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Burkholderia Species Are the Most Common and Preferred Nodulating Symbionts of the Piptadenia Group (Tribe Mimoseae)

Burkholderia Species Are the Most Common and Preferred Nodulating Symbionts of the Piptadenia Group (Tribe Mimoseae)

  • Caroline Bournaud, 
  • Sergio Miana de Faria, 
  • José Miguel Ferreira dos Santos, 
  • Pierre Tisseyre, 
  • Michele Silva, 
  • Clémence Chaintreuil, 
  • Eduardo Gross, 
  • Euan K. James, 
  • Yves Prin, 
  • Lionel Moulin


Burkholderia legume symbionts (also called α-rhizobia) are ancient in origin and are the main nitrogen-fixing symbionts of species belonging to the large genus Mimosa in Brazil. We investigated the extent of the affinity between Burkholderia and species in the tribe Mimoseae by studying symbionts of the genera Piptadenia (P.), Parapiptadenia (Pp.), Pseudopiptadenia (Ps.), Pityrocarpa (Py.), Anadenanthera (A.) and Microlobius (Mi.), all of which are native to Brazil and are phylogenetically close to Mimosa, and which together with Mimosa comprise the “Piptadenia group”. We characterized 196 strains sampled from 18 species from 17 locations in Brazil using two neutral markers and two symbiotic genes in order to assess their species affiliations and the evolution of their symbiosis genes. We found that Burkholderia are common and highly diversified symbionts of species in the Piptadenia group, comprising nine Burkholderia species, of which three are new ones and one was never reported as symbiotic (B. phenoliruptrix). However, α-rhizobia were also detected and were occasionally dominant on a few species. A strong sampling site effect on the rhizobial nature of symbionts was detected, with the symbiont pattern of the same legume species changing drastically from location to location, even switching from β to α-rhizobia. Coinoculation assays showed a strong affinity of all the Piptadenia group species towards Burkholderia genotypes, with the exception of Mi. foetidus. Phylogenetic analyses of neutral and symbiotic markers showed that symbiosis genes in Burkholderia from the Piptadenia group have evolved mainly through vertical transfer, but also by horizontal transfer in two species.


Legumes have developed a symbiosis with a polyphyletic group of bacteria commonly called rhizobia. This symbiosis leads to the formation of a specialized organ, the root nodule, within which rhizobia differentiates into bacteroids. Bacteroids fix atmospheric nitrogen, and feed the plant with combined nitrogen in exchange for carbon compounds derived from photosynthesis by the legume host. Over the last several decades, numerous diversity studies have focused on rhizobia, but their diversity and the number of investigated legumes hosts remain far from being complete due to the large number of legume species (>18000) [1].

Most rhizobia belong to a large diversity of alphaproteobacterial genera: Azorhizobium, Allorhizobium, Bradyrhizobium, Mesorhizobium, Rhizobium, Sinorhizobium (Ensifer), Devosia, Methylobacterium, Ochrobactrum, Phyllobacterium, and more recently Aminobacter [2] and Microvirga [3], whereas Burkholderia and Cupriavidus are members of the betaproteobacteria [4], [5], [6], [7]. The terms α and β-rhizobia have thus been raised to distinguish each class of symbionts [7], [8]. Burkholderia is a highly diversified genus, including more than 70 species that have colonised a wide diversity of niches, ranging from soil and water to plants and animals [9], [10].

Diversity studies of rhizobia and legume host range have shown that the vast majority of nodulating legume species interact with α-rhizobia [7]. To date, β-rhizobia are much more restricted in terms of host range, and most species described so far interact with Mimosa species in their major area of diversification in central Brazil and other parts of the tropical World (for a review see [7]). Mimosa species symbionts include mainly Burkholderia species, such as B. tuberum [11], [12], B. mimosarum [13], [14], [15], [16], B. phymatum, [12], [16], [17], B. nodosa [18], B. sabiae [19], B. symbiotica [20], B. diazotrophica [21], and two species of Cupriavidus: C. taiwanensis [5] and C. necator [22]. Interestingly, further studies have shown that nodulation by Burkholderia could be extended to other legumes, such as some native/endemic African and Australian species in the subfamily Papilionoideae [23], [24]. For example, B. tuberum STM678 nodulates many Cyclopia species [23], and it harbors distinct nodulation genes compared to Mimosa-nodulating burkholderia, suggesting South African and South American burkholderias acquired their nodulation genes in distinct transfer events [8], [14], [12], [7].

The relationship between β-rhizobia and Mimosa spp. was investigated in more depth in two studies reported by [11] and [25]. These authors analyzed the diversity of symbionts from nodules of c. 70 diverse Mimosa species growing in the two major biomes of Brazil (Cerrado and Caatinga) in which the genus Mimosa has evolved and diversified into more than 200 native and endemic species [26]. They concluded on the generic character of nodulation in the genus Mimosa that the preferred symbionts of this genus in Brazil are Burkholderia. Identical tree topologies between neutral and symbiotic markers were also observed on Brazilian and French Guianan Mimosa-nodulating burkholderia [11], [12], indicating a monophyletic origin and single acquisition of symbiotic genes by a Burkholderia ancestor, followed by vertical transfer of nodulation genes during species diversification [11]. On the other hand, based on phylogenies of nodulation genes, Cupriavidus taiwanensis was found to be a more recent symbiont of Mimosa spp., which acquired nodulation genes from a Burkholderia ancestor [8], [12], [27]. Although most species of Mimosa are mainly nodulated by Burkholderia spp., and a few by Cupriavidus spp., some of them can also form effective symbioses with α-rhizobia [8], [11], [28], [29]. Later studies have underlined the existence of genetic and environmental factors that could affect the preference of legumes species for nodulation with either α- or β-rhizobia; these include soil pH or the presence of combined nitrogen [11], [12], [16], [24], [25].

The Mimosa genus is closely related to a number of genera including Piptadenia (P.), Parapiptadenia (Pp.), Pityrocarpa (Py.), Anadenanthera (A.), Stryphnodendron (St.), and Microlobius (Mi.) (syn. Goldmania) within the tribe Mimoseae. Jobson & Luckow [30] have investigated the phylogeny of these different genera and subdivided the Piptadenia genus into three clades: the Piptadenia genus sensu stricto (or Eupiptadenia clade) that is a sister clade to Mimosa, and includes, for example P. flava (type species), P. floribunda, P. gonoacantha, P. paniculata; the Pityrocarpa clade (Py. leucoxylon, Py. monoliformis, Py. obliqua) that is closer to the genera Stryphnodendron and Parapiptadenia than to the Eupiptadenia clade; and finally P. viridiflora that is outgrouped from the previous genera and represent a particular case deserving a new generic name [30]. Interestingly, all these genera contain woody species native to South America, particularly to Brazil [31], [30]. Some of them are currently exploited by locals owing to their economical values, such as A. peregrina [32], [33], Pp. rigida [34], [35] and P. gonoacantha [36]. Although their ability to establish associations with rhizobia is documented [37], [38], [39], [40], [41], [42], [43], information about the rhizobial diversity and symbiotic efficiency on these plant species is scarce. A recent study of symbionts of Parapiptadenia rigida in Uruguay demonstrated that this species is nodulated by rhizobial strains belonging to the genera Burkholderia, Cupriavidus and Rhizobium, among which the Burkholderia genotypes were the predominant group [44]. Symbiosis with β-rhizobia in the tribe Mimoseae thus appears to extend outside Mimosa and be more common than previously expected.

