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Journal of Bacteriology, August 2006, p. 5364-5373, Vol. 188, No. 15
0021-9193/06/$08.00+0     doi:10.1128/JB.00521-06
Copyright © 2006, American Society for Microbiology. All Rights Reserved.

Phase Variation and Genomic Architecture Changes in Azospirillum

Ludovic Vial, Céline Lavire, Patrick Mavingui, Didier Blaha,{dagger} Jacqueline Haurat, Yvan Moënne-Loccoz, René Bally, and Florence Wisniewski-Dyé*

UMR CNRS 5557 Ecologie Microbienne, IFR 41 Bio-Environnement et Santé, Université Lyon 1, 43 Boulevard du 11 Novembre 1918, 69622 Villeurbanne Cedex, France

Received 12 April 2006/ Accepted 25 May 2006


   ABSTRACT
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
The plant growth-promoting rhizobacterium Azospirillum lipoferum 4B generates in vitro at high frequency a stable nonswimming phase variant designated 4VI, which is distinguishable from the wild type by the differential absorption of dyes. The frequency of variants generated by a recA mutant of A. lipoferum 4B was increased up to 10-fold. The pleiotropic modifications characteristic of the phase variant are well documented, but the molecular processes involved are unknown. Here, the objective was to assess whether genomic rearrangements take place during phase variation of strain 4B. The random amplified polymorphic DNA (RAPD) profiles of strains 4B and 4VI differed. RAPD fragments observed only with the wild type were cloned, and three cosmids carrying the corresponding fragments were isolated. The three cosmids hybridized with a 750-kb plasmid and pulse-field gel electrophoresis analysis revealed that this replicon was missing in the 4VI genome. The same rearrangements took place during phase variation of 4BrecA. Large-scale genomic rearrangements during phase variation were demonstrated for two additional strains. In Azospirillum brasilense WN1, generation of stable variants was correlated with the disappearance of a replicon of 260 kb. For Azospirillum irakense KBC1, the variant was not stable and coincided with the formation of a new replicon, whereas the revertant recovered the parental genomic architecture. This study shows large-scale genomic rearrangements in Azospirillum strains and correlates them with phase variation.


   INTRODUCTION
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Azospirillum is a plant growth-promoting rhizobacterium associated with roots of monocots, including important crops, such as wheat, corn, and rice. Both in greenhouse and in field trials, Azospirillum was shown to exert beneficial effects on plant growth and crop yields, under various soil and climatic conditions (15). The actual benefit from biological nitrogen fixation has been questioned, and plant growth promotion by Azospirillum seems to be due mainly to production of phytohormones (9, 15). The most abundant phytohormone produced is indole-3-acetic acid, allowing an increase in the number of lateral roots and root hairs; this results in a higher absorption of water and minerals from the soil (9).

Bacterial populations, especially in soil or in the rhizosphere, have to endure fluctuating environmental conditions. Bacteria have evolved different strategies to adapt to these environments. Phase variation is one adaptive process by which bacteria undergo frequent, usually reversible phenotypic changes resulting from genetic or epigenetic alterations at specific genetic loci (29). This process is used by several bacterial species to generate intrapopulation diversity that increases bacterial fitness and is important in niche adaptation or to escape host defenses (reviewed in references 46 and 55). In contrast to spontaneous mutations, which occur at a frequency of approximately 10–8 to 10–6 mutations per growing cell per generation, phase variation occurs at frequencies higher than 10–5 switches per cell per generation (29). Various mechanisms control phase variation. These include DNA inversion or duplication, deletion, transposition, homologous recombination, slipped-strand mispairing, and differential methylation (reviewed in reference 55).

Phase variation in pathogenic bacteria—for example, switching of type IV pili in Neisseria gonorrhoeae (27), differential expression of surface layer proteins in Campylobacter fetus (16), and loss of virulence in the phytopathogen Ralstonia solanacearum (43)—has been extensively studied. However, phase variation is not restricted to pathogenic bacteria. Indeed, it also occurs during rhizosphere colonization of various plants by several strains of beneficial plant-associated Pseudomonas (1, 48). For instance, the regulation of biocontrol traits (production of antifungal metabolites, chitinases, and biosurfactants) by phase variation was reported for Pseudomonas spp. strains (54).

Azospirillum lipoferum 4B, a strain isolated from a rice rhizosphere, generates in vitro at high frequencies (10–4 to 10–3 per cell per generation) a stable phase variant named 4VI (3, 30). Variant colonies are readily distinguishable from wild-type colonies by the differential absorption of dyes incorporated into the growth medium. The 4VI variant exhibits pleiotropic modifications; it gained assimilation of certain sugars but lost the ability to assimilate other sugars (3), to reduce triphenyl tetrazolium chloride, to bind some dyes, to swim (4), and to reduce nitrous oxide (our unpublished results). A. lipoferum 4T, a nonswimming strain displaying all of the features of the 4VI variant, and strain A. lipoferum 4B have been isolated simultaneously from rice rhizosphere at the same frequency (8). A. lipoferum 4T retains the ability to efficiently colonize rice roots (2). Like 4VI, A. lipoferum 4T was found to be genetically very close to A. lipoferum 4B (3, 8, 28), suggesting that A. lipoferum 4T could in fact be a 4VI variant of strain 4B generated within the soil ecosystem.

We recently showed that inactivating recA in strain 4B resulted in a higher frequency of generation of variants (57), contrasting with many studies of other bacteria showing either no effect or the abolition of phase variation process in recA mutants. This finding suggests the possibility that phase variation is accompanied by genetic rearrangements. As of now, the molecular mechanism underlying these nonreversible changes in A. lipoferum 4B remains to be determined. In this study, the objective was to determine whether genomic rearrangements take place during phase variation of strain 4B and related Azospirillum strains. The current work shows that genomic rearrangements are concomitant with phase variation in three species of Azospirillum.


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Media and bacterial growth conditions. The strains and plasmids used in this study are presented in Table 1. Strains of Escherichia coli were grown in Luria-Bertani (LB) broth or agar at 37°C with ampicillin at 100 µg ml–1 (47). Azospirillum strains were grown at 28°C in modified Luria-Bertani (LBm, containing only 5 g liter–1 of NaCl) or in nitrogen-free basal supplemented with 0.025% LBm (Nfbm) (42). Rhizobium etli CFN42 was grown in tryptone-yeast extract (TY) medium supplemented with 6 mM CaCl2 (11).


