak et al., 2003). The recipient strain was constructed by insertion of a mini-Tn_5_ gene cassette, with the dsRred gene fused to the Escherichia coli ribosomal promoter, rrnB_P1 (Tolker-Nielsen et al., 2000), into the chromosome of P. putida SM1464 by triparental mating according to Herrero et al. (1990). Escherichia coli Mv1190λ_pir/TTN151 and E. coli HB101/pRK600 were used as donor and helper strains, respectively. A clone, which grew at the same rate as SM1464 in minimal medium at 30°C, was selected and named P. putida LM24. Characteristics of the strains and plasmids used are listed in Table 1.

1

Strains and plasmids used in this study

1

Strains and plasmids used in this study

Escherichia coli strains were grown in Luria–Bertani (LB) medium at 30°C. Pseudomonas putida strains were grown at 30°C in FAB minimal medium [1 mM MgCl2, 0.1 mM CaCl2, 0.01 mM FeCl2, 0.15 mM (NH4)SO4, 0.33 mM Na2HPO4, 0.2 mM KH2PO4, 0.5 mM NaCl] (Ramos et al., 2000) containing 10 mM sodium citrate as the sole carbon source. When needed, antibiotics were added at the following concentrations: kanamycin (Km) 25 μg mL−1, nalidixic acid (Nal) 50 μg mL−1, tetracycline (Tc) 10 μg mL−1, and rifampicillin (Rif) 50 μg mL−1.

Rhizosphere microcosms

Microcosms consisted of 50-mL polystyrene tubes (Sarstedt AG & Co., Nümbrecht, Germany) filled with 45 mL of autoclaved vermiculite or 55 g of nonsterile soil. One barley (Hordeum vulgare var. Alexis) or pea (Pisum sativum var. Ping Pong) seed per microcosm was sown at about 1-cm depth. The soil was a sandy loam obtained from Taastrup near Copenhagen, Denmark (Kragelund & Nybroe, 1996). To enable investigation of the effect of root growth on plasmid transfer, the donor was inoculated onto the seeds, while the recipient was inoculated into the vermiculite/soil. Prior to sowing, seeds were coated with the donor strain by soaking in a stationary-phase culture of P. putida KT2442/pKJK5::gfp for 30 min at 20°C. The donor culture was washed twice in M8 buffer (22 mM Na2HPO4·2 H2O, 22 mM KH2PO4, 100 mM NaCl) (Ramos et al., 2000). Donor densities of c. 5 × 106 CFU per pea seed and 7 × 106 CFU per barley seed were achieved. The recipient strain was inoculated into the vermiculite or soil to a density of c. 3 × 105 CFU mL−1 vermiculite or 4 × 106 CFU g−1 soil. A total volume of 30 mL of inoculum in germination buffer [22 mM Na2HPO4, 22 mM KH2PO4, 100 mM NaCl, 1 mM MgSO4, 6 mg L−1 Fe(III) ammonium citrate (28% Fe), 8.2 μM ZnCl2, 0.6 μM H3BO4, 19.8 μM MnCl2, 3.6 μM CoCl2, 35.7 μM NiCl2, 51.1 μM CuCl2, 24.2 μM Na2MoO4] was added to the vermiculite microcosms, while autoclaved water inoculated with the recipient was used to adjust the moisture content of the soil microcosms to 15% (w/w). The microcosms were placed in a glass beaker containing moist filter paper at the bottom, and the beakers placed in transparent plastic bags in a growth chamber at 20–22°C with a 12 h : 12 h light–dark cycle.

Sampling of rhizosphere microcosms

At each sampling time, five replicate microcosms containing vermiculite and five replicate microcosms containing soil were sacrificed. The seedlings were carefully removed and the roots separated from the seed and placed in 10-mL plastic tubes containing 5 mL of cold (4°C) saline. The bacteria were extracted from the rhizosphere by vortexing for 3 min, and dilutions plated on FAB plates containing 10 mM sodium citrate and appropriate antibiotics for the selection of donors, recipients and transconjugants. The plates were incubated at 30°C for 24 h before counting numbers of CFU.