In this study our objectives were (i) to investigate the extent of Burkholderia affinity within the tribe Mimoseae by focusing on native species in the Piptadenia genus and in related genera in the Piptadenia Group described by Jobson & Luckow [30]; (ii) to characterize the diversity of rhizobia and their symbiotic genes in this group of legumes; (iii) to examine further symbiotic specificity within the Piptadenia Group.

To achieve these objectives, we isolated rhizobia from diverse species in the Piptadenia Group in their native areas in Brazil, characterized their taxonomic and symbiotic diversity, and assessed their host specificity via coinoculation assays. We found a large diversity of Burkholderia species, but also α-rhizobia, and with a few exceptions an affinity of most plant species towards Burkholderia rather than to α-rhizobia. We discuss the evolutionary patterns of both taxonomic and symbiotic markers and the putative co-adaptation between Burkholderia and Piptadenia Group species.

Materials and Methods

Bacterial Isolates Sampling and Maintenance

Collection of material was authorized by IBAMA N°. 058/2006. The bacterial collection was built through several sampling campaigns performed between 1984 and 2010, from 9 states in Brazil (Rio de Janeiro, Sao Paulo, Minas Gerais, Bahia, Mato Grosso do Sul, Paranà, Pernambuco, Espirito Santo and Pará). Sample locations were concentrated on the main centers of diversification of the Piptadenia Group (Mata Atlantica) and are presented in Figure S1. Details on sampling (gps coordinates, year, season, soil type and pH) are presented in Table S1. STM strains were sampled from trees from various places of Rio de Janeiro State in April 2010; SFM, BR and CVRD strains were sampled from plants growing in plant nursery beds in various states of Brazil (Table S2). Nodules sampled from the field as described in [37] were dried on silicagel until symbiont isolation. Nodules were then rehydrated for 30 min in sterile distilled water, surface sterilized by immersion 30 seconds in 70% ethanol followed by 1 to 3 min in 3% hydrogen peroxide, and washed three times in sterile distilled water. Nodules were then individually crushed and streaked on yeast mannitol agar (YMA, [45]) plates containing bromophenol blue. All YMA plates were incubated at 28°C. Single colonies were picked and checked for purity by repeated streaking and microscope examination. All pure isolates were stored at −80°C in YM broth plus 20% (w/v) glycerol. For Stryphnodendron species, no nodules could be found on two locations, so a plant trapping approach was developed by growing seedlings on a top soil harvested under each tree, and nodules were harvested and processed as described above. As these rhizobia were trapped, they were not treated as natural symbionts, and were only included on Figure 1C to assess rhizobial patterns among the Piptadenia group. Strains from previous studies on P. flava [42], Pp. rigida [44], [46], as well as P. gonoacantha and Pp. pterosperma [46] were also included in this study. A total of 196 strains were included in the study, of which 63 “representative” strains are listed in Table 1 and were chosen as one strain per host plant per geographical origin per unique 16 S rDNA ribotype (as described in molecular method section).

Figure 1. Comparison between Piptadenia’s group plant phylogeny and the occurrence of alpha and beta-rhizobia as nodule symbionts in the field or during coinoculation experiments.

The plant phylogeny (A) is based on a trnL-F/trnK-matK combined dataset, and was built by parsimony with TNT1.1 (default parameters, on using the Jobson & Luckow [30] dataset (downloaded from Treebase, study number S1763, and amended with the P. trisperma from this study). The % of α and β-rhizobia per legume host (in bold) from field sampling (B) or from the coinoculation experiment (C) are represented as white (Burkholderia) and black (α-rhizobia) squares, with the number of strains sampled within each square. *: statistics of symbionts from [44] and [11]. The grey colored square for Stryphnodendron indicates that % of α-rhizobia originates from a trapping experiment on soil (see Material & Methods section).

Plant Nodulation and Specificity Tests

Seeds of P. gonoacantha, Py. monoliformis, Mi. foetidus and A. colubrina were obtained from Instituto Brasileiro de Florestas (Londrina, Paraná Brasil) or Embrapa Agrobiologia (Seropédica; Brazil). M. pudica and siratro seeds were obtained from B&T World Seeds (Paguignan, France) and from University Cheikh Anta Diop (Dakar, Senegal), respectively. P. gonoacantha seeds were scarified and surface sterilized using 96% H2SO4 and 3% calcium hypochlorite (2 min and 3 min, respectively), while A. colubrina, A. peregrina, and Pp. rigida seeds were immersed for 3 min in 3% calcium hypochlorite; and Mi. foetidus seeds were sterilized with 96% H2SO4 during 30 min. Py. monoliformis seeds were germinated by immersion in concentrated H2SO4 for 5 min, then washed with sterile dH2O and afterwards were soaked in 3% calcium hypochloride for 3 min. All species seeds were then washed four times with sterile dH2O, before being germinated on 0.8% water agar plates at 28°C in the dark. M. pudica and siratro seeds were sterilized and germinated as described in [12].

For nodulation tests, all species were grown in Gibson tubes containing Jensen medium [45] filled with dH2O, except Piptadenia species for which tubes were filled with sterile attapulgite (OIL DRI US Special, IIIR, Damolin) and supplemented with sterile dH2O. All plants were then grown in a chamber at 26°C (relative humidity 40%) with a 16 h light/8 h night cycle. Inoculation was performed by adding 1 ml of exponential bacterial culture grown in broth YM medium. Cross-contamination was investigated by using uninoculated negative controls randomly placed between treatments. Plants were observed for nodulation over a period of 30–60 dai, depending on plant species. Nitrogen fixation was estimated by visual observation of plant vigour and foliage color.

For plant-rhizobium coinoculation experiments, plants were grown in Gibson tubes filled with attapulgite, moistened with sterile water and inoculated with 1 ml of an equal mixture of 12 representative strains calibrated at 10 e6 cells using KOVA slides microscope counting. Plants were incubated in the same conditions than as described above, watered with dH2O and harvested 4 weeks post inoculation. Eight nodules were harvested per nodulated plant (12 replicates), surface sterilized 3 min with 3% calcium hypochlorite, placed in a microplate, crushed using a replicator, grown on YMA microplates, and genotyped on recA gene marker as described in the molecular methods section.

Molecular Methods

Extraction of genomic DNA of bacterial strains was performed using the proteinase K lysis procedure described previously [47]. PCR amplifications were performed with GO-Taq Polymerase (Promega) following the manufacturer instructions. All PCR templates were generated with specific primer sets listed in Table S2. 16 S rDNA amplification was carried out as previously described [12]. A 800-bp recA fragment was amplified by PCR and sequenced using the primers recABurk1F and recABurk1R for β-rhizobia, as described in [11], while two specific couples of primers were used for Bradyrhizobium (TSrecAf and TSrecAr) and Rhizobium (recAf and recAr) using PCR conditions described in [48] and [49], respectively. A 440-bp fragment of nifH was amplified and sequenced on Burkholderia strains as described in [8], and on Rhizobium and Bradyrhizobium strains as described in [50]. A 600-bp fragment of nodC was amplified and sequenced for Burkholderia strains using primers described in [11] except for BSP1, BD, BPL and BSa strains for which a specific primer set (nodCPipF and nodCPipR) was designed (500 bp fragment). For α-rhizobia, 600 bp of nodC for Bradyrhizobium and 800 bp for Rhizobium strains were amplified with primer sets nodCfor540-nodCrev1560 and nodCF-nodCI respectively [51], [50]. DNA sequencing was subcontracted to Genoscreen Inc., using ABI3730 sequencers, and the same primers as used for PCR, except for partial 16 S rRNA sequencing for which an internal primer was used (16 S–1080 r).