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  		TABLE 1. Bacterial strains and plasmids

 
DNA manipulations. Preparations of plasmid DNA or total genomic DNA, restriction digestions, ligations, Southern blotting, and hybridization were carried out according to established protocols (47). Transformation of E. coli DH10B cells was carried out by electroporation using a Gene Pulser apparatus (Bio-Rad Laboratories, Hercules, CA) according to the manufacturer's instructions. A genomic library of A. lipoferum 4B was constructed in E. coli Epi-100 using a pWEB cosmid-cloning kit (Epicentre, Madison, WI). DNA amplifications by PCR were performed according to the Taq polymerase manufacturer's instructions (Fermentas, Burlington, Canada). Sequencing was performed by Genome Express (Genome Express, Grenoble, France) using primers M13 (CGCCAGGGTTTTCCCAGTCACGAC) and T7 (ATTATGCTGAGTGATATCCCTCT). Searches for identity to known DNA and protein sequences in databases were performed using the BLAST algorithm (5). Searches for putative open reading frames (ORFs) were done using the FRAMEPLOT program (33).

RAPD analysis. Random amplified polymorphic DNA (RAPD) analysis was performed on cell lysates, using 0.5 U Taq DNA polymerase (Invitrogen, Cergy Pontoise, France) and primer F1253 (5'-GTTTCCGCCC-3') (22). Cells from a single colony were resuspended in 100 µl of H2O, heated for 10 min, and then cooled immediately on ice. Five microliters of this suspension was used for the PCR in 50 µl reaction mixture. Amplification cycles were as follows: (i) 95°C for 5 min; (ii) 95°C for 45 s, 36°C for 1 min, and 72°C for 2 min (45 cycles); (iii) 72°C for 7 min; and (iv) 60°C for 10 min. PCR products were analyzed by 2% agarose gel electrophoresis.

Plasmid content and pulsed-field gel electrophoresis (PFGE). Plasmid profiles were determined by a modified Eckhardt agarose gel electrophoresis technique as described previously (32). Azospirillum isolates were grown in Nfbm until the optical density at 580 nm reached 0.5, and 150 µl of culture was used per well. Electrophoresis was carried out at 5 V for 30 min and 85 V for 7 h at 4°C on a 0.7% agarose gel containing 1% sodium dodecyl sulfate. Plasmid sizes were estimated by comparison with those of R. etli CFN42 (45) and Azospirillum brasilense Sp7 (14). For Southern blot experiments, plasmids were transferred to GeneScreen Plus nylon filters (PerkinElmer, Zaventem, Belgium). PCR fragments randomly labeled with [{alpha}-32P]dCTP using a random-primed DNA labeling kit (Boehringer Mannheim, Mannheim, Germany) were used as probes.

PFGE was carried out as previously described (40). Briefly, Azospirillum strains were grown at 28°C in LBm until mid-log phase and harvested by centrifugation at 5,000 x g. The pellet was washed twice in TE buffer (10 mM Tris-base [pH 7.5], 0.1 mM EDTA) and finally resuspended in TE to achieve a concentration of 5 x 109 cells per ml. An identical volume of 1.6% low-melting agarose (Euromedex, Mundolsheim, France) prepared in water was added to the cell suspension. The mixture was poured into molds (100 µl well–1; Bio-Rad Laboratories) and cooled at 4°C for 30 min. Plugs were treated with 1.2 mg ml–1 lysozyme (Roche, Meylan, France) for 24 h at 37°C, briefly rinsed with TE, and then treated for 48 h at 37°C with 2 mg ml–1 proteinase K (Euromedex) dissolved in 0.5 M EDTA containing 1% sarkosyl (Sigma, St. Quentin Fallavier, France). The plugs were then washed in TE buffer and stored at 4°C in 0.5 M EDTA until used. PFGE was carried out on a Chef-DRIII system (Bio-Rad Laboratories) at 14°C on a 0.8% agarose gel (pulse field-certified agarose; Bio-Rad Laboratories) in 0.5x TBE (Tris-borate-EDTA) buffer (Euromedex). Conditions of migration for separation of plasmids were 5 V cm–1 for 15 h with a switch time of 20 to 40 s and a field angle of 120°, followed by 3.5 V cm–1 for 30 h with a switch time of 120 to 240 s and a field angle of 120°. For separation of the biggest replicons, the conditions were 4.5 V cm–1 for 60 h with a switch time of 70 to 160 s and a field angle of 120°.

The 750-kb plasmid was purified from a low-melting PFGE agarose gel, digested by EcoRV, labeled with [{alpha}-32P]dCTP, and used as a probe in hybridization experiments.

Phenotypic characterization of variants in Azospirillum strains. The ability of strains to generate variants was analyzed as previously described (3, 57). Briefly, single wild-type colonies were used to inoculate 5 ml of Nfbm medium. After overnight incubation at 28°C with shaking (200 rpm), serial dilutions were plated onto nutrient agar (Difco) supplemented with 0.0005% (wt/vol) bromothymol blue (NAB medium). The morphology of colonies on NAB plates and dye binding were examined after 4 days of incubation at 28°C. For each strain, at least 500 colonies were screened. The stabilities of the variants were tested by growing overnight individual variant colonies in Nfbm medium at 28°C and plating them on NAB. The frequencies of variant emergence were calculated for individual colonies after a 4-day growth on NAB as previously described (57).


   RESULTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Evidence for DNA rearrangements in the A. lipoferum 4B strain. Since phase variation on strain A. lipoferum 4B has been extensively studied and the frequency of variants generated by a 4BrecA mutant was increased up to 10-fold, we focused first on genome rearrangements taking place when 4B switches to 4VI. Strains 4B and 4BrecA as well as their phenotypic variants 4VI and 4VIrecA were compared by RAPD analysis using primers allowing differentiation of A. lipoferum strains (22). RAPD profiles of A. lipoferum 4B and its 4VI variant revealed two strong major reproducible differences: two fragments of approximately 600 bp and 850 bp were repeatedly absent from the 4VI profile (Fig. 1, lanes 1 and 2). The same result was observed for strain 4BrecA and its 4VIrecA variant (Fig. 1, lanes 3 and 4). This finding suggests that genomic rearrangements occur during phase variation and that the same events could take place in the recA mutant.