Parallel to sampling, the significance of mating on the transconjugant-selective plates was assessed. This was carried out by combining extracts of control microcosms inoculated with only donors with extracts of microcosms containing only recipients, and plating the mixture on transconjugant-selective plates. False-positive transconjugants were never detected.

Root colonization patterns

The distribution of donor and transconjugant cells on the roots of the barley and pea plants was assessed at each sampling time. Two replicate roots of each barley and pea microcosm were harvested by carefully removing the plants from the microcosms. For each of the plant species, one root was placed on a FAB agar plate with 10 mM sodium citrate, while the other was placed on a FAB agar plate containing 10 mM sodium citrate and 50 μg mL−1 Nal (Nal was added to prevent mating on the plates). Care was taken to ensure that tweezers were flamed before each touch on the agar plate and that the full length of the roots was in contact with the surface of the agar. The roots were photographed with white light using a CCD camera (Hamamatsu C5985). After one hour of incubation, roots were removed and the plates incubated for 18 h at 30°C. After the incubation, bacterial colonies on plates without Nal were replica-plated onto new FAB plates with rifampicillin and tetracycline to allow growth of the donor cells. Colonies on FAB agar plates containing Nal were replica-plated onto new FAB plates with Nal and tetracycline to allow growth of the transconjugant cells. Following incubation for 24 h at 30°C, plates were illuminated with UV-light and the autofluorescent light from the colonies recorded by a CCD camera. Images of the roots were superimposed on the images of the bacterial colonies using photoshop software (Adobe, Mountain View, CA).

Donor colonization in the presence of growth inhibitors

Pea seedlings inoculated with donors were grown in vermiculite microcosms amended with various concentrations of Nal (0, 50 or 100 mg L−1). After 6 days of incubation, the colonization patterns of three replicate pea seedlings were visualized as described above. To test the potential toxicity of Nal towards the donor strain, batch cultures in germination buffer were treated with 50, 100, 200 or 500 mg L−1 Nal. After one hour of incubation at room temperature, numbers of CFUs in amended cultures were compared with an unamended control.

Bacterial distribution at the microscale level

Rhizosphere chambers were used for in situ visualization of donors, recipients, and transconjugants on the rhizoplane of pea and barley seedlings. The rhizosphere chambers were constructed and used basically as described by Ramos et al. (2000). Briefly, chambers (length, 55 mm; width, 20 mm; depth, 10 mm) were constructed by gluing (with silicone) a bended plastic tube on top of a microscope slide onto which a coverslip (24 × 60 mm) was sealed. The chambers were filled with vermiculite into which 1 mL of recipient culture was inoculated (final density: 5 × 105 CFU mL−1). Pea and barley seeds, pregerminated for 1 day and coated with donor bacteria (see above), were sown just beneath the surface of the vermiculite. As controls, rhizosphere chambers consisting of donor-coated seeds in uninoculated vermiculite, and recipient-coated seeds in uninoculated vermiculite were set up. Rhizosphere chambers were incubated at 20°C in the dark for 4–5 days. To ensure that the roots were growing along the coverslip, growth chambers were placed on a rack at an angle of c. 45°. At various time points, images of the rhizoplane-attached cells were obtained using confocal microscopes (models TCS4D and TCS SP1; 3-channel scanning; Leica Microsystems, Heidelberg GmbH, Germany) equipped with detectors and filter sets that simultaneously monitored Gfp and Dsred. Simulated fluorescence projections and vertical cross-sections were generated using the imaris software package (Bitplane AG, Zürich, Switzerland). Images were processed for display using the photoshop software version 6.0.

Quantitative and qualitative measurements of root exudates

To estimate exudate production by the barley and pea seedlings, sterile seeds were prepared and grown aseptically. The barley seeds were sterilized by sequential treatment with 50% H2SO4 for 1 h, two rinses in sterile water, and 6 min in 0.1% AgNO3. Finally, seeds were rinsed three times in sterile water. Seed coats were removed manually after treatment with H2SO4. Pea seeds were sterilized as previously described for bean (Normander et al., 1998). After sterilization, seeds were pregerminated on moist filter paper for 48 h. To test for sterility, a subsample of 10–20 seeds was placed on LB agar plates for 48 h at 20°C.

