Skip to main page content

• HOME
• Current Issue
• Archive
• Alerts
• About ASM
• Contact us
• Tech Support
• Journals.ASM.org

This Article Abstract Full Text (PDF) Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Reprints and Permissions Copyright Information Books from ASM Press MicrobeWorld Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Pérez-Mendoza, D. Articles by Sanjuán, J. Search for Related Content PubMed PubMed Citation Articles by Pérez-Mendoza, D. Articles by Sanjuán, J.

 Previous Article  |  Next Article 

Journal of Bacteriology, September 2004, p. 5753-5761, Vol. 186, No. 17
0021-9193/04/$08.00+0     DOI: 10.1128/JB.186.17.5753-5761.2004
Copyright © 2004, American Society for Microbiology. All Rights Reserved.

Identification of Functional mob Regions in Rhizobium etli: Evidence for Self-Transmissibility of the Symbiotic Plasmid pRetCFN42d

Daniel Pérez-Mendoza,¹ Ana Domínguez-Ferreras,¹ Socorro Muñoz,¹ María José Soto,¹ José Olivares,¹ Susana Brom,² Lourdes Girard,² José A. Herrera-Cervera,^1, and Juan Sanjuán^1*

Departamento Microbiología del Suelo y Sistemas Simbióticos. Estación Experimental del Zaidín, Consejo Superior de Investigaciones Científicas, Granada, Spain,¹ Programa de Genética Molecular de Plásmidos Bacterianos, Centro de Investigación sobre Fijación de Nitrógeno, UNAM, Cuernavaca, Morelos, México²

Received 29 March 2004/ Accepted 2 June 2004

Top Abstract Introduction Materials and Methods Results Discussion References

 
An approach originally designed to identify functional origins of conjugative transfer (oriT or mob) in a bacterial genome (J. A. Herrera-Cervera, J. M. Sanjuán-Pinilla, J. Olivares, and J. Sanjuán, J. Bacteriol. 180:4583-4590, 1998) was modified to improve its reliability and prevent selection of undesired false mob clones. By following this modified approach, we were able to identify two functional mob regions in the genome of Rhizobium etli CFN42. One corresponds to the recently characterized transfer region of the nonsymbiotic, self-transmissible plasmid pRetCFN42a (C. Tun-Garrido, P. Bustos, V. González, and S. Brom, J. Bacteriol. 185:1681-1692, 2003), whereas the second mob region belongs to the symbiotic plasmid pRetCFN42d. The new transfer region identified contains a putative oriT and a typical conjugative (tra) gene cluster organization. Although pRetCFN42d had not previously been shown to be self-transmissible, mobilization of cosmids containing this tra region required the presence of a wild-type pRetCFN42d in the donor cell; the presence of multiple copies of this mob region in CFN42 also promoted conjugal transfer of the Sym plasmid pRetCFN42d. The overexpression of a small open reading frame, named yp028, located downstream of the putative relaxase gene traA, appeared to be responsible for promoting the conjugal transfer of the R. etli pSym under laboratory conditions. This yp028-dependent conjugal transfer required a wild-type pRetCFN42d traA gene. Our results suggest for the first time that the R. etli symbiotic plasmid is self-transmissible and that its transfer is subject to regulation. In wild-type CFN42, pRetCFN42d tra gene expression appears to be insufficient to promote plasmid transfer under standard laboratory conditions; gene yp028 may play some role in the activation of conjugal transfer in response to as-yet-unknown environmental conditions.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Bacteria grouped within the Rhizobiaceae, Phyllobacteriaceae, and Bradyrhizobiaceae families, collectively known as rhizobia, are able to establish nitrogen-fixing symbiosis with leguminous plants. Many of these organisms contain complex genomes, with one chromosome and one or more large plasmids ranging in size from ca. 100 kb to >2 Mb. A common feature of the genomes of the rhizobia is that genes involved in the symbiotic process are located on independent replicons known as symbiotic plasmids (pSym) or in "symbiotic islands" within the chromosome. In addition to these symbiotic elements, rhizobia may carry additional plasmids, namely, nonsymbiotic or cryptic plasmids, that are not indispensable for symbiosis or simply with no specific function assigned (29, 38, 39, 45).

Rhizobia are difficult to isolate directly from the soil or rhizosphere; they are often isolated by virtue of their ability to nodulate specific legumes, although the presence of large numbers of nonsymbiotic rhizobia in soils is well recognized (32, 47, 52). Acquiring the ability to nodulate leguminous plants provides rhizobia with the capacity to exploit a very exclusive ecological niche and, therefore, some important advantages over a strictly saprophytic lifestyle. Thus, it seems reasonable to think that the gain of the genetic information necessary to nodulate a specific host should be a very important event in the evolution of these soil bacteria.

There is abundant evidence for symbiotic gene transfer among different species or genera of rhizobia. Much of these data come from the analysis of soil populations and the finding that different bacterial species carry similar plasmids (10) or, vice versa, that different plasmids may associate with similar chromosomes (55). After introduction of inoculant strains in soils where no native symbiotic rhizobia are present, sometimes new rhizobial populations arise as a consequence of symbiotic gene transfer from the inoculant to nonsymbiotic rhizobia in the soil (51). However, direct experimental data are required to understand the dynamics of rhizobial DNA exchange. Furthermore, conjugal transfer of pSyms under laboratory conditions appears to occur at negligible frequencies or is undetectable, which raises questions about the actual capacity of these elements for lateral spread. Indeed, few rhizobial nodulation plasmids (pSyms) display consistent and high conjugation frequencies in normal laboratory media. One of these is pRL1JI from Rhizobium leguminosarum, a plasmid carrying genes for nodulation and nitrogen fixation on legumes such as pea, vetch, and lentil. pRL1JI conjugal transfer is regulated by quorum-sensing N-acyl-homoserine lactones (AHLs) through a novel regulatory relay (12). It seems likely that similar regulatory mechanisms operate in other rhizobial nonsymbiotic plasmids, such as pRme41a from Sinorhizobium meliloti (36) or pRetCFN42a from R. etli (53). In other cases, such as the Sym plasmid of Rhizobium sp. strain NGR234, tra gene expression may be regulated by AHLs, but conjugal transfer is negligible under laboratory conditions and not affected by quorum sensing AHLs, suggesting that additional upstream regulatory cascades control tra gene expression (21). Such a hypothetical cascade would perhaps be analogous to the opine response controlling conjugal transfer of Agrobacterium tumefaciens Ti plasmids (17, 26, 35).

In other cases, as for the R. etli symbiotic plasmid pRetCFN42d, the ability to cointegrate with a resident conjugative plasmid, pRetCFN42a, provides an alternative means for lateral spread (9, 53).