Figure 1
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  		FIG. 1. RAPD profiles of A. lipoferum 4B and its phenotypic variant 4VI. Lane 1, wild-type strain 4B; lane 2, 4VI variant; lane 3, strain 4BrecA; lane 4, 4VIrecA variant; lane M, molecular weight marker (123-bp ladder).

 
The two major RAPD fragments present in the 4B profile but absent from that of 4VI were cloned in pGEM-T Easy vector and sequenced. As a RAPD band might contain several amplification products, different clones were analyzed for each band. A unique DNA sequence, RAPD1, was obtained with the 600-bp band (AY913829). From the 850-bp band, two sequences of 854 and 846 bp, named RAPD2 and RAPD3 (DQ242496 and DQ242497, respectively), were recovered. The sequences from RAPD1 and RAPD3 displayed no ORF and no significant similarity to sequences found in databases. A partial ORF encoding a putative transposase (with a higher level of identity to a transposase of Caulobacter crescentus, NP_421536) was found over the 135 bp of the 3' end of the RAPD2 fragment.

In order to determine whether the three RAPD regions were absent from the genome of the 4VI variant or still present but at another location in the genome (both situations could indeed explain the disappearance of bands in an RAPD profile), sets of primers were designed (Table 2). Using these primers, PCR fragments of the expected sizes were obtained from strain 4B, whereas no amplification product was obtained from 4VI (data not shown). To confirm the PCR data, Southern hybridization experiments were carried out on digested genomic DNA using the PCR products obtained from RAPD1, RAPD2, and RAPD3 as probes. The three probes clearly hybridized with DNA from 4B, but no hybridization signal was detected with DNA from 4VI (see Fig. 2 for hybridization with RAPD2; data not shown for RAPD1 and RAPD3). Overall, these findings point to a deletion event taking place when strain 4B switches to the 4VI variant. Moreover, the occurrence of a single hybridization signal with each probe suggested that those three regions were present as single copies in the genome of A. lipoferum 4B.


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  		TABLE 2. Primers and sizes of amplified fragments

 

Figure 2
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  		FIG. 2. Southern blot hybridization of genomic DNA and RAPD profiles of A. lipoferum 4B and the 4VI variant. RAPD2 was used as a probe. Lanes 1, 3, 5, and 7, DNA from 4B digested with BamHI, BglII, EcoRV, and PvuII, respectively; lanes 2, 4, 6 and 8, DNA from 4VI digested with BamHI, BglII, EcoRV, and PvuII, respectively; lane 9, RAPD profile from 4B; lane 10, RAPD profile from 4VI.

 
Searching the extent of the deletion in the 4VI variant genome. To determine the extent of the deletion as well as the putative genetic link between the three RAPD regions, a genomic library of strain 4B was constructed in cosmid pWEB; 400 clones were screened by PCR with specific primers targeting RAPD1, RAPD2, and RAPD3. For each region, a unique positive clone was isolated and the corresponding cosmids were named pR2.51, pR2.52, and pR2.53 for RAPD1, RAPD2 and RAPD3, respectively. These cosmids carried an insert, each about 35 kb DNA, and displayed different restriction patterns (data not shown). The boundaries of each insert were sequenced using the M13 forward and T7 primers.

BLAST analyses revealed no ORF for the 650- and 746-bp sequences of the left borders of cosmids pR2.51 and pR2.52, respectively (DQ242498 and DQ242500). Sequences corresponding to the right borders showed a partial ORF displaying 62% identity with an ORF encoding a signal transduction histidine kinase of Bradyrhizobium japonicum (NP_768355) for pR2.51 (744 bp; DQ242499), and that of pR2.52 (DQ242501) carried an ORF whose product shared 37% identity (55% similarity) with the VirB4 component of the type IV secretion system of Mesorhizobium sp. BNC1 (ZP_00613241). Finally, sequencing of 733 bp of the left border of pR2.53 (DQ242502) revealed a partial ORF with 81% identity to a gene encoding a protein involved in Fe2+ transport of Ralstonia metallidurans (ZP_00596430); the 847 bp of the right boundary of this cosmid (DQ242503) showed 79% identity with an ORF coding for a predicted oxidoreductase of Mesorhizobium sp. BNC1 (ZP_00613187).

Sets of primers for each boundary were designed (Table 2) and used in PCR experiments. PCR fragments of the expected sizes were obtained from strain 4B with the six primer sets; no product was amplified from the 4VI DNA with any of the primer sets (data not shown), suggesting that the entire content of all three cosmids was missing from the 4VI genome. This means that at least 110 kb was deleted during phase variation.

Modification of plasmid content during phase variation. Previous results obtained with the classical Eckhardt technique (18) showed no difference in plasmid pattern between 4B and 4VI, both harboring five plasmids (3, 28). When a modified technique of plasmid separation was performed (see Materials and Methods), the same profile could be observed for both strains (Fig. 3A, lane 1, for strain 4B), and using plasmid profiles of R. etli CFN42 and A. brasilense Sp7 as markers, the sizes of those plasmids were estimated to be 40, 310, 460, 700, and ~1,000 kb. However, when the conditions of migration applied to the gel were chosen to improve separation of high-molecular-weight plasmids (see Materials and Methods), the 40-kb plasmid was no longer visible but a sixth plasmid of 750 kb appeared close to the 700-kb one in strain 4B (Fig. 3B, lane 1). Interestingly, the 750-kb replicon was repeatedly absent from the plasmid profile of 4VI (Fig. 3B, lane 2). The same results were observed for strain 4BrecA and its 4VIrecA variant (data not shown). We also checked the plasmid pattern of the original 4B isolate, which was obtained in 1982 and lyophilized and conserved since, and found it to be identical to that of the 4B strain used throughout this study (data not shown). This indicates that no genetic drift has occurred in the strain. The original 1982 isolate could generate variants at the same frequency, pointing to the fact that the ability to generate variants is an intrinsic property of strain 4B. Moreover, the 750-kb plasmid was also absent from those variants.


Figure 3
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  		FIG. 3. Modification of plasmid pattern when A. lipoferum 4B switches to the 4VI variant. Electrophoretic profile of plasmids in A. lipoferum 4B and 4VI were obtained by a modified Eckhardt procedure (A) with a prolonged migration time (B; see text). (C) Southern blot hybridization of the plasmid profile shown in panel B, using RAPD2 as a probe. Lane 1, strain 4B; lane 2, 4VI variant.