Ten pregerminated seeds were sown in air-tight glass jars (10 cm high and 10 cm wide) containing 200 mL of sterilized vermiculite saturated with 130 mL of germination buffer. The glass jars were incubated in a growth chamber at 20–22°C with a 12 h : 12 h light–dark cycle for 3 or 6 days.

Harvesting of the exudates was performed by the addition of 130 mL of germination buffer followed by gentle stirring for 3 min. The solution was decanted and filtered through a 0.2-μm Micron PES membrane (Frisenette; Ebeltoft; Denmark). To test for sterility, an aliquot of 100 μL was removed before the filtration step and spread on a LB agar plate and incubated at 20°C for 5 days. The sterile exudate solutions were stored at 4°C.

The content of total dissolved organic carbon was measured on a Shimadzu TOC 5000 Analyzer (oxidation at 680°C, Pt-covered alumina beads used as catalyst) (Kroer et al., 1998). To assess potential qualitative differences between the barley- and pea-root exudates in terms of supporting bacterial growth, an exponentially growing culture of P. putida KT2442/pKJK5::gfp, washed twice in saline and incubated for 24 h at 30°C, was inoculated to an OD of 0.005 (OD450 nm) in the exudate solutions and grown in batch at 30°C. Growth was followed by measuring the OD at appropriate time points.

Data analysis

Statistical tests were performed using the statview® software (SAS Institute Inc., NC). The difference between means was tested by anova on log10-transformed data.

Results and discussion

Population changes in rhizospheres

Donor densities in pea rhizospheres were from seven (vermiculite) to 60 (soil) times higher than in barley rhizospheres (P<0.01). For pea, numbers ranged between 7 × 106 and 2 × 109 CFU g−1 root in vermiculite, and between 1 × 106 and 2 × 109 CFU g−1 root in soil (Fig. 1). For barley, densities varied between 2 × 106 and 2 × 108 CFU g−1 root in vermiculite, while they were approximately one order of magnitude lower in soil. Differences in the ability of pseudomonads to colonize different plant roots have previously been reported. For instance, P. putida KT2440 was shown to establish higher population densities in the rhizosphere of broad bean than in that of corn (Molina et al., 2000).

1

Densities of donors, recipients and transconjugants in the rhizospheres of barley (left) and pea (right) grown in vermiculite (○) and soil (●). Lines are means of five replicate microcosms superimposed over individual data points. Plasmid transfer was not assessed at the start of the experiment but was assumed to be zero. Dashed lines, therefore, represent the expected development of numbers of transconjugants. The data shown represent one out of a total of three independent plasmid transfer experiments, all of which produced basically similar results.

Numbers of recipient bacteria extracted from the barley and pea rhizospheres varied between 5 × 106 and 1 × 109 CFU g−1 root in vermiculite and between 1 × 105 and 4 × 108 CFU g−1 root in soil (Fig. 1). Numbers of recipient bacteria from roots grown in vermiculite were significantly higher than those from roots grown in soil (P<0.001). Within each type of growth medium, however, no significant differences (_P_>0.05) between pea and barley were observed. The similar population sizes in pea and barley were probably a result of the inoculation of the recipient into the growth medium.

Maximal numbers of transconjugants were attained after 2–3 days, with the highest densities observed in the pea–vermiculite rhizosphere (c. 4.5 × 107 CFU g−1 root) followed by the pea–soil rhizosphere (c. 2.5 × 106 CFU g−1 root) (Fig. 1). Maximal transconjugant densities in the barley rhizospheres were c. 1.5 × 106 CFU g−1 root in vermiculite and c. 1.5 × 104 CFU g−1 root in soil. In soil microcosms, numbers of transconjugants generally declined after 2–3 days, while numbers were relatively constant in the vermiculite microcosms (Fig. 1).