For most symbiotic plasmids or islands, it is clear that significant conjugal transfer does not occur under laboratory conditions; frequencies of transfer range from very low to undetectable (2, 21, 27, 41, 50), which suggests that either their transfer is precisely controlled or that these elements have lost the capacity for efficient lateral spread. However, genome sequencing is revealing that many of these elements do carry genes potentially involved in conjugal transfer (18, 19, 30, 31). Therefore, it seems necessary to investigate whether pSym plasmids that do not transfer at appreciable rates in the laboratory are actually proficient for self-transmissibility under natural conditions. For this purpose, approaches such as that reported by Turner et al. (54) do not provide an optimal solution, since the presence of tra genes in a particular replicon do not guarantee its conjugation proficiency, as indicated above. We have previously reported an approach for the identification of functional mob regions in S. meliloti (24). The use of merodiploid donor populations allowed the identification of clones carrying functional oriTs under the experimental conditions. This approach allowed us to characterize the mob region of plasmid pRmeGR4a, a cryptic, self-conjugative plasmid of S. meliloti GR4. Indeed, 10 other putative oriTs were identified. Here we report on the disadvantages of this original approach, where a recA-independent recombination event during the construction of merodiploid donor populations resulted in the downstream selection of false Mob⁺ clones. The approach has been modified to prevent such disadvantages, and its efficacy was tested during the identification of mob regions in the R. etli CFN42 genome. This has allowed us to obtain the first evidence that the Sym plasmid of this strain may be self-transmissible and that its transfer is subject to regulation.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Bacterial strains and plasmids. All bacteria and plasmids used in the present study are listed in Table 1. R. etli and S. meliloti strains were grown at 30°C on TY medium (tryptone-yeast extract-CaCl₂) (4). Escherichia coli and A. tumefaciens were grown on Luria-Bertani medium (44). When required, antibiotics were added at the following concentrations: nalidixic acid, 20 µg/ml; spectinomycin, 50 µg/ml for R. etli and 200 µg/ml for S. meliloti; kanamycin (Km), 50 µg/ml for R. etli and A. tumefaciens and 200 µg/ml for S. meliloti; gentamicin (Gm), 10 µg/ml; rifampin (Rif), 50 µg/ml; streptomycin (Sm), 100 µg/ml for R. etli, 200 µg/ml for S. meliloti, and 25 µg/ml for E. coli; and tetracycline (Tc), 2 µg/ml for Rhizobium and 10 µg/ml for E. coli.

View this table: [in this window] [in a new window]

TABLE 1. Bacterial strains and plasmids

Bacterial matings. Donor strains grown to an approximate optical density at 600 nm of 0.2 and recipient Rhizobium, Sinorhizobium, or E. coli strains grown to late exponential phase were washed and mixed in a 1:1 donor/recipient ratio. Mating mixtures were resuspended in 50 µl of TY medium and loaded onto a sterile nitrocellulose filter of 0.45-µm pore size. Filter mating mixtures were deposited on TY-agar plates and incubated overnight at 30°C. Cells were resuspended by vigorous vortexing and diluted in liquid medium. To calculate transfer frequencies, donor, recipient, and transconjugant CFU were counted after mating disruption and plating of serial dilutions. Transconjugants were selected on plates supplemented with appropriate antibiotics. The transfer frequency was expressed as the number of transconjugants per output recipient. Donor and recipient spontaneous resistance to selective antibiotics was also determined.

Plasmid profiles. Plasmids profiles were visualized by the Eckhardt procedure (14) as modified by Hynes and McGregor (28).

DNA hybridizations. For DNA hybridization, total genomic DNAs of S. meliloti or R. etli strains were isolated by standard procedures (37) and digested with endonuclease EcoRI, electrophoresed on 0.8% agarose gels, and transferred to positively charged nylon membranes by the method of Southern (44). Blots of intact plasmid profiles electrophoresed in 0.8% agarose-1% sodium dodecyl sulfate-Eckhardt gels were transferred to positively charged nylon membranes as well (44). DNA hybridization probes were digoxigenin labeled according to manufacturer instructions (Roche, Barcelona, Spain). Hybridization and membrane washes were carried out under high-stringency conditions. Membranes were prepared for chemiluminiscent detection (Roche) and exposed to Kodak X-Omat film (Sigma).

PCR, cloning, and sequencing. Total DNA was prepared from mid-exponential-phase R. etli cells. PCR primers Yp028F and Yp028R were designed to match conserved regions of yp028 gene of pRetCFN42d of R. etli (NC_004041). The primer sequences of Yp028F (GGATCCTCCATCACGTTGAGCAGC) and Yp028R (GGATCCGGCATCAACCTCTGAGAC) correspond to positions 140473 to 140490 and 141118 to 141135, respectively, of the R. etli pRetCFN42d replicon sequence (19), with BamHI restriction sites at the 5' end of the primers (underlined) to facilitate subsequent clonings. These primers were used at a final concentration of 50 pmol in 50-µl amplification reactions containing 1x PCR buffer, 200 µM deoxynucleoside triphosphates, 1.5 mM MgCl₂, and 1 U of Taq polymerase (Sigma). The PCR profile used was as follows: an initial denaturation of 94°C for 10 min, followed by 30 cycles of 94°C for 60 s, 52°C for 60 s, and 72°C for 60 s, followed in turn by a final extension of 72°C for 10 min. The PCR product was checked and digested according to standard procedures (44). For all cloning procedures, standard DNA techniques were used as described previously by Sambrook et al. (44). Several EcoRI fragments from cosmid pRe182R1a were cloned into pBluescript (48). Sequencing was carried out with a Perkin-Elmer ABI Prism 373 automated sequencer. DNA sequence edition, translation, and analysis were performed by using the Vector NTI 5.0 software package and the program BLAST from the NCBI network service (1).

Construction of a traA mutant derivative of pRetCFN42d. Two primers, RetraAF (TCGTTGGTGTGGGCGAGCA) and RetraAR (GCGCAGCCGCCGATGCTCA), were used to amplify a 2,489-bp fragment from the R. etli CE3 pSym (from positions 143900 to 146388 of accession number 004041 [19]). The PCR product was cloned into pGEM-T Easy cloning vector; a 673-bp EcoRV fragment from traA was removed and replaced by a Gm resistance (Gm^r) gene cassette from SmaI-digested pMS255 (3). The construction was cloned as an EcoRI fragment into plasmid pK18mobSacB (46) and introduced by conjugation into strain CFNX195 of R. etli. Allele replacement was selected as described previously (46), and traA mutants were verified after hybridization with a labeled plasmid pK18traAGm BamHI digested as a probe.