 
To establish whether the deleted regions identified following RAPD analysis and cosmid library screening were located on the missing 750-kb plasmid or not, DNA hybridizations of plasmid profiles were carried out using PCR products RAPD1, RAPD2, and RAPD3 as probes. As suspected, the three probes clearly hybridized with the 750-kb plasmid missing from the variant, as illustrated for RAPD2 (Fig. 3C). This result clearly indicates that a major deletion event (of at least 110 kb of plasmid DNA) occurs when strain 4B switches to 4VI, explaining why revertants cannot be observed in this strain. However, whether the 750-kb plasmid was entirely lost or part of it integrated into a bigger replicon was explored by PFGE analysis and hybridization.

It is known that in PFGE, higher-molecular-weight circular DNAs cannot enter the agarose; thus, only a fraction of randomly linearized replicons is expected to migrate during electrophoresis. PFGE of undigested DNA revealed the presence of plasmids with banding patterns similar to those described above for 4B and 4VI (Fig. 4A). The genomic architecture of the nonmotile strain A. lipoferum 4T isolated from the same rice rhizosphere as strain 4B and displaying the same features as the 4VI variant was also analyzed by PFGE. Strain 4T carries replicons similar to those in the 4VI variant, and in particular, it lacks the 750-kb plasmid (Fig. 4A, lane 3). Interestingly, an additional replicon of 400 kb was detected in strain 4T. When using appropriate conditions of migration for separating the bigger replicons, two similar replicons of approximately 2,200 kb and <1,600 kb were detected in the three strains (Fig. 4C). To ascertain that the 750-kb replicon is lost, the corresponding plasmid band was purified and used as a probe to hybridize PFGE profiles of small and large replicons. The only hybridization signal was obtained with the 750-kb plasmid of strain 4B (Fig. 4B). No hybridization signal was detected on bigger replicons of the 4VI variant (Fig. 4D). These results clearly demonstrate that the 750-kb plasmid was entirely lost in the variants. The fact that the additional plasmid of strain 4T did not hybridize indicates that the 400-kb plasmid is not a remnant of the 750-kb plasmid.


Figure 4
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  		FIG. 4. PFGE of undigested DNA of Azospirillum wild-type strains and variants. (A) Profile showing well-separated plasmids. (B) Southern blot hybridization of the plasmid profile shown in panel A, using the 750-kb plasmid from strain 4B as a probe. (C) Separation of the bigger replicons. (D) Southern blot hybridization of the profile shown in panel C, using the 750-kb plasmid as a probe. A. lipoferum 4B (lane 1), the 4VI variant (lane 2), and A. lipoferum 4T (lane 3). M, Saccharomyces cerevisiae marker.

 
Link between phenotypic features and plasmid loss. Phase variation is linked with pleiotropic modifications in Azospirillum; notably, the 4VI variant and strain 4T have lost the ability to swim and to reduce nitrous oxide. Attempts to isolate loci involved in flagellin synthesis of polar flagellum were unsuccessful. Primers were designed and used to amplify nosZ, a gene already characterized for several Azospirillum strains and whose product is involved in reducing nitrous oxide. A product of 520 bp was amplified from genomic DNA from strain 4B; no amplification was obtained with DNA from the 4VI variant and from strain 4T. The 4B PCR product was sequenced (DQ465016) and appeared to be 94% identical to nosZ of A. brasilense Sp7 (AF361791). This PCR product was used as a probe on plasmid gel and was found to hybridize with the 750-kb plasmid of strain 4B (data not shown); as expected, no hybridization signal was detected with 4VI and 4T, which is consistent with these strains displaying no nitrous oxide-reducing activity.

Occurrence of phase variation in Azospirillum species. To explore the phase variation phenomena in Azospirillum, a collection of 27 strains (Table 1) belonging to different species was screened by using an approach similar to that used to identify variants generated by A. lipoferum 4B (see Materials and Methods) (3). Five strains (Table 3) belonging to three different Azospirillum species produced variant colonies with the same phenotypes as the 4VI variant: those variants did not fix bromothymol blue (i.e., they were translucent on NAB medium), they were nonmotile (as seen on soft agar and confirmed by microscopy), and they lost the ability to reduce triphenyl tetrazolium chloride into formazan. The other strains failed to produce variants or the frequency of appearance of variants was under the detection threshold.


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  		TABLE 3. Stabilities and frequencies of variants

 
The stabilities of the variants were examined by subculturing the corresponding colonies overnight (eight generations) and plating them onto NAB. As previously reported (3) and confirmed in this study, the 4VI variant from A. lipoferum 4B was found to be stable, generating identical and homogeneous colonies. A similar stable phase variant was also generated by A. brasilense strain WN1 (Table 3). In contrast, subculturing variant colonies from A. brasilense Wb1 and A. irakense KBC1 produced a mixture of wild-type and variant colonies (at a ratio of about 10:1), suggesting that a reversion event had occurred. For strains A. brasilense PH1 and A. lipoferum MRB16, subculturing of variants yielded almost only wild-type colonies, and thus, those variants were considered highly unstable and could not be further analyzed. In all cases, reversion to a wild-type phenotype was confirmed by the observation that cell motility and TTC reduction were restored. The frequencies of appearance of variants were determined for strains generating stable or weakly unstable variants and were found to be in the same order of magnitude as the frequency previously calculated for strain 4B (Table 3).

Genomic rearrangements in other Azospirillum strains. To investigate whether phase variation was also linked to genomic rearrangements in other Azospirillum strains, the genomes of stable and weakly unstable variants were compared to those of the corresponding wild-type strains by a modified Eckhardt gel electrophoresis. For A. brasilense Wb1, no difference in plasmids contents was seen between the wild type and the variant, implying that genetic modifications not detected by this method might occur in that particular strain (Fig. 5A). A plasmid pattern change was observed for A. brasilense WN1, where the wild-type cells harbor five plasmids of approximately 190, 260, 500, 570, and >650 kb, whereas cells from its stable variant carry only four plasmids, with disappearance of the 260-kb replicon (Fig. 5B). Plasmid profile and PFGE analyses of A. irakense KBC1 revealed two plasmids of 1,050 and 1,400 kb, while the plasmid profile of its variant constantly displayed three plasmids of 160, 1,050, and 1,240 kb (Fig. 5C, lanes 1 and 2; data not shown for PFGE). The 1240-kb plasmid is usually seen as a faint band in the gel, which may indicate that only some cells bear this replicon; it seems that an excision event in the 1,400-kb plasmid generates two new replicons. Several revertant colonies obtained from KBC1 variants showed the same phenotypic features and the same plasmid profile as the wild-type strain (Fig. 5C, lane 3), indicating that excision occurring in the 1,400-kb plasmid was followed by cointegration and that these events were probably not accompanied by a loss of genetic material.