Transfer ratios

Transfer ratios (transconjugants/donors; T/D) of pea rhizospheres were significantly higher (P<0.001) than those of barley rhizospheres when comparing data within the same growth medium (Fig. 2). In pea, mean T/D values of 4.0±0.8 × 10−2 in vermiculite and 3.5±0.7 × 10−3 in soil were observed, while in barley, the equivalent values were 5.9±1.4 × 10−3 and 2.0±0.5 × 10−4, respectively (Fig. 2). Thus, in vermiculite, the average T/D of pea was seven times higher than that of barley. In soil it was 17 times higher. The highest T/D value (0.15) was measured in one of the pea–vermiculite microcosms on day 6 (Fig. 2).

2

Transfer ratios (transconjugants/donors) in the rhizospheres of barley (solid lines) grown in vermiculite (○) and soil (●), and pea (dashed lines) grown in vermiculite (◻) and soil (▪). Lines are means of five replicate microcosms superimposed over individual data points.

Sengeløv et al. (2001) found T/D values of 4.4 × 10−3 in the barley rhizosphere using the same donor strain, plasmid, and soil type as in our study. Thus, their transfer ratios were a factor of 20 higher than those of our barley–soil microcosms. The relatively high transfer ratios observed by Sengeløv et al. were probably a result of the fact that the donor and recipient cells were inoculated at higher densities (4 × 108 donors and 6 × 108 recipients per g of root). Moreover, because both cell types were inoculated into the soil, establishment of mating pairs was possible in the rhizosphere and bulk soil throughout the experiment, whereas in our case mating pair formation relied on spreading of the donor with the root.

Schwaner & Kroer (2001) reported the pea rhizosphere to be 10 times more conducive to plasmid transfer than the barley rhizosphere. Similarly, we showed the pea rhizosphere to be from seven to 17 times more conducive. Thus, despite the fact that we used other bacterial strains, a different plasmid and a different inoculation procedure, our data confirmed the apparent species-specific difference between pea and barley. Differences in the activity of the bacteria have been suggested to be an important determinant for transfer (Lilley et al., 1994; Sørensen & Jensen, 1998). However, Schwaner & Kroer (2001) measured the activity of the bacterial populations and were not able to establish a correlation between transfer and activity, suggesting that activity was not limiting for transfer. We did not measure the activity of the bacteria, but because the donor density in the pea rhizospheres was from seven to 60 times higher than in the barley rhizospheres (Fig. 1), density rather than activity probably was responsible for the elevated transfer in pea. The question to be asked is what caused the difference in density.

Bacterial density and distribution pattern in the rhizosphere

To investigate the differences in donor density in the pea and barley rhizospheres, we set up a series of experiments to determine the colonization patterns of the two plants by the donor. The root print procedure, in which the rhizoplane bacteria were replica-plated onto agar plates, was used to visualize on a large scale which parts of the roots were colonized.

In the pea–vermiculite rhizosphere, the donor established a population of cells along the whole length of the root (Fig. 3). In the barley rhizosphere, on the other hand, only the spermosphere and upper parts of the root were colonized (Fig. 3). The recipient bacteria were present ubiquitously along the roots of both plant species (not shown). The distribution of the transconjugants strictly followed the donor colonization pattern, as transconjugants were observed along the whole length of the pea roots while they were found only at the root base of barley (Fig. 3).

3

Distribution of donors (green) and transconjugants (blue) in the rhizosphere of barley and pea grown in vermiculite. At day 0, both an inoculated (green colonies) and a noninoculated (control) seed were placed on the agar dish. Note that on day 6 only the upper parts of the barley roots are shown. The data shown represent one out of a total of three independent experiments, all of which gave similar results.

In soil rhizospheres, the donor colonization patterns were similar to those in vermiculite; however, transconjugant colonies were only observed at the root base (not shown). In barley, this distribution of transconjugants was expected based on the donor colonization. The distribution of transconjugants in the pea rhizosphere, however, was surprising. The average donor densities in the pea rhizospheres in vermiculite and soil during the first 6 days of incubation were not significantly different (_P_>0.05) (Fig. 1). The recipient concentration, on the other hand, was c. 10 time slower in soil than in vermiculite (Fig. 1), indicating that colonization of the rhizosphere by the recipient was negatively affected by the indigenous soil bacteria. This relatively poor recipient colonization in soil was probably the reason why identical transconjugant distributions were not observed in the vermiculite and soil rhizospheres of pea.