Cell transformations. Bacterial transformation was carried out by electroporation by using an electro-cell manipulator apparatus (BTX 600; BTX, San Diego, Calif.). Electrocompetent cells were prepared according to the instructions of the manufacturer and stored at –80°C. For electroporation, cells were thawed on ice and mixed with plasmid DNA (0.3 to 0.5 µg/ml of cell suspension) and then transferred to a 0.2-cm electrode gap chilled cuvette. A field strength of 2.5 kV/cm, a 6.8-ms pulse length, and a 129- set resistance was applied; cells were then immediately suspended in 1 ml of TY or Luria-Bertani medium and then incubated at 30°C for 15 h or 37°C for 1 h for R. etli or E. coli, respectively. Appropriate dilutions were plated on selective media.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Analysis of putative mob regions previously identified in S. meliloti. Herrera-Cervera et al. (24) reported an approach to identify DNA regions with the ability to convert a nontransmissible vector into a mobilizable plasmid and thus to identify functional origins of conjugative transfer (oriT and mob). RecA^– S. meliloti merodiploid populations were obtained by transferring an S. meliloti gene library from E. coli into S. meliloti by triparental matings using pRK2013 as a helper plasmid (16). The S. meliloti merodiploids were used as donors en masse in matings with S. meliloti recipient strains and transconjugants carrying vector-encoded antibiotic resistance selected. Eleven putative mob regions were identified, most of which originated from plasmid replicons. This approach allowed identification and further characterization of the oriT from the conjugative plasmid pRmeGR4a (24), as well as the mob region from pRmeGR4b cloned in cosmid pRmOR65, a nonsymbiotic plasmid that can be mobilized in trans by pRmeGR4a (22).

During the analysis of the remaining nine putative mob regions, we found that all of these nine cosmids showed unusually high transfer frequencies from either S. meliloti (24) or E. coli donors and therefore behaved as self-transmissible Mob⁺ Tra⁺ plasmids. In contrast, cosmids pRmOR69 (oriT from pRmeGR4a) and pRmOR65 (oriT from pRmeGR4b) displayed a Mob⁺ Tra^– phenotype. After removing all of the EcoRI insert from the nine Mob⁺ cosmids to theoretically obtain the empty vector pLAFR1, we observed that the nine empty vectors (form here on named pLAFR1*) still maintained the Mob⁺ Tra⁺ character, in contrast to the Mob⁺ Tra^– properties of the original vector pLAFR1. This suggested that the Mob⁺ Tra⁺ capacity of the nine cosmids was due to a change in the cosmid vector pLAFR1. After digestion with Eco47III the restriction profiles of the pLAFR1* molecules were compared to those of vector pLAFR1 and plasmid pRK2013. Whereas the empty vectors derived from mob cosmids pRmOR69 and pRmOR65 had restriction profiles identical to the original pLAFR1 vector, the remaining nine pLAFR1* molecules with Mob⁺ Tra⁺ phenotype showed restriction patterns that appeared to be chimeras derived from both pLAFR1 and the mobilizing plasmid pRK2013 (data not shown). However, the pLAFR1* cosmids carried no resistance to Km, as does pRK2013. In conclusion, 9 of the 11 mob cosmids isolated by Herrera-Cervera et al. (24) did not contain any S. meliloti oriT. These cosmids probably were the result of recombination events between the pLAFR1 vector and the mobilizing plasmid pRK2013 during construction of the S. meliloti merodiploid populations. However, the recombination between the two plasmids must have been a recA-independent phenomenon, since all of the strains used by Herrera-Cervera et al. were RecA defective (24).

A modified approach to identify Mob⁺ regions in rhizobial genomes. The above results indicated that, although the approach reported by Herrera-Cervera et al. might be useful for identifying rhizobial functional oriTs, a modification was needed in order to prevent or at least reduce the excessive selection of false Mob⁺ clones. As a general rule, we recommend against using the suicide helper plasmid pRK2013 when the transfer of Mob⁺ plasmids from E. coli to rhizobial strains is needed. Alternatively, this can be done by using mobilizing E. coli strains (i.e., S17-1 [49]) instead of mobilizing vectors or, when feasible, by direct introduction of cosmids or plasmids into the rhizobial strains by electroporation. We have tested both alternatives as a modification to the oriT cloning strategy of Herrera-Cervera et al. (24).

The cosmids from the S. meliloti GR4 gene library were isolated from pooled E. coli HB101 clones by standard methods and electroporated into the mobilizing strain S17-1. The resulting clones were pooled and used as donors en masse in matings with the S. meliloti strains GR4KLR and GRM10KR (24), yielding merodiploid populations derived from both rhizobial strains. From here, we repeated the experiments of Herrera-Cervera et al. (24). When we used the new GRM10KR merodiploid population as the donor in matings, no transconjugants were obtained, in contrast to the 10^–5 transconjugants per recipient obtained by Herrera-Cervera et al. (24). However, when the GR4KLR merodiploid population was the donor, transconjugants arose at a frequency of 10^–7, a finding similar to that obtained by Herrera-Cervera et al. (24). The cosmids acquired by 24 of the transconjugants were studied by restriction analysis, and all of them were found to correspond to cosmid pRmOR69, carrying the oriT of plasmid pRmeGR4a (24). Thus, the modified approach resulted in the selection of the mob region of pRmeGR4a plasmid without the appearance of any false Mob⁺ clones. We were unable to isolate additional putative oriT-containing regions from S. meliloti GR4. The fact that we could not isolate the mob region from plasmid pRmeGR4b in this experiment may be due to the fact that mobilization of this plasmid by pRmeGR4a is at least 10-fold less efficient than transfer of pRmeGR4a itself.

We also applied the modified approach to the isolation of mob regions from R. etli CE3. This strain carries six plasmids, ranging from 180 to 630 kb in size (9). To date, only the cryptic plasmid pRetCFN42a (abbreviated as p42a) has been shown to be self-transmissible (9, 53). Transfer of pRetCFN42d (the pSym; abbreviated as p42d) has also been detected (8), but this was shown to be fully dependent on the presence of p42a. The mechanism for pSym transfer appeared to require its cointegration with p42a.

Merodiploid populations of R. etli strains CE3 and CFNX182, a CE3 derivative cured of plasmid p42a (see Table 1), were obtained after introducing by electroporation cosmids from a R. etli CE3 gene library made in vector pLAFR1 (25) into each strain. Each merodiploid population was used as donor en masse in matings with R. etli CFNX218Rif or E. coli HB101 as recipients.

Using the CFNX182 merodiploids as donors, HB101 Tc^r transconjugants arose at frequency of 1.6 x 10^–4 (Table 2). Cosmids from 20 of such transconjugants were isolated by standard procedures and subjected to restriction analysis with endonuclease EcoRI. All putative mob cosmids showed similar EcoRI patterns (data not shown), with at least five EcoRI fragments in common, indicating that all of them contained overlapping DNA inserts. Furthermore, one of these cosmids showed a restriction profile identical to cosmid pC-13, which was recently characterized by Tun-Garrido et al. (53), which contains the oriT and all transfer functions from plasmid p42a. To maintain the nomenclature used by Tun-Garrido et al. (53), we named the two different overlapping cosmids pC13a and pC13b, respectively. The selection of the p42a mob region demonstrated that our approach could also be applied to R. etli. When R. etli CFNX218Rif was used as a recipient in matings with the CFNX182 merodipoids, CFNX218Rif Tc^r transconjugants arose at frequency of 3 x 10^–5 (Table 2). Cosmids from 32 such transconjugants were isolated and subjected to restriction analysis with EcoRI. Cosmids from 27 transconjugants showed similar EcoRI patterns to any one of members of the pC-13 cosmid family described above (with the appearance of a new member of this family, pC13c), whereas the remaining five cosmids were of three overlapping types (which we designated pRe182R1b, pRe182R1c, and pRe182R1d), with restriction patterns completely different to the pC-13 family, suggesting that they could represent a new mob region.