For the identification of the rhizobial nodule occupant from the plant coinoculation experiment, DNA from surface-sterilized crushed nodule were heat-shocked in 20 µl using a PCR machine (1 cycle of 2 min at 96°C, followed by 7 cycles of 10 seconds at 96°C then 4°C, with a final cycle of 2 min at 4°C), and 1 µl was used to amplify a recA gene fragment as described above. PCR templates were sequenced as previously described.

Plant phylogenetic markers trnL-F and trnk-matK of Stryphnodendron sp. and P. trisperma were PCR-amplified and sequenced as described by [30], from leaves collected from the field.

Phylogenetic and Sequence Analyses

Nucleotide sequences from 16 S rDNA, recA, nodC and nifH genes were corrected with CHROMAS PRO v1.33 (Technelysium Pty Ltd), aligned using Muscle3.6 [52], and alignments were manually curated with GENEDOC [53]. Screening and classification of each 16 S rDNA haplotype (unique sequence in our dataset) was performed using MOTHUR using the unique.seqs command [54]. Phylogenetic trees were constructed by neighbor joining and likelihood methods using MEGA5 [55] and PAUP4 [56], or by Bayesian analyses using Mr Bayes [57] using priors from a GTR+I+G model (with parameters previously estimated by ML under PAUP4). Parsimony analyses on trnL-F+trnK-matK plant markers were performed on TNT1.1 (default parameters, using the Jobson & Luckow [30] dataset (downloaded from Treebase,, study number S1763), amended with new sequence from this study (P. trisperma). Bootstrapping analyses were conducted on MEGA5.

Nucleotide Sequence Accession Numbers

The sequences have been deposited in EMBL database under accession numbers HE983632 to HE983823 and HF536727 to HF536767, and are listed by gene for each strain in Table S3. Sequence alignments are available upon request.


Building a Collection of Rhizobia from Piptadenia Group Species

We sampled at least one representative species of the different clades and genera in the Piptadenia Group (see species in bold in Figure 1A): Piptadenia sensu stricto (the “Eupiptadenia” clade e.g. P. gonoacantha, P. paniculata, P. adiantoides), the Pityrocarpa clade (Py. moniliformis), the P. viridiflora clade, and the Anadenanthera, Parapiptadenia, Stryphnodendron and Pseudopiptadenia clades. The species assignment of each plant host was confirmed by the Botanical garden of Rio de Janeiro (R. Ribeiro). For five species (Stryphnodendron adstringens, P. trisperma, P. paniculata, P. gonoacantha and A. peregrina), the taxonomic position was confirmed by sequencing the trnL-F marker, and comparing it to the Jobson & Luckow dataset [30]. Among all sampled legume species, we confirmed that 17 of them are nodulated, and one, Piptadenia trisperma is a new report for nodulation (according to nodulation data in GRIN database, and [58]). The rhizobial collection was composed of 196 isolates and details are presented in Table 1, where they are classified according to original host legume, representative strain and site of isolation. Representative strains for each location and legume host were chosen according to their 16 S rDNA sequence haplotype (named ribotype in Table 1): one strain per legume host and per sampling location with a unique 16 S rDNA sequence based on a 800 bp alignment was kept for further analyses, and designated as representative strains (for a total of 63 for strains from this work, and 79 when including reference strains from other studies).

Nodulation ability of the representative strains was confirmed either on their original host or on M. pudica and siratro when seeds of the original host could not be obtained (see Table 1, Nodulation column), except for 15 strains which were not tested. No nodulation was observed on M. pudica or siratro by five strains (STM7339, STM7342, STM7420, STM7426, STM7444), but these strains were kept as the original host could not be tested. Details about nodule size, shape and functionality for each Burkholderia species are given in Table S4 and Figure S2, together with pictures of nodules on M. pudica and P. gonoacantha. All nodulating strains appeared to fix nitrogen with M. pudica (based on red color within nodules usually linked to presence of leghaemoglobin, and plant development, see Table S4), except some B. sabiae (carrying nodC1 variant) and B. nodosa (nodC5 variant) strains that produced ineffective nodules. Three nodulation groups were observed on M. pudica as described in Table S4 (all nodules being indeterminate): small inefficient nodules (BSa, BN, Figure S2A), small round shape nodules (BSP1, BD, BPL, BSP4, Figure S2B), and elongated nodules with a peanut shape (BSa, BSP3, Figure S2C).

We also incorporated in our analyses strains BR9001 to BR9004, and BR4812 that were isolated from the Piptadenia Group [46] but which were not previously characterized at the taxonomic level, as well as strains from other studies: two from P. flava (UPRM8020 & UPRM8021) [42], two from P. viridiflora (JPY570, JPY565, Gross et al., unpublished), one each from P. stipulacea (JPY584), Pp. blanchetti (EG100) and Ps. bahiana (EG118) (Gross et al., unpublished), and three frequent genotypes of Burkholderia and Rhizobium from a study of Pp. rigida symbionts in Uruguay [44]. Based on their 16 S rDNA sequence haplotype, the bacterial collection could be grouped into three genera, with 107 strains in the genus Burkholderia, 48 in Rhizobium and 41 in Bradyrhizobium.

In order to establish if patterns between the occurrence of particular symbionts and plant phylogeny existed, we mapped the % of α- and β-rhizobia (as means across all sampling locations) per legume species onto the trnL-trnF matK-trnK phylogeny (Figure 1B). Although some legume species were poorly sampled, Burkholderia symbionts were still found along all the phylogeny of the Piptadenia Group species, and in the majority of most sampled species. A few legume species were devoid of Burkholderia symbionts, but these were mainly poorly sampled species, and so the presence of β-rhizobia in these species cannot be excluded.

Taxonomic Characterization of the Piptadenia Symbionts Collection

To further characterize the rhizobial collection at the taxonomic level and assign a putative species to each strain, a phylogenetic analysis of all representative strains based on a partition of two neutral markers (643 bp of the variable part of the 16 S rRNA gene, and a 644 bp fragment of the recA gene), was built following a Bayesian analysis (see Mat&Methods). The recA gene is a more highly resolved phylogenetic marker than the 16 S rDNA gene, and has proven to be a valuable tool for discriminating species within the Burkholderia genus [11], [12], [59]. We also included type strains of the most closely related species in the genera Burkholderia, Rhizobium and Bradyrhizobium. A phylogenetic tree of the 16 S-recA dataset is presented in Figure 2A (only for the genus Burkholderia, as α-rhizobia are presented in Figure S3); and individual gene marker phylogenies are available as Figure S4. The bacterial collection was clustered into 9 highly supported clades (posterior probabilities >0.9, bootstrap values >75%) in the genus Burkholderia (107 strains), 8 clades in Rhizobium (48 strains) and 4 in Bradyrhizobium (41 strains). In Burkholderia, most strains belonged to clades associated with the species B. nodosa (BN, 31 strains), B. phenoliruptrix (BPL, 27 strains), B. sabiae (BSa, 21 strains), and B. diazotrophica (BD, 18 strains). Other strains belonged to B. phymatum (BP, one strain), B. caribensis (BC, two strains), and three potentially undescribed species: Burkholderia sp. 1, 3 and 5 (BSP1, two strains, BSP3, four strains, BSP4, one strain). Burkholderia strains were identified among all legume species in this study, except for Mi. foetidus, Ps. psilostachya and P. adiantoides. For these three species, few rhizobial isolates were obtained (3–7), all belonging to Rhizobium and/or Bradyrhizobium genera. Each clade includes strains isolated from various hosts from the Piptadenia Group, except the BSP3 clade which was sampled only from P. gonoacantha. Some clades (BSa, BP, BC, BD, BN) contained strains isolated both from the genus Mimosa and the Piptadenia Group, while others were only found associated to the Piptadenia Group (BPL, BSP1, BSP3, BSP4) or to Mimosa species (BM, BSy, BSP2, BT). In the case of the α-rhizobia (Figure S3), strains were clustered into 8 new clades of Rhizobium and 4 of Bradyrhizobium. Rhizobium sp. 1 (RSP1), R. tropici and RSP3 were the most represented clades with 25, 10 and 8 strains, respectively. R. tropici strains were found on the most important number of legume host (6 species). Strains of R. leucaenae and R. gallicum were also found on A. peregrina and P. flava [42], respectively. Rhizobium isolates from Pp. rigida from the Taulé et al. study [44] were grouped within the R. mesoamericanum species (UYPR7.63 & STM3625; Figure S4), a frequently sampled symbiotic species from Mimosa pudica [12], [60], [61]. In Bradyrhizobium, four clades could be distinguished and these were named Bradyrhizobium sp1 (BrSP1) to BrSP4, with no close relationship to any described species (except BrSP4 that was closely related to Br. elkanii). No obvious pattern could be deduced between host plants and α-rhizobial clades. Strains trapped from two soils by Stryphnodendron sp. were only characterized on the basis of their 16 S rDNA, and these grouped closed to Bradyrhizobium sp. 1 (Figure S4).