To analyse differences in bacterial density and distribution at the microscale level, we took advantage of the fluorescent reporter genes inserted into the plasmid and chromosome of the recipient, which enabled microscopic discrimination between donor, recipient and transconjugant cells at the single-cell level in a noninvasive way.

Pea roots were generally more densely colonized by the donor than barley roots (Fig. 4), and dense layers of donor bacteria were often found at the root base. On barley roots, cells were found in scattered mono-layered microcolonies. Recipient cells were the dominant colonizer of barley roots, while donor and recipient microcolonies were more equally distributed on the pea root. Motile bacteria were frequently seen on pea roots, whereas they were rare on barley roots.

4

Micrographs of hot-spots on the root surface of 3-day-old pea (a) and barley (b and c) seedlings colonized with green fluorescent donor cells, red fluorescent recipient cells, and orange/yellow fluorescent transconjugant cells. Micrograph (c) is a magnification of the lower-left microcolony in (b). The bar in (a) represents 5 μm, while the bar in (b) represents 10 μm.

Transconjugants were found mixed among donor and recipient cells (Fig. 4). In pea, transconjugants were most often observed in the outer layer of the bacterial biofilm, suggesting that proliferation of the donors happened before conjugation took place. In transconjugant-containing microcolonies, no apparent differences in T/D between microcolonies were observed, indicating that the ‘intrinsic’ transfer efficiency between donors and recipients was the same in both rhizospheres.

In conclusion, both the large-scale and microscale observations suggested that the high transfer frequency in the pea rhizosphere was a result of the combined effects of the high density and uniform distribution of the donor, which in turn facilitated high numbers of donor-to-recipient encounters.

Production and utilization of root exudates

Root exudation per seedling was higher for pea than for barley. The average amount of water-soluble exudates, extracted from both seeds and roots after 6 days of incubation, were 2.71±0.40 mgC plant−1 day−1 for barley and 4.70±0.86 mgC plant−1 day−1 for pea; that is, exudation was almost twofold higher in pea. Since the dry weight of the roots was approximately the same for the two plant species, but the dry weight of the pea seed was several times higher than that of barley (data not shown), the difference in exudation rate was probably the result of differences in the exudation from the seeds.

During seed germination, nutrients are released and bacterial populations concentrate around the emerging embryo (Tombolini et al., 1999). Our experimental method did not allow us to quantitate the relative importance of seed and root exudates. However, since dense layers of donor bacteria were often found at the root base of pea, exudation from the large-size pea seeds probably accounted for a substantial fraction of the measured exudates.

No qualitative difference between pea and barley root exudates with respect to supporting growth was observed. Growth of the donor on undiluted exudates showed no lag phase (Fig. 5a), and the maximal biomass yield was linearly correlated (_r_=0.995) with the total organic carbon content of the exudates (Fig. 5b). Thus, differences in the amount, but not in the quality, of the exudates appeared to be responsible for the relatively high donor densities in the pea rhizosphere.

5

Root exudates as growth substrate for Pseudomonas putida LM24/pKJK5::gfp. (a) Growth on barley exudate, pea exudate and germination buffer harvested after 6 days. (b) Correlation between the concentration of root exudate and the maximal biomass yield. Germination buffer (△); barley (◻) and pea (○) exudates harvested after 3 days; barley (▪) and pea (●) exudates harvested after 6 days.

Mechanism of dispersal in the pea rhizosphere

The fact that a higher exudation rate in the pea rhizosphere allowed a higher donor biomass to be achieved compared with barley did not explain the mechanism by which the donor was spread longitudinally on the pea root. To distinguish dispersal resulting from continuous colonization of the elongating root by cell divisions from dispersal by passive transport/motility, an experiment was set up in which the donor was prevented from multiplying by addition of the bacteriostatic antibiotic Nal to the vermiculite.