View this table: [in this window] [in a new window]

TABLE 2. Isolation of Mob+ cosmids from a R. etli cosmid librarya

Using the CE3-derived merodiploid population as donor, Tc^r transconjugants arose at frequencies of 7.68 x 10^–6 and 1.17 x 10^–4 when HB101 or R. etli CFNX218 Rif^r strains were used as recipients, respectively. Cosmids from the transconjugants obtained from each mating were analyzed and identified as members of either of the two cosmid families described above (Table 2). A new member of the pRe182R1 cosmid family, pRe182R1a, was identified.

In summary, our modified approach for identifying oriT regions appeared to work correctly in R. etli, since it had been able to select for the only previously known functional mob region in strain CE3, located in plasmid p42a. In addition, we were able to isolate a new functional mob region not reported previously.

Characterization of a mob region in the R. etli CFN42 symbiotic plasmid. The putative mob region present in the cosmid family pRe182R1 was found to belong to the symbiotic plasmid p42d after cosmid pRe182R1a was used as a probe in hybridizations either against blotted digested genomic DNAs from various R. etli strains or against blotted Eckhardt-type gels containing intact plasmids from these strains. Specific hybridization signals were absent only in strains cured of p42d (Fig. 1).

View larger version (63K): [in this window] [in a new window]

FIG. 1. Replicon localization of the Mob+ DNA identified in cosmid pRe182R1a. Blots of EcoRI-digested genomic DNAs (A) and of intact plasmid profiles (Eckhardt gels) (B) hybridized against digoxigenin-labeled pRe182R1a as a probe are shown. Lanes: M, digoxigenin-labeled DNA molecular weight marker; 1, R. etli CE3 (wild type); 2, R. etli CFNX182(p42a–); 3, R. etli CFNX183(p42b–); 4, R. etli CFNX184(p42c–); 5, R. etli CFNX89(p42d–); 6 R. etli CFNX185(p42e); 7, R. etli CFNX186(p42f–); 8, R. etli CFNX218(p42a–, p42b–, p42c–, p42d–, p42e, p42f–).

To confirm that the isolated cosmids of the pRe182R1 family could indeed be mobilized from R. etli, individual cosmids were introduced back into CFNX182, and the corresponding strains were separately used in matings with CFNX218Rif as recipient. All of the overlapping cosmids showed similar transfer frequencies of 10^–4 transconjugants per recipient. To test the influence of p42d background on the mobilization of this cosmid family, pRe182R1a was introduced into strain CFNX2001 (cured of p42a and p42d), and the resulting strain was crossed with CFNX218Rif and HB101 as recipients. No transconjugants were obtained from these matings, indicating that mobilization of these cosmids required the presence of p42d in the donor cell. To further characterize this oriT, we cloned and end sequenced several EcoRI fragments from cosmid pRe182R1a. Comparison with the genome sequence of plasmid p42d (19) demonstrated that we had isolated a putative mob region containing two divergently transcribed tra operons and an oriT located in this symbiotic plasmid (Fig. 2A [19]). As described above, our data suggested that this cloned mob region was functional and that its mobilization required plasmid p42d, despite the fact that this plasmid has never been shown to be self-transmissible. This apparent paradox could be explained if both the intact plasmid and the cloned mob region (in multicopy) were necessary for the plasmid to show self-transfer in laboratory media. If so, then the cloned mob region would promote transfer of the entire pSym. Indeed, strain CFNX667 (RecA^–) carrying cosmid pRe182R1a was able to donate the pSym p42d to either R. etli CFNX218Rif (1.38 x 10^–3 transconjugants per recipient) or A. tumefaciens GMI9023 (frequency of 2.56 x 10^–5) in media where the transfer of p42d is otherwise undetectable. This suggested that multiple copies of the p42d mob region allowed conjugation of the symbiotic plasmid to occur at detectable rates in standard laboratory conditions. Furthermore, a derivative of cosmid pRe182R1a carrying an 11-kb HindIII deletion that removed this mob region (plasmid pReOR182RaHindIII) also lost the ability to promote transfer of p42d.

View larger version (20K): [in this window] [in a new window]

FIG. 2. (A) Physical map of the pRe182R1 cosmid family containing the mob region of the R. etli symbiotic plasmid p42d. Arrows indicate the transcription directions of the identified genes according to Gonzalez et al. (19). The Dtr (DNA transfer and replication) genes are shown as black arrows. The gene coding for the hypothetical coupling protein TraG gene is shown by a striped arrow. The DNA insert in each member of the cosmid family is indicated by dotted bars. oriT, origin of transfer; E, EcoRI; H, HindIII. (B) Identification of an ORF promoting conjugal transfer of the p42d symbiotic plasmid. Various constructs were derived from the 7.7-kb HindIII fragment cloned in pJBdp1 and tested for pSym transfer from strain CFNX667. "(+)" or "(–)" indicates promotion or no promotion, respectively, of p42d conjugal transfer. In pTEYp028 and pTEYp028R, the dotted box indicates the trp promoter, and the white arrow indicates the yp028 ORF. E, EcoRI; H, HindIII; B, BamHI.

Overexpression of the yp028 gene promotes R. etli pSym transfer in standard media. Since all overlapping cosmids pRe182R1a to pRe182R1d displayed the same capacity to promote p42d conjugation, it seemed obvious that the gene or genes responsible for this effect would be located in a DNA fragment common to all of them. This would be a DNA fragment flanked by yp026 and traD (see Fig. 2A). Indeed, a pJB3 derivative including a 7.7-kb HindIII fragment carrying the mob region (pJBdp1) had the same effect as the entire cosmids in promoting p42d transfer (Fig. 2B and Table 3). Several fragments were subsequently subcloned into pJB3Tc19 and tested for promotion of p42d conjugal transfer. As outlined in Fig. 2B and Table 3, clone pJBdp2, containing an intact yp028 open reading frame (ORF), maintained the ability to promote conjugation of p42d, but this property was lost when a BglII deletion removed the C-terminal half yp028 (plasmid pJBdp3). The results indicated that it was the presence of this ORF yp028 that was responsible for promoting conjugation of p42d. Since this effect was only observed when this gene was cloned in multicopy vectors (p42d contains an intact copy of yp028), we reasoned that a change in yp028 expression resulted in the observed effects on p42d conjugal transfer. Indeed, when the yp028 coding sequence was placed under the control of the trp promoter (plasmid pTEYp028), it had the effect of promoting conjugal transfer of p42d at high frequencies (Fig. 2B and Table 3). In contrast, plasmid pTEYp028R containing the same yp028 cloned in the opposite orientation to the trp promoter (thus containing a promoterless yp028) was unable to promote conjugal transfer of the p42d Sym plasmid (Fig. 2B; Table 3). These results strongly suggested that both the presence of an intact yp028 and expression of the ORF were necessary to promote conjugal transfer of the R. etli pSym. Transconjugants were verified to carry the p42d plasmid after the plasmid profiles were visualized (Fig. 3). We observed that most of the transconjugants had acquired only plasmid p42d from CFNX195(pTEyp028) (Fig. 3, lane 3), whereas in some cases both p42d and p42b were transferred (Fig. 3, lane 2). Cointegration of p42d and p42b has previously been observed (7). As a result, the transconjugants carrying these two plasmids may have been generated through the transfer of a p42b-p42d cointegrate and its subsequent resolution into the wild-type plasmids in the recipient.