Figure 5
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  		FIG. 5. Electrophoretic profile of plasmids of Azospirillum wild-type strains and their corresponding variants. Electrophoretic profile of plasmids obtained by a modified Eckhardt procedure with A. brasilense Wb1 (A), A. brasilense WN1 (B), and A. irakense KBC1 (C). In panel C, lane 2 corresponds to a mixture of cells bearing the two excised replicons (1,240 kb and 160 kb, i.e., variants) and cells containing the cointegrate 1,400-kb plasmid (revertants). Lane 1, wild type; lane 2, variant; lane 3, revertant; M, S. cerevisiae marker.

 
To investigate the possibility that the different plasmids involved in phase variation share common sequences, the 750-kb plasmid DNA was used as a probe to hybridize to plasmid profiles of strains generating variants, i.e., A. brasilense WN1 and Wb1 and A. irakense KBC1. No signal was detected, even with prolonged exposure time (data not shown), suggesting that these plasmids do not share homologous sequences. However, the possibility that diverging orthologs are localized on these plasmids cannot be excluded.


   DISCUSSION
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
In this study, phase variation and DNA rearrangement phenomena in Azospirillum strains were investigated by using media containing dyes, plasmid and PFGE profiles, and RAPD and Southern hybridization of genomic DNA patterns. In addition to the well-described phase variant generated by A. lipoferum 4B, the ability to generate a variant was extended to two other species, A. brasilense and A. irakense. Phase variants of Azospirillum appeared at frequencies of 10–4 to 10–3 per cell per generation, which is similar to those reported for many other bacteria (16, 54). Although the majority of the variants were unstable and reverted to the original phenotypes in these bacteria, variants from strains A. lipoferum 4B and A. brasilense WN1 exhibited stable phenotypes over multiple subculturing. Reversion is common in phase variation phenomena, where revertants tend to recover phenotypic and genetic characteristics of the wild-type strains (25, 54).

The genus Azospirillum comprises eight species (10, 17, 35, 37, 44, 51, 59), and most of them harbor multiple replicons consisting of a chromosome(s) and plasmids (14, 24). So far, the composite genome of Azospirillum has been described as being relatively stable, at least for A. brasilense: indeed, the PFGE profile after field experiments and several isolation procedures remained unchanged for Sp7 and Sp245 (38). Interestingly, here we showed that modification of genome architecture in Azospirillum species occurs in vitro at high frequency (>10–4) during phase variation. Most strains studied present different genomic rearrangements, implying plasmids of various sizes. For strain 4B, all of the data obtained point to the loss of a 750-kb plasmid during conversion from 4B to 4VI. Disappearance of a plasmid in the variant of A. brasilense WN1, excision followed by temporary formation of new replicons in the variant of A. irakense KBC1, and an undetermined discrete event in strain A. brasilense Wb1 were evidenced. These contrasting changes might indicate the involvement of different genetic events. As no similar sequence was found between the 750-kb plasmid of A. lipoferum 4B and the other plasmids involved in phase variation, the participation of common sequences in the genetic rearrangements is unlikely.

DNA rearrangements have previously been involved in bacterial phase variation. For instance, in the human pathogen Yersinia pestis, a deletion of a 102-kb fragment occurs at a frequency of 10–5 and leads to loss of pigmentation and avirulence in a mouse model (23). Antigenic variants of Coxiella burnetii arose by deletions of up to 29 kb (31, 58). Phase variation based on recombinational events of S-layer proteins encoded by genes between the chromosome and plasmids has been also demonstrated for Bacillus stearothermophilus (49). The molecular origin for lipopolysaccharide phase variation in Legionella pneumophila relies on a 30-kb unstable element located on the chromosome in the wild type, whereas excision from the chromosome and replication as a high-copy plasmid result in the avirulent phenotype variant (36). Moreover, major DNA rearrangements between the chromosome and plasmids that are not involved in phase variation were also well documented for many bacteria, including the plant-associated bacterium Rhizobium. These genomic rearrangements arose at 10–4 to 10–3; they include deletion, amplification, inversion, and cointegration (13, 26, 40, 41, 60). Some of these rearrangements have biological consequences (26, 39). Accordingly, it is well established that plasticity is a common feature of bacterial genomes. Our study constitutes the first to demonstrate such genomic rearrangements in Azospirillum genomes.

Unraveling the mechanism by which phase variants arise is not always straightforward, in particular when genetic alterations are foreboded and no markers of the altered regions are available, as was the case for the Azospirillum strains studied here. Classical genetic techniques, such as functional complementation of the variant or transposon mutagenesis, which were used successfully to characterize DNA regions or loci involved in phase variation in certain bacteria (25, 54), were found unproductive in A. lipoferum 4B, to be due mainly to the extremely low frequency of plasmid transfer (7). In this study, by combining RAPD analysis with a genomic library construction, we were able to isolate cosmids containing DNA region involved in phase variation in strain 4B. To date, no Azospirillum genome has been entirely sequenced and the implementation of standard genetic protocols is difficult with A. lipoferum (53). Sequencing the entire content of the isolated cosmids or of the deleted plasmid might allow the elucidation of the mechanism underlying these genetic rearrangements. In strain 4B, the absence of a functional RecA protein does not impede these genomic rearrangements and the frequency of variants generated by 4BrecA is increased up to 10-fold (57). RecA has been shown to affect plasmid stability in two contrasting ways. For instance, in E. coli, RecA expression causes plasmid multimerization with concomitant instability (50), whereas stabilization of plasmid by recA-dependent functions was reported for Zymomonas mobilis (56). Our results suggest that RecA plays a role in the maintenance of the 750-kb plasmid in A. lipoferum 4B, and knocking out the recA gene increases its instability.