Passive transport or motility rather than growth appeared to be the primary mechanism(s) of donor spreading, as addition of Nal had no effect on the ability of the donor to colonize the pea rhizoplane (Fig. 6). Obviously, if the added Nal sorbed to the vermiculite, the free concentration would have been reduced and the effect of Nal potentially eliminated. We did not measure the free concentration of Nal in the microcosms, but growth of the seedlings was inhibited at 100 mg L−1 Nal (Fig. 6), suggesting that sorption to the vermiculite was insignificant. Toxicity of Nal towards the donor strain could potentially also have affected the experimental results. However, a control experiment demonstrated that Nal concentrations up to 500 mg L−1 (i.e. five times the maximal concentration in the microcosms) had no effect on the survival (culturability) of the donor cells in batch cultures (not shown).

6

Distribution of donors (green colonies) on the rhizoplane of pea grown in vermiculite amended with 0 mg, 50 mg and 100 mg Nal. Triplicate roots at each Nal concentration are shown.

Colonization of the entire pea root by P. fluorescens has previously been reported (Dandurand et al., 1997), and passive movement with flowing water rather than motility was proposed as the major mechanism of spreading (Bowers & Parke, 1993). In our case, passive transport by water movement was unlikely because microcosms were not watered during the incubation. Furthermore, had water movement been important, a more complete colonization of the barley rhizosphere would have been expected. Therefore, plant-specific differences between pea and barley probably were responsible for the different colonization patterns.

Among the factors generally believed to be important for root colonization are nutrient availability and features of the root surface (Dandurand et al., 1997). In pea, the higher exudate production resulted in a higher donor biomass relative to barley, and, as mentioned above, dense donor populations were observed at the root base. Root surface features, on the other hand, did not appear to play an important role. Rather, the growth rate of the roots impacted the colonization. Measurements of root growth demonstrated that the barley roots grew almost 10 times faster than the pea roots. The growth rate of the barley root was 4.8 mm h−1 (calculated on the basis of the total length of the branched roots), while the pea root grew at a rate of 0.5 mm h−1. Thus, the ability of the donor to colonize the entire pea root appeared to be the combined result of relatively dense populations at the root base, which acted as a reservoir for colonization, and a relatively slow growth of the root.

Conclusion

Our observations confirmed that the pea rhizosphere is several times more conducive to conjugal plasmid transfer than the barley rhizosphere. We also showed that, compared with barley, a relatively high production of exudates by the pea seed/root resulted in a higher density of donor cells in the pea rhizosphere. Furthermore, it was demonstrated that the pea rhizosphere was uniformly colonized by the donor, while only the upper part of the barley root was colonized. Finally, spreading to the deeper parts of the pea root was probably the result of passive transport and/or motility.

In our experimental set up, in which the donor was coated onto the seeds, while the recipient was inoculated into the plant growth medium, a likely scenario explaining the high plasmid transfer ratio in the pea rhizosphere is that a high exudation rate during seed germination allowed the establishment of a dense donor population. The relatively slow growth of the root enabled this population to act as a reservoir for colonization (either passively or resulting from motility) of the growing root. The dense and uniform distribution of the donor cells resulted in a high frequency of donor–recipient encounters, and, hence, a high ratio of transfer. In the barley rhizosphere, on the other hand, the seed exuded at a relatively slow rate, while the root elongated rapidly. Therefore, scattered donor populations mainly on the upper parts of the roots resulted in relatively few donor–recipient encounters. Thus by affecting the cell density and distribution, exudation and root growth rate appear to be the key parameters controlling horizontal plasmid transfer in the rhizosphere.

Acknowledgements

We thank Heidi Irming and Lilly Mathiesen for their technical assistance. Bjarne Munk Hansen is thanked for his advice during the study. L.M. was supported by a grant from the Danish Directorate for Food, Fisheries and Agri Business (grant no. 3304-03-4).

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Author notes

Editor: Kornelia Smalla

© 2006 Federation of European Microbiological Societies. Published by Blackwell Publishing Ltd. All rights reserved