Figure 2. Comparison of phylogenies between neutral and symbiotic markers in beta-rhizobia.

The neutral marker phylogeny (A) is based on a partition of 800 bp of 16 S rDNA and 600 bp of recA genes, and was built by a Bayesian analysis with priors estimated by ML (see Experimental procedure section). Numbers at ach nodes indicates posterior probabilities (upper number) and bootstraps values % (lower number) from a ML analyses built in parallel (with a GTR+I+G model, 1000 bootstrap replicates). Bootstraps are only indicated when >50%. Node values in bold indicates supported nodes (both by posterior probabilities and bootstrap) retained for clades delineation. The nodC phylogeny is based on 437 bp alignments, and was built by ML using a GTR+I+G model, with 1000 bootstraps replicates (% indicated at tree nodes if >50%). Abbreviations used: B.: Burkholderia, C.: Cupriavidus, BPL: Burkholderia phenoliruptrix, BT: B. tuberum, BSa: B. sabiae, BD: B. diazotrophica, BSy: B. symbiotica, BM: B. mimosarum, BN: B. nodosa, BSP1,2,3,4: Burkholderia sp. 1 to 4, T indicates species type strains. Accession numbers are listed in Table S3.

Distribution of Rhizobial Species per Host Plant and Sampling Locations

A distribution of rhizobial species according to their plant host and site of sampling is presented in Figure 3. We observed a strong site sampling effect since the diversity pattern of encountered species was completely divergent for the same host sampled on different sites. This pattern heterogeneity of symbionts was even found between α and β-rhizobia, since P. gonoacantha and A. peregrina nodules hosted mainly Rhizobium strains in some sites but Burkholderia species on other sites, indicating no strict specificity of these species for α or β-rhizobia. Burkholderia nodosa was particularly frequent on the three different sampling sites of Pp. pterosperma, and was also found in P. trisperma, A. peregrina and P. gonoacantha. Burkholderia sabiae was also a promiscuous species in the Piptadenia Group (found in A. peregrina, P. viridiflora, P. trisperma, A. peregrina). On the other hand, BSP1 and BSP3 species were only found in P. gonoacantha nodules. On the plant side, some species exhibited the same pattern of symbionts diversity in the same geographical area (for example P. trisperma and Pp. pterosperma in Cabo Frio, RJ; Figure 3). However, the opposite could also be found, as P. gonoacantha and P. paniculata in Cabo Frio also exhibited divergent patterns of symbiotic association, one being nodulated by β- and the other by α-rhizobial species.

Figure 3. Distribution of rhizobial species according to host plant and sampling regions.

The % of each species per legume host was calculated according to sampled isolates listed in Table 1. Sampling region is listed below each histogram while on top are indicated the number of isolates. Abbreviations: SER, RJ: Seropédica, Rio de Janeiro; MGS: Mato Grosso do Sul; PAR: Paranà; SP: Sao Paulo; CB, RJ: Cabo Frio, Rio de Janeiro; BU, RJ: Buzios, Rio de Janeiro; ES: Espirito Santo; MG: Minas Geraïs; PHF, SP: Paraibuna horto florestal, Sao Paulo; BA: Xique-Xique, Bahia; IFSP, SP: IFSP, Sao Paulo; Ac: Anadenanthera colubrina, Ap: A. peregrina, Mf: Microlobius foetidus; Pg: Piptadenia gonoacantha; Ppan: P. paniculata; Pt: P. trisperma; Pv: P. viridiflora; Pppt: Parapiptadenia pterosperma; Ppr: Parapiptadenia rigida.

Phylogeny of Symbiosis-related Genes of Piptadenia’s Rhizobia

Phylogenetic analyses were performed on fragments of the nodC (involved in the synthesis of the Nod factor core) and nifH (involved in nitrogen fixation) genes to evaluate the origin and evolution of symbiosis in Piptadenia Group microsymbionts. The nodC and nifH data sets used for phylogenies included representative strains listed in Table 1, as well as reference strains from each clade of the Bontemps et al. [11] and Mishra et al. [12] diversity studies on Mimosa species. A nodC ML phylogeny is shown in Figure 2B, while the nifH ML tree is presented as Figure S5. All nodC sequences from Burkholderia symbiotic species of the Mimoseae tribe were monophyletic compared to α-rhizobia. Burkholderia strains from the Piptadenia Group were clustered into five nodC clades (nodC1 to nodC5 on Figure 2B). Some nodC clades corresponded to already described bacterial species, such as B. nodosa (nodC5), BSP3 (nodC2), while other species had strains carrying different alleles of nodC (e.g. B. sabiae with nodC1 or nodC3 alleles; B. phenoliruptrix with nodC3 and nodC4 alleles). In the case of B. phenoliruptrix, strains carrying either nodC3 or nodC4 were both efficient symbionts of M. pudica, while B. sabiae nodC variants (nodC1, nodC3) exhibited different nodulation phenotypes on M. pudica (inefficient/efficient, Table S4). Some clades carried strains sampled from Mimosa and Piptadenia Group species (nodC3, C5), while others were specific to either the Piptadenia Group (nodC1, C2, C4) or to Mimosa spp. (nodC6, C7). The large nodC3 clade could be divided into 2 sub-clades corresponding to different species: nodC3-1 hosted B. sabiae and BSP1, while nodC3-2 contained B. phymatum, B. phenoliruptrix and B. diazotrophica strains. The large nodC3 clade hosted strains from every studied legume species, except for P. rigida that was restricted to the nodC1 clade. Overall, no clear pattern was identified concerning nodC cladogenesis and legume hosts (except the nodC2 clade that was specific to P. gonoacantha). A better correspondence was found between the nodC and 16 S-recA trees. However, although several 16 S-recA clades corresponded to nodC clades, horizontal gene transfer was also identified in species which carried different nodC alleles (e.g. B. sabiae with BSa-1 and 2, and B. phenoliruptrix with BPL-1 & 2, Figure 2B). The nifH gene ML tree (Figure S5) reflected the same cladogenesis as the nodC tree, but with a few exceptions e.g. some strains in the nodC1 clade (UYPR1.313, UYPR3.611 and Prigida2) were clustered in a different clade in the nifH tree (nifH3).