View this table: [in this window] [in a new window]

TABLE 3. Identification of an ORF promoting conjugal transfer of the R. etli symbiotic plasmid p42d

View larger version (55K): [in this window] [in a new window]

FIG. 3. Eckhardt-type gel electrophoresis showing plasmid profiles of transconjugants that had acquired the Sym plasmid p42d from CFNX195(pTEYp028) using CFNX218Rif(pe) as recipient. Lanes: 1, CFNX195(pTEYp028); 2, type I transconjugant; 3, type II transconjugant; 4, recipient strain CFNX218Rif. The bands corresponding to plasmids p42d and p42b are indicated.

yp028 is located downstream of traA and putatively encodes a protein of 171 amino acids that shows no sequence homology to any protein sequence of known function described in the databases. Yp028 displayed some sequence conservation (<30% sequence identity and 47% sequence similarity) with two small ORFs of unknown function, SMa0974 (1235562) from S. meliloti pSymA and Atu5116 (1136889), from the A. tumefaciens pAtC58 cryptic plasmid.

yp028-dependent transfer of p42d requires traA. To determine whether the effect of overexpressing yp028 on the promotion of conjugal transfer of p42d was indeed dependent on p42d conjugal transfer genes, we investigated the effect of a traA mutation on p42d conjugal transfer. A p42d-traA deletion mutant derived from CFNX195 was obtained as described in Materials and Methods. Plasmid pTEYp028 was introduced into the mutant strain by electroporation, and the transfer of p42d was determined in matings with A. tumefaciens GMI9023. No transconjugants were obtained (Table 3), demonstrating that the high-frequency conjugal transfer of the symbiotic plasmid p42d promoted by yp028 required a wild-type traA gene, which likely encodes the relaxase of this pSym.

Top Abstract Introduction Materials and Methods Results Discussion References

 
The origin of transfer (oriT or mob) of conjugative or mobilizable elements is the only known cis-acting function required for DNA transfer. Its presence in a plasmid or a transposon is usually suggestive of the conjugative or mobilizable capacity of such genetic elements. Among the gram-negative bacteria known as rhizobia, genome sequencing is demonstrating that most, if not all plasmids and chromosomal islands in these bacteria contain an oriT, including the symbiotic plasmid and islands that carry genes important for the establishment of nitrogen-fixing symbiosis with legumes (18, 19, 30, 31, 40). In other cases, the search for oriTs by PCR amplification of putatively conserved traC-traA intergenic regions has provided similar results in R. leguminosarum (54). Neither approach, however, is able to determine whether the mob regions identified are actually functional in the genetic background where they are found. Indeed, relatively few rhizobial Sym plasmids and symbiotic islands have been found to conjugate at high frequencies under laboratory conditions (12, 21), despite the sequence data and other evidence indicating they may have self-transmissible (Mob⁺ Tra⁺) or mobilizable (Mob⁺) capacities under natural conditions (51, 55). Therefore, reliable approaches are needed that allow the identification of Mob⁺ capabilities in these bacteria, opening the possibility to study the putative regulation of their conjugal transfer. This was the aim of the approach described by Herrera-Cervera et al. (24), which allowed the identification of 11 putative oriTs in strain GR4 of S. meliloti. Among these were the mob region from the conjugative plasmid pRmeGR4a and that of the mobilizable plasmid pRmeGR4b. However, as shown in the present study, the remaining nine putative oriT-containing clones did not represent any functional mob region. These were actually false Mob⁺ clones generated after a recA-independent recombination event between the cosmid vector pLAFR1 and the helper plasmid pRK2013 during the construction of merodiploid rhizobial populations used as donors of putative Mob⁺ clones. Such recombination led to the generation of hybrid Mob⁺ Tra⁺ plasmids (resembling pRK2013) able to replicate in S. meliloti (as does pLAFR1). The subsequent experimental selection for Mob⁺ clones converted a probably unusual event into a frequently selected one. We modified the procedure to obtain merodiploid populations, avoiding the use of helper plasmids during transfer of the rhizobial gene library from E. coli into rhizobial strains, and verified that for S. meliloti GR4 no false Mob⁺ clones were ever obtained. Similarly, the procedure was applied to identify Mob⁺ clones in a R. etli CFN42 cosmid library. In addition to showing that no artificial Mob⁺ clones were isolated in this case, we demonstrated that there are two functional mob regions in the genome of this bacterium that can be isolated by this procedure. One corresponded to the previously characterized transfer region of the cryptic, conjugative plasmid p42a (53), indicating the reliability of this approach and its application not only to S. meliloti but also to R. etli. The second was identified as a mob region in the symbiotic plasmid p42d of this strain. This was somewhat surprising since pSym p42d has never been shown to have conjugative capabilities (9, 19, 53). Although conjugal transfer of p42d has been detected previously under laboratory conditions, this transfer always relied on the cryptic plasmid p42a and required cointegration between p42d and p42a (9, 53). Furthermore, we observed that multiple copies of the mob region from p42d promoted conjugal transfer of this pSym in the absence of the cryptic plasmid p42a. Interestingly, we found that the presence of pSym was needed for mobilization of the cloned mob region. This apparent paradox was solved after identifying a small ORF, yp028, located adjacent to traA, which was responsible for this effect. Most likely, overexpression of yp028 was leading to conjugal transfer of the Sym plasmid p42d, providing the first evidence that this plasmid may be self-transmissible. The second piece of evidence comes from the fact that conjugal transfer of the R. etli pSym promoted by yp028 was dependent on the p42d traA gene. traA likely encodes the relaxase, which is essential for processing the oriT and for the initiation of DNA transfer. Thus, we have arrived at three conclusions: (i) the previously sequenced mob region of this pSym (19) is functional; (ii) conjugal transfer of this symbiotic plasmid under laboratory conditions can be promoted by overexpression of the yp028 gene; and (iii) plasmid p42d probably contains all functions needed for conjugal transfer and therefore may be regarded as a Mob⁺ Tra⁺, self-transmissible plasmid. In addition to Dtr genes, p42d carries a complete set of virB-like genes (19), similar to those identified in A. tumefaciens and involved in DNA transfer to plants or in pAtC58 plasmid conjugation (11, 33). These results support the hypothesis that the presence of transfer-related genes and particularly oriT-processing genes in rhizobial plasmids probably indicates that these elements actually have conjugative capabilities. The fact that these elements often display negligible or undetectable conjugal transfer rates in standard laboratory media does not preclude their conjugative potential under natural conditions and probably reflects a rather strict regulation of transfer functions. Based on our results, we suggest that conjugal transfer functions of the R. etli pSym may be silent under laboratory conditions and that transfer can be activated upon an environmental signal with the participation of yp028. Since overexpression of yp028 leads to pSym conjugal transfer, we speculate that activation of yp028 gene expression is a critical point during activation of conjugal transfer. The pRetCFN42d genome sequence annotation (19) includes a computational prediction that the yp028 promoter may be of the ₅₄ class, which would involve activation by a transcriptional regulator binding upstream of the promoter. Adequate experiments, however, are necessary to confirm this prediction.