Several phenotypes altered here are identical in variants obtained from Azospirillum strains; noticeably, phase variation is always accompanied by loss of motility due to polar flagellum, suggesting that genes implicated in flagellum synthesis or regulation could undergo rearrangements. The lost region represents about 1/10 of the genome of A. lipoferum 4B, which suggests that additional phenotypes altered in the 4VI variant remain to be identified. Sequencing the 750-kb plasmid will provide insight into the biological significance of this major genomic rearrangement occurring in vitro (4VI variant) and in nature (A. lipoferum 4T). It is known that Pseudomonas brassicacearum, a major root colonizer of Arabidopsis thaliana, generates variants with higher abilities to swim and swarm, and thus, phase variation was correlated with the high colonization powers of these bacteria (1). In the case of Azospirillum, it is important to point out that loss of swimming ability in variant 4VI and in strain 4T is directly linked to enhanced swarming motility (4). Thus, the nonswimming Azospirillum strains are expected to keep the ability to move along plant roots. The ability to generate variants was not correlated to a particular Azospirillum species or to a specific plant host, and thus, the adaptive significance of phase variation in Azospirillum remains to be established.


   ACKNOWLEDGMENTS
 
We are grateful to Gladys Alexandre (University of Tennessee) and Philippe Normand (UMR CNRS 5557 Ecologie Microbienne) for helpful discussions, Sajjad Mirza (NIGBE, Faisalabad, Pakistan) for providing Azospirillum strains, and Joël Pothier (UMR CNRS 5557 Ecologie Microbienne) for technical help. We also thank the reviewers for useful comments.

This research was supported by a fellowship from the Ministère de la Recherche et des Nouvelles Technologies to L.V. and by funding from CNRS.


   FOOTNOTES
 
* Corresponding author. Mailing address: UMR CNRS 5557 Ecologie Microbienne, 43 Boulevard du 11 Novembre 1918, 69622 Villeurbanne Cedex, France. Phone: 33 4 72 44 58 89. Fax: 33 4 72 43 12 23. E-mail: wisniews{at}biomserv.univ-lyon1.fr. Back

{dagger} Present address: UMR CNRS INSA 5122 Microbiologie et Génétique, IFR 41 Bio-Environnement et Santé, Université Lyon 1, 43 Boulevard du 11 Novembre 1918, 69622 Villeurbanne Cedex, France. Back