In the case of α-rhizobia from the Piptadenia Group (Figure S2B), nodC sequences from Rhizobium strains (R. tropici, RSP1, RSP2, R. leucaenae) were clustered in a clade (nodC12) together with R. tropici strains from diverse Mimosa species isolated from Papua New Guinea (NGR181) and Puerto Rico (UPRM8021) [28], [42]. There was no sequence variation on nodC fragment between UPRM8021 and the bean symbiont R. tropici CIAT899T (not shown). The only exceptions were R. gallicum strains from Piptadenia flava (e.g. UPRM8060) that grouped close to the R. etli symbiovar (sv) mimoseae clade (nodC8), and to a R. mesoamericanum strain from Pp. rigida (nodC13) that grouped close to S. meliloti 1021. The nodC and/or nifH genes from some Rhizobium strains isolated from nodules of P. paniculata (STM7339, STM7342) could not be amplified: these strains did not nodulate M. pudica, siratro or P. gonoacantha, and could not be tested on their original host P. paniculata. The symbiotic nature of these two strains thus remains uncertain. Bradyrhizobium strains from P. paniculata and Mi. foetidus clustered in three different clades (nodC9 to nodC11), with the P. paniculata isolates (nodC9, C11) being separated from Mi. foetidus ones (nodC10). Notably, Bradyrhizobium isolates from P. paniculata could not be tested on their original host, but they did nodulate siratro but not M. pudica. As observed for Burkholderia symbionts, the nifH tree (Figure S5) reflected the nodC groupings of the α-rhizobia.

Assessment of Symbiotic Specificity in the Piptadenia Group Using Competition Assays

In order to assess plant specificity in the Piptadenia Group, we carried out a coinoculation experiment with 12 rhizobial genotypes (6 Burkholderia, 3 Rhizobium and 3 Bradyrhizobium strains, representative of species and nodC clades) on 6 Piptadenia Group species (Mf, Pm, Ppr, Pg, Ap, Ac) for which seeds were available. The final range of nodules analyzed were 12 to 60 on 5 to 9 plant replicates. The nodule occupants were then identified as described in the Material and Methods section. Table 2 shows the percentage of nodule occupancy per bacterial genotype for each plant host. The percentage for α or β-rhizobia was also included in Figure 1C for comparison with wild nodule sampling % and plant phylogeny. Overall, all plant species showed a preference for Burkholderia genotypes, except for Mi. foetidus which was nodulated preferentially by bradyrhizobia. One Burkholderia genotype, STM7353 (BN, nodC5) was the most prevalent (in 38 to 85% of nodules) over all other genotypes on all plant hosts (except for Mf, 9% of nodules), followed by STM7317 (BPL, nodC4) that occupied 30 to 33% of plant hosts (except on Mf, 9%, and Pg, 3%). Interestingly, the two B. sabiae genotypes tested (each carrying either the nodC1 or nodC3 allele) did not show the same nodulation pattern: STM7373 (nodC1) was present at around 20% on Mf and Ac while STM7315 (nodC3) was absent, and conversely STM7315 was present at 20% on Pm while STM7373 was absent. Such results indicate potential host plant specificity towards several nodC alleles, although competitiveness for nodulation also interferes with these results. Genotypes from the Rhizobium and Bradyrhizobium genera were poorly competitive, and only STM7379 (RT, nodC2) and STM7378 (BrSP4) were identified in one host, Mi. foetidus (these two strains originating from nodules of this species). It is interesting to note that if Mi. foetidus favors α-rhizobia symbionts, this plant species selected its native genotypes rather than α-rhizobia from P. paniculata, thus underlining specific affinities between this host and its symbionts. Another case of potential host plant specificity is that of STM7296 (BSP3, nodC2), as this poorly competitive strain was only detected on its original plant host, P. gonoacantha.

Table 2. Nodule occupancy (%) in a coinoculation experiment on 6 species of the Piptadenia group.


Burkholderia Species are Common and Highly Diverse Symbionts of the Piptadenia Group

Burkholderia symbiont diversity has been intensively studied on collections isolated from nodules of different species of Mimosa, most of them originating from Brazil [11], [14]. In the present study we extended the nodulation by Burkholderia symbionts to other species in the Piptadenia Group, including six genera that are sister clades to Mimosa in the tribe Mimoseae. The Piptadenia Group contains about 70 species (if excluding the genus Mimosa, that is also part of this group), which are common in tropical and subtropical America, but are mainly native to Brazil [30]. The capacity of nodulation among these clades has been investigated in previous works (see Introduction), but one of them, P. trisperma, is a new report of nodulation from this study. A large diversity of Burkholderia was found, with nine species, five being already described as symbionts of Mimosa species (B. sabiae, B. phymatum, B. caribensis, B. diazotrophica, B. nodosa; [8], [18], [19], [17], [21]), one was not previously reported as containing symbiotic strains (B. phenoliruptrix, [10], [62]), and BSP1, BSP3 and BSP4 are three putative new species, whose taxonomic status is currently being investigated (Bournaud et al., unpublished). This diversity of symbiotic Burkholderia species mirrored the one discovered on 47 species of Mimosa by Bontemps et al [11] in Brazil, which also had seven clades, of which only three overlap with this study and are symbionts of both Mimosa and Piptadenia (B. nodosa, B. diazotrophica and B. sabiae). Overall, diversity studies on symbionts of mimosoid legumes reveal that four rhizobial species were found only associated to the Piptadenia Group (BPL, BSP1, BSP3, BSP4) while four different ones are found only on Mimosa species (BSP2, B. symbiotica, B. mimosarum, B. tuberum). B. tuberum is a frequent species in Mimosa spp. nodules in Brazil and French Guiana [11], [12] but was not detected in Piptadenia group nodules. However, these patterns are only based on our current knowledge, and given the high sampling site effect observed on the diversity of symbionts in this group, it is likely that each bacterial species could also be found in both groups of legume hosts.

The legume ability to form a symbiosis with Burkholderia is not antagonistic with the presence of α-rhizobia, since the two kinds of symbionts were found on six plant species (Table 1, Figure 1B), and some species had only α-rhizobia strains sampled from them. No clear conclusion about affinities of plant species towards α or β-rhizobia could be drawn for several legumes for which the nodule sampling was limited (i.e. Py. monoliformis, P. flava, P. stipulacea, Pseudopiptadenia spp), or for which the site sampling effect was too strong (Fig. 3, P. gonoacantha, A. peregrina). In order to investigate this affinity, we conducted coinoculation tests with 12 genotypes of α and β-rhizobia, and could conclude that Py. monoliformis, Pp. rigida, P. gonoacantha, A. peregrina and A. colubrina had a strong preference towards Burkholderia genotypes, while Mi. foetidus had an affinity for two genotypes of Rhizobium and Bradyrhizobium, as was also found in the wild sampling study. Another species, P. paniculata, remains ambiguous in terms of symbiont preference. Unfortunately we could not investigate this topic due to the unavailability of seeds. In the case of Stryphnodendron species, trapping assays on soils recovered from the rhizosphere of two trees (identified as St. adstringens) trapped only Bradyrhizobium species. The species St. pulcherrimum species is nodulated by strains close to R. tropici but also by B. nodosa in French Guiana (Christine Le Roux, unpublished data), indicating that this genus is not only nodulated by α-rhizobia. The Stryphnodendron genus thus warrants a more focused study in order to characterize its symbionts.