At this point, we cannot speculate about the actual role of gene yp028 or whether such regulation is exerted at transcriptional or posttranscriptional levels. However, our results open a new window for investigating the actual capabilities of rhizobial symbiotic plasmids for lateral spread.

 
This study was supported by grant BIO99-0904 from MCyT to J.S. and by a CSIC-CONACYT cooperation grant to J.S. and S.B. D.P.-M. and A.D.-F. were supported by MCYT and MECD predoctoral fellowships, respectively. M.J.S. and J.A.H.-C. were supported by an MCYT postdoctoral contract.

We are grateful to G. Dávila (CIFN-UNAM [Mexico]) for sharing part of the sequence of pRetCFN42d prior to publication. Michael Hynes is acknowledged for critical comments on the manuscript.

 
* Corresponding author. Mailing address: Departamento Microbiología del Suelo y Sistemas Simbióticos, Estación Experimental del Zaidín. Profesor Albareda 1, 18008 Granada, Spain. Phone: 34958181600, ext. 219. Fax: 34958129600. E-mail: juan.sanjuan{at}eez.csic.es.

Present address: Departamento Biología Vegetal, Facultad de Ciencias, Universidad de Granada, 18071 Granada, Spain.

Top Abstract Introduction Materials and Methods Results Discussion References

 