   REFERENCES
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

  1. Achouak, W., S. Conrod, V. Cohen, and T. Heulin. 2004. Phenotypic variation of Pseudomonas brassicacearum as a plant root-colonization strategy. Mol. Plant-Microbe Interact. 17:872-879.[Medline]
  2. Alexandre, G., C. Jacoud, D. Faure, and R. Bally. 1996. Population dynamics of a motile and a non-motile Azospirillum lipoferum strain during rice colonization and motility variation in the rhizosphere. FEMS Microbiol. Ecol. 19:271-278.[CrossRef]
  3. Alexandre, G., and R. Bally. 1999. Emergence of a laccase-positive variant of Azospirillum lipoferum occurs via a two-step phenotypic switching process. FEMS Microbiol. Lett. 174:371-378.[CrossRef][Medline]
  4. Alexandre, G., R. Rohr, and R. Bally. 1999. A phase variant of Azospirillum lipoferum lacks a polar flagellum and constitutively expresses mechanosensing lateral flagella. Appl. Environ. Microbiol. 65:4701-4704.[Abstract/Free Full Text]
  5. Altschul, S. F., T. L. Madden, A. A. Schäffer, 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]
  6. Baldani, V. L. D., M. A. B. Alvarez, J. I. Baldani, and J. Döbereiner. 1986. Establishment of inoculated Azospirillum spp. in the rhizosphere and in roots of field grown wheat and sorghum. Plant Soil 90:37-40.[CrossRef]
  7. Bally, R., and A. Givaudan. 1988. Mobilization and transfer of Azospirillum lipoferum plasmid by the Tn5-Mob transposon into a plasmid-free Agrobacterium tumefaciens strain. Can. J. Microbiol. 34:1354-1357.
  8. Bally, R., D. Thomas-Bauzon, T. Heulin, J. Balandreau, J. Richard, and J. de Ley. 1983. Determination of the most frequent N2-fixing bacteria from the rhizosphere of rice. Can. J. Microbiol. 29:881-887.
  9. Bashan, Y., G. Holguin, and L. E. de-Bashan. 2004. Azospirillum-plant relationships: physiological, molecular, agricultural, and environmental advances (1997-2003). Can. J. Microbiol. 50:521-577.[CrossRef][Medline]
  10. Ben Dekhil, S., M. Cahill, E. Stackebrandt, and L. I. Sly. 1997. Transfer of Conglomeromonas largomobilis subsp. largomobilis to the genus Azospirillum as Azospirillum largomobile comb. nov., and elevation of Conglomeromonas largomobilis subsp. parooensis to the new type species of Conglomeromonas, Conglomeromonas parooensis. sp. nov. Syst. Appl. Microbiol. 20:72-77.
  11. Beringer, J. F. R factor transfer in Rhizobium leguminosarum. J. Gen. Microbiol. 84:188-198.
  12. Blaha, D., C. Prigent-Combaret, M. S. Mirza, and Y. Moënne-Loccoz. 2006. Phylogeny of the 1-aminocyclopropane-1-carboxylic acid deaminase-encoding gene acdS in phytobeneficial and pathogenic Proteobacteria and relation with strain biogeography. FEMS Microbiol. Ecol. 56:455-470.
  13. Brom, S., L. Girard, C. Tun-Garrido, A. Garcia-de los Santos, P. Bustos, V. Gonzalez, and D. Romero. 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/Free Full Text]
  14. Caballero-Mellado, J., L. López-Reyes, and R. Bustillos-Cristales. 1999. Presence of 16S rRNA genes in multiple replicons in Azospirillum brasilense. FEMS Microbiol. Lett. 178:283-287.[CrossRef]
  15. Dobbelaere, S., A. Croonenborghs, T. Amber, D. Ptacek, J. Vanderleyden, P. Dutto, C. Labandera-Gonzalez, J. Caballero-Mellado, J. F. Aguirre, Y. Kapulnik, B. Shimon, S. Burdman, D. Kadouri, S. Sarig, and Y. Okon. 2001. Responses of agronomically important crops to inoculation with Azospirillum. Aust. J. Plant Physiol. 28:1-9.
  16. Dworkin, J., and M. J. Blaser. 1996. Generation of Campylobacter fetus S-layer protein diversity utilizes a single promoter on an invertible DNA segment. Mol. Microbiol. 19:1241-1253.[Medline]
  17. Eckert, B., O. B. Weber, G. Kirchhof, A. Halbritter, M. Stoffels, and A. Hartmann. 2001. Azospirillum doebereinerae sp. nov., a nitrogen-fixing bacterium associated with the C4-grass Miscanthus. Int. J. Syst. Evol. Microbiol. 51:17-26.[Abstract]
  18. Eckhardt, T. 1978. A rapid method for the identification of plasmid desoxyribonucleic acid in bacteria. Plasmid 1:584-588.[CrossRef][Medline]
  19. Elbeltagy, A., K. Nishioka, T. Sato, H. Suzuki, B. Ye, T. Hamada, T. Isawa, H. Mitsui, and K. Minamisawa. 2001. Endophytic colonization and in planta nitrogen fixation by a Herbaspirillum sp. isolated from wild rice species. Appl. Environ. Microbiol. 67:5285-5293.[Abstract/Free Full Text]
  20. Eskew, D. L., D. D. Focht, and P. Ting. 1977. Nitrogen fixation, denitrification and pleomorphic growth in a highly pigmented Spirillum lipoferum. Appl. Environ. Microbiol. 34:582-585.[Abstract/Free Full Text]
  21. Fagès, J., and D. Mulard. 1988. Isolement de bactéries rhizosphériques et effet de leur inoculation en pots chez Zea mays. Agronomie 8:309-314.
  22. Fani, R., G. Damiani, C. Di Serio, E. Gallori, A. Grifoni, and M. Bazzicalupo. 1993. Use of random amplified polymorphic DNA (RAPD) for generating specific DNA probes for microorganisms. Mol. Ecol. 2:243-250.[Medline]
  23. Fetherston, J. D., P. Schuetze, and R. D. Perry. 1992. Loss of the pigmentation phenotype in Yersinia pestis is due to the spontaneous deletion of 102 kb of chromosomal DNA which is flanked by a repetitive element. Mol. Microbiol. 6:2693-2704.[Medline]
  24. Givaudan, A., and R. Bally. 1991. Similarities between large plasmids of Azospirillum lipoferum. FEMS Microbiol. Lett. 78:245-251.
  25. Grewal, S. I., B. Han, and K. Johnstone. 1995. Identification and characterization of a locus which regulates multiple functions in Pseudomonas tolaasii, the cause of brown blotch disease of Agaricus bisporus. J. Bacteriol. 177:4658-4668.[Abstract/Free Full Text]
  26. Guo, X., M. Flores, P. Mavingui, S. I. Fuentes, G. Hernández, G. Dávila, and R. Palacios. 2003. Natural genomic design in Sinorhizobium meliloti novel genomic architectures. Genome Res. 13:1810-1817.[Abstract/Free Full Text]
  27. Hagblom, P., E. Segal, E. Billyard, and M. So. 1985. Intragenic recombination leads to pilus antigenic variation in Neisseria gonorrhoeae. Nature 315:156-158.[CrossRef][Medline]
  28. Haurat, J., D. Faure, R. Bally, and P. Normand. 1994. Molecular relationship of an atypical Azospirillum strain 4T to other Azospirillum species. Res. Microbiol. 145:633-640.[Medline]
  29. Henderson, I. R., P. Owen, and J. P. Nataro. 1999. Molecular switches—the ON and OFF of bacterial phase variation. Mol. Microbiol. 33:919-932.[CrossRef][Medline]
  30. Heulin, T., P. Weinhard, and J. Balandreau. 1983. Motility changes in Azospirillum lipoferum, p. 89-94. In W. Klingmüller (ed.), Azospirillum. II. Experentia supplementum, vol. 28. Birkhäuser Verlag, Basel, Switzerland.
  31. Hoover, T. A., D. W. Culp, M. H. Vodkin, J. C. Williams, and H. A. Thompson. 2002. Chromosomal DNA deletions explain phenotypic characteristics of two antigenic variants, phase II and RSA 514 (crazy), of the Coxiella burnetii Nine Mile strain. Infect. Immun. 70:6726-6733.[Abstract/Free Full Text]