Symbiosis Genes in Burkholderia from Mimosoid Legumes are Monophyletic and have been Transferred both Vertically and Horizontally Among Bacterial Species

The Burkholderia strains from Mimosa and from the other Piptadenia Group species are monophyletic in terms of their nodC and nifH phylogenies, indicating a common ancestor at the origin of their interaction with members of the tribe Mimoseae. The comparison of symbiotic and neutral markers revealed that vertical gene transfer is the main process for the dissemination of symbiotic genes within Burkholderia species. This pattern has been already observed for Mimosa symbionts [11], [12], and has allowed for the description of the symbiotic nature of Mimosa-nodulating burkholderias as being “ancient”, as it has been tentatively dated at c. 50 MY ago [11]. However, we found also that two species of Burkholderia (B. phenoliruptrix and B.sabiae) harbor strains with different nodC alleles, which were either inherited by vertical transfer (nodC3/BSa-2, nodC4/BPL-2), or were acquired by horizontal gene transfer from another Burkholderia clade (nodC1/BSa-1, nodC3/BPL-1). Horizontal gene transfer thus also played a role in the co-adaptation between Burkholderia species and their legume hosts.

Host Specificity between Burkholderia Genotypes and Piptadenia Group Species is not Strong

Although gene phylogenies have informed us about the ancient character of the symbiosis between burkholderias and Piptadenias, host specificity between the different partners is not obvious. Host plants are dispersed all over the Burkholderia phylogenies (Figure 2A,B), with no specificity pattern, except for that between BSP3 and P. gonoacantha. We tried to assess the specificity of the interactions between legume and rhizobial partners using co-inoculation experiments, and detected i) a high affinity of most tested plants towards Burkholderia genotypes, ii) promiscuous strains with high competitivity for nodulation (B. nodosa, B. phenoliruptrix), as well as some trends towards specificity (e.g. P. gonoacantha/BSP3, Mi. foetidus/Bradyrhizobium). Given the strong site sampling effect detected in the sampling analyses (Figure 3), and the absence of strict host specificity, it is realistic to hypothesize that most Piptadenia Group and Burkholderia species interact with a relaxed host specificity, and that plant selection targets nodulation genes from the Burkholderia nodC monophyletic clade or from diverse α-rhizobia when no symbiotic Burkholderia are present, and there are few exclusions in order to maximize their chances of finding a compatible partner. The environmental (especially soil) conditions would thus play a crucial role in the survival and biogeography of the symbionts, and could be the origin of the different diversity patterns observed in our results. Several parameters have already been found to affect Burkholderia symbionts diversity and/or competition, such as soil pH [12], [24], [16], altitude [11], or nitrogen sources [28].


In this study we extended the symbiotic interaction with Burkholderia from Mimosa to the wider Piptadenia Group in the tribe Mimoseae. Given the ancient and diverse character of this interaction, symbiosis with Burkholderia species might be present in other genera of this tribe, and possibly even in other mimosoid tribes, such as the Ingeae, as Barrett and Parker [63] have isolated Burkholderia strains from a member of this tribe (Abarema macradenia). The fact that B. phymatum STM815 is able to nodulate several legumes in the subfamily Mimosoideae (including species in Leucaena, Prosopis, and Acacia) give clues on the possible extent of the host range of these symbionts [7]. Symbiosis with Burkholderia could thus be much more common and ecologically significant than anticipated, encompassing several tribes in the Mimosoideae.

Given the relaxed specificity between Piptadenia Group species and their symbionts, and the diversification of Burkholderia into many species, larger samplings of each species in relation to soils and environmental parameters are required in order to get a clearer picture of the diversity and phylogeography of β-rhizobia, and how they have co-evolved with species in the Mimosoideae.

Supporting Information

Figure S1.

Sampling sites of nodules, soil and plant material.


Figure S2.

Section of M. pudica and P. gonoacantha nodules induced by different Burkholderia species. Legend: sections (40 micrometer-deep) of M. pudica nodules at 21 days post-inoculation (A to C) induced by B. sabiae STM7373 (A), B. phenoliruptrix STM7317 (B), or Burkholderia sp. 3 STM7296 (C). Sections of P. gonoacantha nodules at 60 dpi (D to E) induced by Burkholderia sp. 3 STM7296 (D) or Burkholderia sp. 1 JPY565 (E). Scale bar: 500 micrometer on all but B (1000 µm).


Figure S3.

Phylogenies of neutral and symbiotic markers in alpha-rhizobia from the Piptadenia group. The phylogeny of neutral markers (A) is based on a 16 S-recA partition and was built by a Bayesian analysis described in Figure 2 legend, while the symbiotic gene nodC phylogeny (B) was built by Maximum Likelihood with 1000 bootstraps replicates. See Figure 2 legend and Table S3 for sequence accession numbers.


Figure S4.

Comparison of phylogenies of neutral markers in rhizobia from the Piptadenia group. Phylogenies of 16 S rDNA (A) and recA (B) were built by Neighbor Joining from a distance matrix corrected by the Kimura 2 method, and with 1000 bootstraps replicates). See Figure 2 legend for abbreviations and Table S3 for accession numbers.


Figure S5.

Phylogeny of the nifH gene in alpha and beta-rhizobia from the Piptadenia group of legumes. The phylogeny was built by ML using a GTR model with 1000 bootstraps replicates. See Figure 2 legend for abbreviations and Table S3 for accession numbers.


Table S1.

Sampling sites characteristics and date of nodule collection. Legend: £ pH range indicated for the region when soil was not harvested for pH determination. ND: Not Determined.


Table S2.

Primers used for PCR amplification and sequencing.


Table S3.

Accession numbers of strains from this study and reference strains. Abbreviations: B. : Burkholderia, Br : Bradyrhizobium, R. : Rhizobium, C: Cupriavidus, Pseudo.: Pseudomonas, %: T at the end of the strain name indicates the type strain of a species.


Table S4.

Nodulation data of representative strains inoculated on Mimosa pudica (at 21 days post-inoculation-dpi) and Piptadenia gonoacantha (60 dpi). Legend: £: Efficiency of symbiosis was estimated as effective or ineffective according to plant development and presence of leghaemoglobin in nodules. %: Letters refer to Figure S2 pictures showing nodules sections. NT: Not Tested, ND: Not Determined. Scale bar on all nodules pictures is 1000 mm. BPL: Burkholderia phenoliruptrix, BSa: B. sabiae, BSP1-3: Burkholderia sp. 1 to 3, BN: B. nodosa, BD: B. diazotrophica.



This work is dedicated to the memory of Robson Dumas Ribeiro, botanist at the Botanical garden of Rio de Janeiro. We would like to thank Robson Dumas Ribeiro for its invaluable help on spotting and sampling Piptadenia trees. Collection of material was authorized by IBAMA No. 058/2006.

Author Contributions

Conceived and designed the experiments: CB SMF EG EKJ YP LM. Performed the experiments: CB SMF JMFdS PT MS CC YP LM. Analyzed the data: CB SMF JMFdS EKJ YP LM. Contributed reagents/materials/analysis tools: SMF JMFdS EG PT MS EKJ YP LM. Wrote the paper: CB SMF EKJ YP LM.