1
1. Altschul, S. F., T. L. Madden, A. A. Schaffer, J. Zhang, Z. Zhang, W. Miller, and D. J. Lipman. 1997. Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res. 25:3389-3402.[Abstract/Free Full Text]
2
2. Banfalvi, Z., E. Kondorosi, and A. Kondorosi. 1985. Rhizobium meliloti carries 2 megaplasmids. Plasmid 13:129-138.[CrossRef][Medline]
3
3. Becker, A., M. Schmidt, W. Jäger, and A. Pühler. 1995. New gentamicin-resistance and lacZ promoter-probe cassettes suitable for insertion mutagenesis and generation of transcriptional fusions. Gene 162:37-39.[CrossRef][Medline]
4
4. Beringer, J. E. 1974. R factor transfer in Rhizobium leguminosarum. J. Gen. Microbiol. 84:188-198.[Abstract/Free Full Text]
5
5. Blatny, J. M., T. Brautaset, H. C. Winther-Larsen, K. Haugan, and S. Valla. 1997. Construction and use of a versatile set of broad-host-range cloning and expression vectors based on the RK2 replicon. Appl. Environ. Microbiol. 63:370-379.[Abstract]
6
6. Boyer, H. W., and D. Roulland-Dussoix. 1969. A complementation analysis of the restriction and modification of DNA in Escherichia coli. J. Mol. Biol. 41:459-472.[CrossRef][Medline]
7
7. Brom, S., A. García de los Santos, G. M. de Lourdes, G. Dávila, R. Palacios, and D. Romero. 1991. High-frequency rearrangements in Rhizobium leguminosarum bv. phaseoli plasmids. J. Bacteriol. 173:1344-1346.[Abstract/Free Full Text]
8
8. Brom, S., A. García de los Santos, T. Stepkowsky, M. Flores, G. Davila, D. Romero, and R. Palacios. 1992. Different plasmids of Rhizobium leguminosarum bv. phaseoli are required for optimal symbiotic performance. J. Bacteriol. 174:5183-5189.[Abstract/Free Full Text]
9
9. Brom, S., A. García-de los Santos, L. Cervantes, R. Palacios, and D. Romero. 2000. In Rhizobium etli symbiotic plasmid transfer, nodulation competitivity and cellular growth require interaction among different replicons. Plasmid 44:34-43.[CrossRef][Medline]
10
10. Brom, S., L. Girard, A. García-de los Santos, J. M. Sanjuán-Pinilla, J. Olivares, and J. Sanjuán. 2002. Conservation of plasmid-encoded traits among bean-nodulating Rhizobium species. Appl. Environ. Microbiol. 68:2555-2561.[Abstract/Free Full Text]
11
11. Chen, L., Y. Chen, D. W. Wood, and E. W. Nester. 2002. A new type IV secretion system promotes conjugal transfer in Agrobacterium tumefaciens. J. Bacteriol. 184:4838-4845.[Abstract/Free Full Text]
12
12. Danino, V. E., A. Wilkinson, A. Edwards, and J. A. Downie. 2003. Recipient-induced transfer of the symbiotic plasmid pRL1JI in Rhizobium leguminosarum bv. viciae is regulated by a quorum-sensing relay. Mol. Microbiol. 50:511-525.[CrossRef][Medline]
13
13. Ditta, G., S. Stanfield, D. Corbin, and D. R. Helinski. 1980. Broad host range DNA cloning system for gram-negative bacteria: construction of a gene bank of Rhizobium meliloti. Proc. Natl. Acad. Sci. USA 77:7347-7351.[Abstract/Free Full Text]
14
14. Eckhardt, T. 1978. A rapid method for the identification of plasmid desoxyribonucleic acid in bacteria. Plasmid 1:584-588.[CrossRef][Medline]
15
15. Egelhoff, T. T., and S. R. Long. 1985. Rhizobium meliloti nodulation genes: identification of nodDABC gene products, purification of nodA protein, and expression of nodA in Rhizobium meliloti. J. Bacteriol. 164:591-599.[Abstract/Free Full Text]
16
16. Figurski, D. H., and D. R. Helinski. 1979. Replication of an origin-containing derivative of plasmid RK2 dependent on a plasmid function provided in trans. Proc. Natl. Acad. Sci. USA 76:1648-1652.[Abstract/Free Full Text]
17
17. Fuqua, W. C., and S. C. Winans. 1994. A LuxR-LuxI type regulatory system activates Agrobacterium Ti plasmid conjugal transfer in the presence of a plant tumor metabolite. J. Bacteriol. 176:2796-2806.[Abstract/Free Full Text]
18
18. Galibert, F., T. M. Finan, S. R. Long, A. Pühler, P. Abola, F. Ampe, F. Barloy-Hubler, M. J. Barnett, A. Becker, P. Boistard, G. Bothe, M. Boutry, L. Bowser, J. Buhrmester, E. Cadieu, D. Capela, P. Chain, A. Cowie, R. W. Davis, S. Dreano, N. A. Federspiel, R. F. Fisher, S. Gloux, T. Godrie, A. Goffeau, B. Golding, J. Gouzy, M. Gurjal, I. Hernandez-Lucas, A. Hong, L. Huizar, R. W. Hyman, T. Jones, D. Kahn, M. L. Kahn, S. Kalman, D. H. Keating, E. Kiss, C. Komp, V. Lelaure, D. Masuy, C. Palm, M. C. Peck, T. M. Pohl, D. Portetelle, B. Purnelle, U. Ramsperger, R. Surzycki, P. Thebault, M. Vandenbol, F. J. Vorholter, S. Weidner, D. H. Wells, K. Wong, K. C. Yeh, and J. Batut. 2001. The composite genome of the legume symbiont Sinorhizobium meliloti. Science 293:668-672.[Abstract/Free Full Text]
19
19. González, V., P. Bustos, M. A. Ramírez-Romero, A. Medrano-Soto, H. Salgado, I. Hernández-González, J. C. Hernández-Celis, V. Quintero, G. Moreno-Hagelsieb, L. Girard, O. Rodríguez, M. Flores, M. A. Cevallos, J. Collado-Vides, D. Romero, and G. Dávila. 2003. The mosaic structure of the symbiotic plasmid of Rhizobium etli CFN42 and its relation to other symbiotic genome compartments. Genome Biol. 4:R36-R36.13.[CrossRef][Medline]
20
20. Hanahan, D. 1983. Studies on transformation of Escherichia coli with plasmids. J. Mol. Biol. 166:557-580.[Medline]
21
21. He, X., W. Chang, D. L. Pierce, L. O. Seib, J. Wagner, and C. Fuqua. 2003. Quorum sensing in Rhizobium sp. strain NGR234 regulates conjugal transfer (tra) gene expression and influences growth rate. J. Bacteriol. 185:809-822.[Abstract/Free Full Text]
22
22. Herrera-Cervera, J. A., J. Olivares, and J. Sanjuán. 1996. Ammonia inhibition of plasmid pRmeGR4a conjugal transfer between Rhizobium meliloti strains. Appl. Environ. Microbiol. 62:1145-1150.[Abstract]
23
23. Herrera-Cervera, J. A., F. I. Rodríguez-Alonso, J. Olivares, and J. Sanjuán. 1997. Evaluation of the recA-based containment system in Rhizobium meliloti GR4. FEMS Microbiol. Ecol. 22:49-56.
24
24. Herrera-Cervera, J. A., J. M. Sanjuán-Pinilla, J. Olivares, and J. Sanjuán. 1998. Cloning and identification of conjugative transfer origins in the Rhizobium meliloti genome. J. Bacteriol. 180:4583-4590.[Abstract/Free Full Text]
25
25. Huerta-Zepeda, A., L. Ortuno, P. G. Du, S. Duran, A. Lloret, H. Merchant-Larios, and J. Calderón. 1997. Isolation and characterization of Rhizobium etli mutants altered in degradation of asparagine. J. Bacteriol. 179:2068-2072.[Abstract/Free Full Text]
26
26. Hwang, I., D. M. Cook, and S. K. Farrand. 1995. A new regulatory element modulates homoserine lactone-mediated autoinduction of Ti plasmid conjugal transfer. J. Bacteriol. 177:449-458.[Abstract/Free Full Text]
27
27. Hynes, M. F., K. Brucksch, and U. Priefer. 1988. Melanin production encoded by a cryptic plasmid in a Rhizobium leguminosarum strain. Arch. Microbiol. 150:326-332.[CrossRef]
28
28. Hynes, M. F., and N. F. McGregor. 1990. Two plasmids other than the nodulation plasmid are necessary for formation of nitrogen-fixing nodules by Rhizobium leguminosarum. Mol. Microbiol. 4:567-574.[Medline]
29
29. Johnston, A. W. B., G. Hombrecher, N. J. Brewin, and M. C. Cooper. 1982. Two transmissible plasmids in Rhizobium leguminosarum strain 300. J. Gen. Microbiol. 128:85-93.
30
30. Kaneko, T., Y. Nakamura, S. Sato, E. Asamizu, T. Kato, S. Sasamoto, A. Watanabe, K. Idesawa, A. Ishikawa, K. Kawashima, T. Kimura, Y. Kishida, C. Kiyokawa, M. Kohara, M. Matsumoto, A. Matsuno, Y. Mochizuki, S. Nakayama, N. Nakazaki, S. Shimpo, M. Sugimoto, C. Takeuchi, M. Yamada, and S. Tabata. 2000. Complete genome structure of the nitrogen-fixing symbiotic bacterium Mesorhizobium loti. DNA Res. 7:331-338.[Abstract]
31
31. Kaneko, T., Y. Nakamura, S. Sato, K. Minamisawa, T. Uchiumi, S. Sasamoto, A. Watanabe, K. Idesawa, M. Iriguchi, K. Kawashima, M. Kohara, M. Matsumoto, S. Shimpo, H. Tsuruoka, T. Wada, M. Yamada, and S. Tabata. 2002. Complete genomic sequence of nitrogen-fixing symbiotic bacterium Bradyrhizobium japonicum USDA110. DNA Res. 9:189-197.[Abstract]
32
32. Laguerre, G., M. P. Fernandez, V. Edel, P. Normand, and N. Amarger. 1993. Genomic heterogeneity among french Rhizobium strains isolated from Phaseolus vulgaris L. Int. J. Syst. Bacteriol. 43:761-767.[Abstract/Free Full Text]
33
33. Lai, E. M., and C. I. Kado. 2000. The T-pilus of Agrobacterium tumefaciens. Trends Microbiol. 8:361-369.[CrossRef][Medline]
34
34. Leemans, J., G. Soberon, M. A. Cevallos, L. Fernandez, M. A. Pardo, H. de la Vega, M. Flores, C. Quinto, and R. Palacios. 1984. General organization in Rhizobium phaseoli nif plasmids, p. 710. In C. Veeger and W. E. Newton (ed.), Advances in nitrogen fixation research. Nijhoff-Junk-Pudoc, The Hague, The Netherlands.
35
35. Li, P. L., and S. K. Farrand. 2000. The replicator of the nopaline-type Ti plasmid pTiC58 is a member of the repABC family and is influenced by the TraR-dependent quorum-sensing regulatory system. J. Bacteriol. 182:179-188.[Abstract/Free Full Text]
36
36. Marketon, M. M., and J. E. Gonzalez. 2002. Identification of two quorum-sensing systems in Sinorhizobium meliloti. J. Bacteriol. 184:3466-3475.[Abstract/Free Full Text]
37
37. Meade, H. M., S. R. Long, G. B. Ruvkun, S. E. Brown, and F. M. Ausubel. 1982. Physical and genetic characterization of symbiotic and auxotrophic mutants of Rhizobium meliloti induced by transposon Tn5 mutagenesis. J. Bacteriol. 149:114-122.[Abstract/Free Full Text]
38
38. Mercado-Blanco, J., and J. Olivares. 1993. Stability and transmissibility of the cryptic plasmids of Rhizobium meliloti GR4: their possible use in the construction of cloning vectors for rhizobia. Arch. Microbiol. 160:477-485.[CrossRef]
39
39. Mercado-Blanco, J., and N. Toro. 1996. Plasmids in rhizobia: the role of nonsymbiotic plasmids. Mol. Plant-Microbe Interact. 9:535-545.
40
40. Perret, X., C. Freiberg, A. Rosenthal, W. J. Broughton, and R. Fellay. 1999. High-resolution transcriptional analysis of the symbiotic plasmid of Rhizobium sp. NGR234. Mol. Microbiol. 32:415-425.[CrossRef][Medline]
41
41. Pretorius-guth, I. M., A. Pühler, and R. Simon. 1990. Conjugal transfer of megaplasmid-2 between Rhizobium meliloti strains in alfalfa nodules. Appl. Environ. Microbiol. 56:2354-2359.[Abstract/Free Full Text]
42
42. Quinto, C., H. Delavega, M. Flores, J. Leemans, M. A. Cevallos, M. A. Pardo, R. Azpiroz, M. D. Girard, E. Calva, and R. Palacios. 1985. Nitrogenase reductase: a functional multigene family in Rhizobium phaseoli. Proc. Natl. Acad. Sci. USA 82:1170-1174.[Abstract/Free Full Text]
43
43. Rosenberg, C., and T. Huguet. 1984. The pAtC58 plasmid of Agrobacterium tumefaciens is not essential for tumor-induction. Mol. Gen. Genet. 196:533-536.[CrossRef]
44
44. Sambrook, J., E. F. Fritsch, and T. Maniatis. 1989. Molecular cloning: a laboratory manual, 2nd ed.Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.
45
45. Santos, A. G. D., S. Brom, and D. Romero. 1996. Rhizobium plasmids in bacteria legume interactions. World J. Microbiol. Biotechnol. 12:119-125.[CrossRef]
46
46. Schafer, A., A. Tauch, W. Jäger, J. Kalinowski, G. Thierbach, and A. Pühler. 1994. Small mobilizable multi-purpose cloning vectors derived from the Escherichia coli plasmids pK18 and pK19: selection of defined deletions in the chromosome of Corynebacterium glutamicum. Gene 145:69-73.[CrossRef][Medline]
47
47. Segovia, L., D. Pinero, R. Palacios, and E. Martínez-Romero. 1991. Genetic structure of a soil population of nonsymbiotic Rhizobium leguminosarum. Appl. Environ. Microbiol. 57:426-433.[Abstract/Free Full Text]
48
48. Short, J. M., J. M. Fernandez, J. A. Sorge, and W. D. Huse. 1988. Lambda ZAP: a bacteriophage lambda expression vector with in vivo excision properties. Nucleic Acids Res. 16:7583-7600.[Abstract/Free Full Text]
49
49. Simon, R., U. Priefer, and A. Pühler. 1983. A broad host range mobilization system for in vivo genetic-engineering transposon mutagenesis in gram-negative bacteria. Bio/Technology 1:784-791.[CrossRef]
50
50. Sullivan, J. T., B. D. Eardly, P. van Berkum, and C. W. Ronson. 1996. Four unnamed species of nonsymbiotic rhizobia isolated from the rhizosphere of Lotus corniculatus. Appl. Environ. Microbiol. 62:2818-2825.[Abstract]
51
51. Sullivan, J. T., and C. W. Ronson. 1998. Evolution of rhizobia by acquisition of a 500-kb symbiosis island that integrates into a phe-tRNA gene. Proc. Natl. Acad. Sci. USA 95:5145-5149.[Abstract/Free Full Text]
52
52. Tong, Z. K., and M. J. Sadowsky. 1994. A selective medium for the isolation and quantification of Bradyrhizobium japonicum and Bradyrhizobium elkanii strains from soils and inoculants. Appl. Environ. Microbiol. 60:581-586.[Abstract/Free Full Text]
53
53. Tun-Garrido, C., P. Bustos, V. Gonzalez, and S. Brom. 2003. Conjugative transfer of p42a from Rhizobium etli CFN42, which is required for mobilization of the symbiotic plasmid, is regulated by quorum sensing. J. Bacteriol. 185:1681-1692.[Abstract/Free Full Text]
54
54. Turner, S. L., K. A. L. Knight, and J. P. W. Young. 2002. Identification and analysis of rhizobial plasmid origins of transfer. FEMS Microbiol. Ecol. 42:227-234.[CrossRef]
55
55. Vlassak, K. M., and J. Vanderleyden. 1997. Factors influencing nodule occupancy by inoculant rhizobia. Crit. Rev. Plant Sci. 16:163-229.[CrossRef]