  32. 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]
  33. Ishikawa, J., and K. Hotta. 1999. FramePlot: a new implementation of the Frame analysis for predicting protein-coding regions in bacterial DNA with a high G+C content. FEMS Microbiol. Lett. 174:251-253.[CrossRef][Medline]
  34. Kabir, M., D. Faure, T. Heulin, W. Achouak, and R. Bally. 1995. Azospirillum populations in soils infested by a parasitic weed (Striga) under Sorghum cultivation in Mali, West Africa. Eur. J. Soil Biol. 32:157-163.
  35. Khammas, K. M., E. Ageron, P. A. Grimont, and P. Kaiser. 1989. Azospirillum irakense sp. nov., a nitrogen-fixing bacterium associated with rice roots and rhizosphere soil. Res. Microbiol. 140:679-693.[Medline]
  36. Lüneberg, E., B. Mayer, N. Daryab, O. Kooistra, U. Zahringer, M. Rohde, J. Swanson, and M. Frosch. 2001. Chromosomal insertion and excision of a 30 kb unstable genetic element is responsible for phase variation of lipopolysaccharide and other virulence determinants in Legionella pneumophila. Mol. Microbiol. 39:1259-1271.[CrossRef][Medline]
  37. Magalhaes, F. M., J. I. Baldani, S. M. Souto, J. R. Kuykendall, and J. Döbereiner. 1983. A new acid-tolerant Azospirillum species. An. Acad. Bras. Ciênc. 55:417-430.
  38. Martin-Didonet, C. C., L. S. Chubatsu, E. M. Souza, M. Kleina, F. G. Rego, L. U. Rigo, M. G. Yates, and F. O. Pedrosa. 2000. Genome structure of the genus Azospirillum. J. Bacteriol. 182:4113-4116.[Abstract/Free Full Text]
  39. Mavingui, P., M. Flores, D. Romero, E. Martinez-Romero, and R. Palacios. 1997. Generation of Rhizobium strains with improved symbiotic properties by random DNA amplification (RDA). Nat. Biotechnol. 15:564-569.[CrossRef][Medline]
  40. Mavingui, P., M. Flores, X. Guo, G. Davila, X. Perret, W. J. Broughton, and R. Palacios. 2002. Dynamics of genome architecture in Rhizobium sp. strain NGR234. J. Bacteriol. 184:171-176.[Abstract/Free Full Text]
  41. Mavingui, P., T. Laeremans, M. Flores, D. Romero, E. Martinez-Romero, and R. Palacios. 1998. Genes essential for Nod factor production and nodulation are located on a symbiotic amplicon (AMPRtrCFN299pc60) in Rhizobium tropici. J. Bacteriol. 180:2866-2874.[Abstract/Free Full Text]
  42. Nelson, L. M., and R. Knowles. 1978. Effect of oxygen and nitrate on nitrogen fixation and denitrification by Azospirillum brasilense grown in continuous culture. Can. J. Microbiol. 24:1395-1403.[Medline]
  43. Poussier, S., P. Thoquet, D. Trigalet-Demery, S. Barthet, D. Meyer, M. Arlat, and A. Trigalet. 2003. Host plant-dependent phenotypic reversion of Ralstonia solanacearum from non-pathogenic to pathogenic forms via alterations in the phcA gene. Mol. Microbiol. 49:991-1003.[CrossRef][Medline]
  44. Reinhold, B., T. Hurek, I. Fendrik, B. Pot, M. Gillis, K. Kersters, S. Thielemans, and J. De Ley. 1987. Azospirillum halopraeferens sp. nov., a nitrogen-fixing organism associated with roots of kallar grass (Leptochloa fusca (L.) Kunth). Int. J. Syst. Bacteriol. 37:43-51.
  45. Romero, D., S. Brom, J. Martinez-Salazar, M. L. Girard, R. Palacios, and G. Davila. 1991. Amplification and deletion of a nod-nif region in the symbiotic plasmid of Rhizobium phaseoli. J. Bacteriol. 173:2435-2441.[Abstract/Free Full Text]
  46. Salaün, L., L. A. Snyder, and N. J. Saunders. Adaptation by phase variation in pathogenic bacteria. Adv. Appl. Microbiol. 52:263-301.
  47. Sambrook, J., E. F. Fritsch, and T. Maniatis. 1989. Molecular cloning: a laboratory manual, 2nd ed. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y.
  48. Sánchez-Contreras, M., M. Martin, M. Villacieros, F. O'Gara, I. Bonilla, and R. Rivilla. 2002. Phenotypic selection and phase variation occur during alfalfa root colonization by Pseudomonas fluorescens F113. J. Bacteriol. 184:1587-1596.[Abstract/Free Full Text]
  49. Scholz, H. C., E. Riedmann, A. Witte, W. Lubitz, and B. Kuen. 2001. S-layer variation in Bacillus stearothermophilus PV72 is based on DNA rearrangements between the chromosome and the naturally occurring megaplasmids. J. Bacteriol. 183:1672-1679.[Abstract/Free Full Text]
  50. Summers, D. K., and D. J. Sherratt. 1984. Multimerization of high copy number plasmids causes instability: ColE1 encodes a determinant essential for plasmid monomerization and stability. Cell 36:1097-1103.[CrossRef][Medline]
  51. Tarrand, J. J., N. R. Krieg, and J. Döbereiner. 1978. A taxonomic study of the Spirillum lipoferum group, with descriptions of a new genus, Azospirillum gen. nov. and two species, Azospirillum lipoferum (Beijerinck) comb. nov. and Azospirillum brasilense. sp. nov. Can. J. Microbiol. 24:967-980.[Medline]
  52. Trân Vân, V., O. Berge, D. Faure, R. Bally, P. Hebbar, and T. Heulin. 1997. A simple method for the isolation of Azospirillum strains associated with the rhizosphere of rice. Can. J. Microbiol. 43:486-490.
  53. Vande Broek, A., A. van Gool, and J. Vanderleyden. 1989. Electroporation of Azospirillum brasilense with plasmid DNA. FEMS Microbiol. Lett. 61:177-181.
  54. van den Broek, D., T. F. Chin-A-Woeng, K. Eijkemans, I. H. Mulders, G. V. Bloemberg, and B. J. Lugtenberg. 2003. Biocontrol traits of Pseudomonas spp. are regulated by phase variation. Mol. Plant-Microbe Interact. 16:1003-1012.[Medline]
  55. van der Woude, M. W., and A. J. Baumler. 2004. Phase and antigenic variation in bacteria. Clin. Microbiol. Rev. 17:581-611.[Abstract/Free Full Text]
  56. Vartholomatos, G., M. A. Typas, and C. Drainas. 1993. An ultraviolet-sensitive mutant of Zymomonas mobilis affecting the stability of its natural plasmid pZMO2. Plasmid 29:10-18.[CrossRef][Medline]
  57. Vial, L., J. F. Pothier, P. Normand, Y. Moënne-Loccoz, R. Bally, and F. Wisniewski-Dyé. 2004. Construction of a recA mutant of Azospirillum lipoferum and involvement of recA in phase variation. FEMS Microbiol. Lett. 236:291-299.
  58. Vodkin, M. H., and J. C. Williams. 1986. Overlapping deletion in two spontaneous phase variants of Coxiella burnetii. J. Gen. Microbiol. 132:2587-2594.[Medline]
  59. Xie, C. H., and A. Yokota. 2005. Azospirillum oryzae sp. nov., a nitrogen-fixing bacterium isolated from the roots of the rice plant Oryza sativa. Int. J. Syst. Evol. Microbiol. 55:1435-1438.[Abstract/Free Full Text]
  60. Zhang, X. X., B. Kosier, and U. B. Priefer. 2001. Symbiotic plasmid rearrangement in Rhizobium leguminosarum bv. viciae VF39SM. J. Bacteriol. 183:2141-2144.[Abstract/Free Full Text]

Journal of Bacteriology, August 2006, p. 5364-5373, Vol. 188, No. 15
0021-9193/06/$08.00+0     doi:10.1128/JB.00521-06
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