  1. 1. Lewis G, Schrire B, Mackinder B, Lock M (2005) Legumes of the World. Kew, Royal Botanic Gardens: Kew Publishing.
  2. 2. Maynaud G, Willems A, Soussou S, Vidal C, Mauré L, et al. (2012) Molecular and phenotypic characterization of strains nodulating Anthyllis vulneraria in mine tailings, and proposal of Aminobacter anthyllidis sp. nov., the first definition of Aminobacter as legume-nodulating bacteria. Systematic and Applied Microbiology 35: 65–72.
  3. 3. Ardley J, Parker M, De Meyer S, Trengove R, O’Hara G, et al. (2012) Strains of Gram-negative, rod-shaped, non-spore-forming bacteria were isolated from nitrogen-fixing nodules of the native legumes Listia angolensis (from Zambia) and Lupinus texensis (from Texas, USA). International Journal of Systematic and Evolutionary Microbiology 62: 2579–2588.
  4. 4. Moulin L, Munive a, Dreyfus B, Boivin-Masson C (2001) Nodulation of legumes by members of the beta-subclass of Proteobacteria. Nature 411: 948–950.
  5. 5. Chen WM, Laevens S, Lee TM, Coenye T, De Vos P, et al. (2001) Ralstonia taiwanensis sp. nov., isolated from root nodules of Mimosa species and sputum of a cystic fibrosis patient. International Journal of Systematic and Evolutionary Microbiology 51: 1729–1735.
  6. 6. Cummings SP, Gyaneshwar P, Vinuesa P, Farruggia FT, Andrews M, et al. (2009) Nodulation of Sesbania species by Rhizobium (Agrobacterium) strain IRBG74 and other rhizobia. Environmental Microbiology 11: 2510–2525.
  7. 7. Gyaneshwar P, Hirsch AM, Moulin L, Chen W-M, Elliott GN, et al. (2011) Legume-nodulating betaproteobacteria: diversity, host range, and future prospects. Molecular Plant-Microbe Interactions 24: 1276–1288.
  8. 8. Chen W, Moulin L, Bontemps C, Vandamme P (2003) Legume Symbiotic Nitrogen Fixation by beta-Proteobacteria Is Widespread in Nature. Journal of Bacteriology 185: 7266–7272.
  9. 9. Compant S, Nowak J, Coenye T, Clément C, Ait Barka E (2008) Diversity and occurrence of Burkholderia spp. in the natural environment. FEMS Microbiology Reviews 32: 607–626.
  10. 10. Suárez-Moreno ZR, Caballero-Mellado J, Coutinho BG, Mendonça-Previato L, James EK, et al. (2012) Common features of environmental and potentially beneficial plant-associated Burkholderia. Microbial Ecology 63: 249–266.
  11. 11. Bontemps C, Elliott GN, Simon MF, Dos Reis Júnior FB, Gross E, et al. (2010) Burkholderia species are ancient symbionts of legumes. Molecular Ecology 19: 44–52.
  12. 12. Mishra RPN, Tisseyre P, Melkonian R, Chaintreuil C, Miché L, et al. (2012) Genetic diversity of Mimosa pudica rhizobial symbionts in soils of French Guiana: Investigating the origin and diversity of Burkholderia phymatum and other beta-rhizobia. FEMS Microbiology Ecology 79: 487–503.
  13. 13. Chen W-M, James EK, Chou J-H, Sheu S-Y, Yang S-Z, et al. (2005) Beta-rhizobia from Mimosa pigra, a newly discovered invasive plant in Taiwan. The New phytologist 168: 661–675.
  14. 14. Chen W, De Faria SM, Pitard RM, Simo JL, Prescott AR, et al. (2005) Proof that Burkholderia Strains Form Effective Symbioses with Legumes: a Study of Novel Mimosa-Nodulating Strains from South America. Applied and Environmental Microbiology 71: 7461–7471.
  15. 15. Chen W-M, James EK, Coenye T, Chou J-H, Barrios E, et al. (2006) Burkholderia mimosarum sp. nov., isolated from root nodules of Mimosa spp. from Taiwan and South America. International Journal of Systematic and Evolutionary Microbiology 56: 1847–1851.
  16. 16. Liu X, Wei S, Wang F, James EK, Guo X, et al. (2012) Burkholderia and Cupriavidus spp. are the preferred symbionts of Mimosa spp. in southern China. FEMS Microbiology Ecology 80: 417–426.
  17. 17. Elliott GN, Chen W-M, Chou J-H, Wang H-C, Sheu S-Y, et al. (2007) Burkholderia phymatum is a highly effective nitrogen-fixing symbiont of Mimosa spp. and fixes nitrogen ex planta. The New Phytologist 173: 168–180.
  18. 18. Chen W-M, De Faria SM, James EK, Elliott GN, Lin K-Y, et al. (2007) Burkholderia nodosa sp. nov., isolated from root nodules of the woody Brazilian legumes Mimosa bimucronata and Mimosa scabrella. International Journal of Systematic and Evolutionary Microbiology 57: 1055–1059.
  19. 19. Chen W-M, De Faria SM, Chou J-H, James EK, Elliott GN, et al. (2008) Burkholderia sabiae sp. nov., isolated from root nodules of Mimosa caesalpiniifolia. International Journal of Systematic and Evolutionary Microbiology 58: 2174–2179.
  20. 20. Sheu S-Y, Chou J-H, Bontemps C, Elliott GN, Gross E, et al. (2012) Burkholderia symbiotica sp. nov., isolated from root nodules of Mimosa spp. native to North East Brazil. International Journal of Systematic and Evolutionary Microbiology 62: 2272–2278.
  21. 21. Sheu S, Chou J, Bontemps C, Elliott G, Gross E, et al. (2013) Burkholderia diazotrophica sp. nov., isolated from root nodules of Mimosa spp. International Journal of Systematic and Evolutionary Microbiology 63: 435–41.
  22. 22. Da Silva K, Florentino LA, Da Silva KB, De Brandt E, Vandamme P, et al. (2012) Cupriavidus necator isolates are able to fix nitrogen in symbiosis with different legume species. Systematic and Applied Microbiology 35: 175–182.
  23. 23. Elliott GN, Chen W-M, Bontemps C, Chou J-H, Young JPW, et al. (2007) Nodulation of Cyclopia spp. (Leguminosae, Papilionoideae) by Burkholderia tuberum. Annals of Botany 100: 1403–1411.
  24. 24. Garau G, Yates RJ, Deiana P, Howieson JG (2009) Novel strains of nodulating Burkholderia have a role in nitrogen fixation with papilionoid herbaceous legumes adapted to acid, infertile soils. Soil Biology and Biochemistry 41: 125–134.
  25. 25. Dos Reis FB, Simon MF, Gross E, Boddey RM, Elliott GN, et al. (2010) Nodulation and nitrogen fixation by Mimosa spp. in the Cerrado and Caatinga biomes of Brazil. The New Phytologist 186: 934–946.
  26. 26. Simon MF, Grether R, De Queiroz LP, Särkinen TE, Dutra VF, et al. (2011) The evolutionary history of Mimosa (Leguminosae): toward a phylogeny of the sensitive plants. American Journal of Botany 98: 1201–1221.
  27. 27. Amadou C, Pascal G, Mangenot S, Glew M, Bontemps C, et al. (2008) Genome sequence of the beta-rhizobium Cupriavidus taiwanensis and comparative genomics of rhizobia. Genome Research 18: 1472–1483.
  28. 28. Elliott GN, Chou J-H, Chen W-M, Bloemberg G V, Bontemps C, et al. (2009) Burkholderia spp. are the most competitive symbionts of Mimosa, particularly under N-limited conditions. Environmental Microbiology 11: 762–778.
  29. 29. Wang E, Rogel M, García-de los Santos A, Martínez-Romero J, Cevallos M, et al. (1999) Rhizobium etli bv. mimosae, a novel biovar isolated from Mimosa affinis. International journal of systematic bacteriology 49 Pt 4: 1479–1491.
  30. 30. Jobson RW, Luckow M (2007) Phylogenetic Study of the Genus Piptadenia (Mimosoideae: Leguminosae ) using Plastid trnL-F and trnK/matK Sequence Data. Systematic Botany 32: 569–575.