---

Journal of Bacteriology, September 2004, p. 5753-5761, Vol. 186, No. 17
0021-9193/04/$08.00+0     DOI: 10.1128/JB.186.17.5753-5761.2004
Copyright © 2004, American Society for Microbiology. All Rights Reserved.

This article has been cited by other articles:

• Sepulveda, E., Perez-Mendoza, D., Ramirez-Romero, M. A., Soto, M. J., Lopez-Lara, I. M., Geiger, O., Sanjuan, J., Brom, S., Romero, D. (2008). Transcriptional Interference and Repression Modulate the Conjugative Ability of the Symbiotic Plasmid of Rhizobium etli. J. Bacteriol. 190: 4189-4197 [Abstract] [Full Text]  
• MacLean, A. M., Finan, T. M., Sadowsky, M. J. (2007). Genomes of the Symbiotic Nitrogen-Fixing Bacteria of Legumes. Plant Physiol. 144: 615-622 [Full Text]  
• Perez-Mendoza, D., Lucas, M., Munoz, S., Herrera-Cervera, J. A., Olivares, J., de la Cruz, F., Sanjuan, J. (2006). The Relaxase of the Rhizobium etli Symbiotic Plasmid Shows nic Site cis-Acting Preference. J. Bacteriol. 188: 7488-7499 [Abstract] [Full Text]  
• Perez-Mendoza, D., Sepulveda, E., Pando, V., Munoz, S., Nogales, J., Olivares, J., Soto, M. J., Herrera-Cervera, J. A., Romero, D., Brom, S., Sanjuan, J. (2005). Identification of the rctA Gene, Which Is Required for Repression of Conjugative Transfer of Rhizobial Symbiotic Megaplasmids. J. Bacteriol. 187: 7341-7350 [Abstract] [Full Text]  
• Brom, S., Girard, L., Tun-Garrido, C., Garcia-de los Santos, A., Bustos, P., Gonzalez, V., Romero, D. (2004). Transfer of the Symbiotic Plasmid of Rhizobium etli CFN42 Requires Cointegration with p42a, Which May Be Mediated by Site-Specific Recombination. J. Bacteriol. 186: 7538-7548 [Abstract] [Full Text]  

This Article Abstract Full Text (PDF) Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Reprints and Permissions Copyright Information Books from ASM Press MicrobeWorld Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Pérez-Mendoza, D. Articles by Sanjuán, J. Search for Related Content PubMed PubMed Citation Articles by Pérez-Mendoza, D. Articles by Sanjuán, J.