  31. 31. Lewis G (1991) Five new taxa of Piptadenia (Leguminosae: Mimosoideae) from Brazil. Kew Bull 46: 159–168.
  32. 32. Scotti MR, Corrêa EJA (2004) Original article Growth and litter decomposition of woody species inoculated with rhizobia and arbuscular mycorrhizal fungi in Semiarid Brazil. 61: 87–95.
  33. 33. Pagano MC, Cabello MN, Bellote AF, Sá NM, Scotti MR (2008) Intercropping system of tropical leguminous species and Eucalyptus camaldulensis, inoculated with rhizobia and/or mycorrhizal fungi in semiarid Brazil. Agroforestry Systems 74: 231–242.
  34. 34. De Souza C, Haas AS, Von Poser GL, Schapoval EES, Elisabetsky E (2004) Ethnopharmacological studies of antimicrobial remedies in the south of Brazil. Journal of Ethnopharmacology 90: 87–95.
  35. 35. Schmidt C a, Murillo R, Bruhn T, Bringmann G, Goettert M, et al. (2010) Catechin derivatives from Parapiptadenia rigida with in vitro wound-healing properties. Journal of natural products 73: 2035–2041.
  36. 36. Correa M, De Azeredo Pena L (1984) Dicionário das plantas úteis do Brasil e das exóticas cultivadas, vol.4. Rio de Janeiro: Ministério da Agricultura, Instituto Brasileiro de Desenvolvimento Florestal.
  37. 37. De Faria S, Franco A, Jesus R De, Menandro MDS, Baitello JB, et al. (1984) New Nodulating Legume South-East. New Phytologist 98: 317–328.
  38. 38. De Faria SM, Diedhiou AG, De Lima HC, Ribeiro RD, Galiana A, et al. (2010) Evaluating the nodulation status of leguminous species from the Amazonian forest of Brazil. Journal of experimental botany 61: 3119–3127.
  39. 39. De Souza Moreira F, Da Silva M, De Faria S (1992) Occurrence of nodulation in legume species in the Amazon region of Brazil. New Phytologist 121: 563–570.
  40. 40. Gross E (2002) Nodule Ultrastructure and Initial Growth of Anadenanthera peregrina (L.) Speg. var. falcata (Benth.) Altschul Plants Infected with Rhizobia. Annals of Botany 90: 175–183.
  41. 41. Jesus E da C, Schiavo J, De Faria SM (2005) Dependence on arbuscular mycorrhizal fungi for nodulation and growth of tropical woody legume species. Rev Árvore 29: 545–552.
  42. 42. Zurdo-Piñeiro JL, Velázquez E, Lorite MJ, Brelles-Mariño G, Schröder EC, et al. (2004) Identification of fast-growing rhizobia nodulating tropical legumes from Puerto Rico as Rhizobium gallicum and Rhizobium tropici. Systematic and applied microbiology 27: 469–477.
  43. 43. Freitas ADS, Sampaio EVSB, Santos CERS, Fernandes AR (2010) Biological nitrogen fixation in tree legumes of the Brazilian semi-arid caatinga. Journal of Arid Environments 74: 344–349.
  44. 44. Taulé C, Zabaleta M, Mareque C, Platero R, Sanjurjo L, et al. (2012) New Betaproteobacterial Rhizobium Strains Able To Efficiently Nodulate Parapiptadenia rigida (Benth.) Brenan. Applied and Environmental Microbiology 78: 1692–1700.
  45. 45. Vincent J (1970) A manual for the pratical study of root-nodule bacteria. I.B.P. Han. Ltd, Oxford: Blackwell Scientific Publications.
  46. 46. Franco AA, De Faria S (1997) The Contribution of N2-Fixing Tree Legumes To Land Reclamation And Sustainability In The Tropics. Soil Biology and Biochemistry 29: 897–903.
  47. 47. Moulin L, Béna G, Boivin-Masson C, Stępkowski T (2004) Phylogenetic analyses of symbiotic nodulation genes support vertical and lateral gene co-transfer within the Bradyrhizobium genus. Molecular Phylogenetics and Evolution 30: 720–732.
  48. 48. Stepkowski T, Moulin L, Krzyzanska A, McInnes A, Law IJ, et al. (2005) European Origin of Bradyrhizobium Populations Infecting Lupins and Serradella in Soils of Western Australia and South Africa. Applied and Environmental Microbiology 71: 7041–7052.
  49. 49. Gaunt MW, Turner SL, Rigottier-Gois L, Lloyd-Macgilp S a, Young JP (2001) Phylogenies of atpD and recA support the small subunit rRNA-based classification of rhizobia. International journal of systematic and evolutionary microbiology 51: 2037–2048.
  50. 50. Laguerre G, Nour SM, Macheret V, Sanjuan J, Drouin P, et al. (2001) Classification of rhizobia based on nodC and nifH gene analysis reveals a close phylogenetic relationship among Phaseolus vulgaris symbionts. Microbiology (Reading, England) 147: 981–993.
  51. 51. Sarita S, Sharma PK, Priefer UB, Prell J (2005) Direct amplification of rhizobial nodC sequences from soil total DNA and comparison to nodC diversity of root nodule isolates. FEMS Microbiology Ecology 54: 1–11.
  52. 52. Edgar R (2004) MUSCLE: multiple sequence alignment with high accuracy and high throughput. Nucleic Acids Research 32: 1792–1797.
  53. 53. Nicholas K, Nicholas B (n.d.) Genedoc: a tool for editing and annotating multiple sequence alignments. Distributed by the authors.
  54. 54. Schloss PD, Westcott SL, Ryabin T, Hall JR, Hartmann M, et al. (2009) Introducing mothur: open-source, platform-independent, community-supported software for describing and comparing microbial communities. Applied and Environmental Microbiology 75: 7537–7541.
  55. 55. Tamura K, Peterson D, Peterson N, Stecher G, Nei M, et al. (2011) MEGA5: molecular evolutionary genetics analysis using maximum likelihood, evolutionary distance, and maximum parsimony methods. Molecular biology and evolution 28: 2731–2739.
  56. 56. Swofford D (1998) PAUP*. Phylogenetic analysis using parsimony (*and Other Methods): Version 4.
  57. 57. Ronquist F, Huelsenbeck JP (2003) MrBayes 3: Bayesian phylogenetic inference under mixed models. Bioinformatics 19: 1572–1574.
  58. 58. Sprent JI (2009) Legume Nodulation: a global perspective. Chichester UK: Wiley-Blackwells.
  59. 59. Payne GW, Vandamme P, Morgan SH, Lipuma JJ, Coenye T, et al. (2005) Development of a recA Gene-Based Identification Approach for the Entire Burkholderia Genus. 71: 3917–3927.
  60. 60. López-López A, Rogel-Hernández M a, Barois I, Ortiz Ceballos AI, Martínez J, et al. (2012) Rhizobium grahamii sp. nov. from Dalea leporina, Leucaena leucocephala, Clitoria ternatea nodules, and Rhizobium mesoamericanum sp. nov. from Phaseolus vulgaris, siratro, cowpea and Mimosa pudica nodules. International Journal of Systematic and Evolutionary Microbiology 62: 2264–2271.
  61. 61. Klonowska A, Chaintreuil C, Tisseyre P, Miché L, Melkonian R, et al. (2012) Biodiversity of Mimosa pudica rhizobial symbionts (Cupriavidus taiwanensis, Rhizobium mesoamericanum) in New Caledonia and their adaptation to heavy metal-rich soils. FEMS Microbiology Ecology 81: 618–635.
  62. 62. Coenye T, Henry D, Speert DP, Vandamme P (2004) Burkholderia phenoliruptrix sp. nov., to Accommodate the Strain AC1100. Systematic and Applied Microbiology 27: 623–627.
  63. 63. Barrett CF, Parker M a (2005) Prevalence of Burkholderia sp. nodule symbionts on four mimosoid legumes from Barro Colorado Island, Panama. Systematic and Applied Microbiology 28: 57–65.