Plant Microbiology

Distribution and Phylogeny of Microsymbionts Associated with Cowpea (Vigna unguiculata) Nodulation in Three Agroecological Regions of Mozambique

Ifeoma N. Chidebe, Sanjay K. Jaiswal, Felix D. Dakora
Volker Müller, Editor
DOI: 10.1128/AEM.01712-17

ABSTRACT

Cowpea derives most of its N nutrition from biological nitrogen fixation (BNF) via symbiotic bacteroids in root nodules. In Sub-Saharan Africa, the diversity and biogeographic distribution of bacterial microsymbionts nodulating cowpea and other indigenous legumes are not well understood, though needed for increased legume production. The aim of this study was to describe the distribution and phylogenies of rhizobia at different agroecological regions of Mozambique using PCR of the BOX element (BOX-PCR), restriction fragment length polymorphism of the internal transcribed spacer (ITS-RFLP), and sequence analysis of ribosomal, symbiotic, and housekeeping genes. A total of 122 microsymbionts isolated from two cowpea varieties (IT-1263 and IT-18) grouped into 17 clades within the BOX-PCR dendrogram. The PCR-ITS analysis yielded 17 ITS types for the bacterial isolates, while ITS-RFLP analysis placed all test isolates in six distinct clusters (I to VI). BLASTn sequence analysis of 16S rRNA and four housekeeping genes (glnII, gyrB, recA, and rpoB) showed their alignment with Rhizobium and Bradyrhizobium species. The results revealed a group of highly diverse and adapted cowpea-nodulating microsymbionts which included Bradyrhizobium pachyrhizi, Bradyrhizobium arachidis, Bradyrhizobium yuanmingense, and a novel Bradyrhizobium sp., as well as Rhizobium tropici, Rhizobium pusense, and Neorhizobium galegae in Mozambican soils. Discordances observed in single-gene phylogenies could be attributed to horizontal gene transfer and/or subsequent recombinations of the genes. Natural deletion of 60 bp of the gyrB region was observed in isolate TUTVU7; however, this deletion effect on DNA gyrase function still needs to be confirmed. The inconsistency of nifH with core gene phylogenies suggested differences in the evolutionary history of both chromosomal and symbiotic genes.

IMPORTANCE A diverse group of both Bradyrhizobium and Rhizobium species responsible for cowpea nodulation in Mozambique was found in this study. Future studies could prove useful in evaluating these bacterial isolates for symbiotic efficiency and strain competitiveness in Mozambican soils.

INTRODUCTION

For millennia, humans have utilized legumes as a source of food, animal fodder, traditional medicine, shelter, and fuel (1). In Africa's diverse cultures, grain legumes constitute an integral component of cropping systems, especially in the rural communities. In Mozambique, for example, cowpea is the most important food grain legume, grown extensively on smallholder fields and estimated at nearly four million hectares in land area. The high protein content of cowpea grain and leaves (2) can augment the low-protein diets of rural Mozambican households, with potential to eliminate protein calorie malnutrition. Cowpea grain also contains 57% carbohydrates (2), essential amino acids, dietary fiber, and abundant minerals (3) and thus contributes to a balanced diet.

In addition to their nutritional importance, legumes are very highly valued in the agroecosystem as they have the most efficient biological nitrogen fixation (BNF) system known in nature (4). BNF in legumes occurs through a symbiotic partnership between the host plant and soil bacteria collectively known as rhizobia, which provide biologically fixed atmospheric N to the legume in exchange for plant photosynthate. The distinctive feature of this partnership is the presence of specialized symbiotic organs, or nodules, which are developed on the roots and occasionally stems of legumes following a series of molecular exchanges and morphological modifications in the two partners (5).

Legume fixed N can meet more than half of the total N needs of a legume (6). As a result, plant growth and productivity are often less affected by soil N deficiency (7). Through N contribution in cropping systems, the legume/rhizobium symbiosis has the potential to promote and sustain agricultural productivity, particularly in the low-input systems of Sub-Saharan Africa (8).

Although they all are capable of forming root or stem nodules, rhizobia can be phylogenetically and metabolically very diverse (4). Currently, there are about 15 genera of symbiotic N2-fixing bacteria belonging to the phylum Proteobacteria in the alpha, beta, and gamma classes. Representative N2 fixers are found in the genera Agrobacterium, Allorhizobium, Azorhizobium, Bradyrhizobium, Burkholderia, Cupriavidus, Devosia, Herbaspirrilum, Mesorhizobium, Methylobacterium, Ochrobactrum, Phyllobacterium, Rhizobium, Shinella, and Sinorhizobium (9). To date, more than 113 nodulating species have been identified within these genera. However, knowledge gaps still exist regarding the genetic diversity and biogeographic distribution of the N2-fixing microsymbionts of many legumes (10), especially in Sub-Saharan Africa, where the bacterial symbionts have not been studied for many indigenous legumes. This therefore emphasizes the need to assess the rhizobial genetic diversity within the region and determine the taxonomic affiliations of the bacteria nodulating native legumes.

A determination of the taxonomy and phylogeny of rhizobia is key to improving legume productivity via BNF in diverse environments. Not only has this increased our understanding of the diversity and genetic relatedness of rhizobia, but it also has significantly improved rhizobial classification through (i) the redefinition of species lineages (10, 11), (ii) the identification of rhizobia from more genera of alpha- and betaproteobacteria (12–14), and (iii) the discovery of novel species (15). Such studies are important for generating a genetic resource base from which highly adapted and efficient rhizobial strains are selected for inoculant production (15). Therefore, well-tested, quick methods are required for differentiating rhizobial isolates (16). PCR of the BOX element (BOX-PCR) and PCR-restriction fragment length polymorphism (RFLP) analyses have become powerful tools for detecting diversity among rhizobial isolates at the genome level (16, 17).

Sequencing of the 16S rRNA gene, which is functionally conserved and universally distributed, has become one of the most important tools for defining kingdom and genera but not species (18, 19). To overcome this, the 16S rRNA phylogenetic analysis is usually complemented with data from more rapidly evolving loci, such as the protein-coding housekeeping genes. These genes have a much higher level of sequence divergence, especially in relation to the 16S rRNA gene, but are sufficiently well conserved to retain genetic information for characterizing bacteria at the intra- and interspecific levels (20). Previous studies of cowpea rhizobia have reported nodulation by different species in the genera Bradyrhizobium, Rhizobium, Sinorhizobium, Ralstonia, Achromobacter, and Microvirga (7, 15, 21–24). Clearly, this suggests that the phylogeny of rhizobia nodulating cowpea is still unclear and requires further study (22, 25).

Cowpea is cultivated in all the 10 agroecological zones (AEZs) of Mozambique and contributes significantly to protein calorie requirements in rural households. However, to date, little information exists on the diversity and phylogeny of cowpea-nodulating bacteria in many African countries, including Mozambique. The aim of this study was to determine the diversity and phylogenetic relationships of cowpea rhizobia isolated from experimental sites located in three different agroecological regions of Mozambique using BOX-PCR and RFLP of the internal transcribed spacer (ITS-RFLP) analyses and sequence analysis of 16S rRNA, nifH, and housekeeping genes (recA, glnII, gyrB, and recA).

RESULTS

Bacterial isolation from root nodules.A total of 122 bacterial isolates were obtained from root nodules of both cowpea varieties (IT-1263 and IT-18) planted at Muriaze, Ruace, and Sussundenga in Mozambique. Of the 122 isolates, 39 originated from Muriaze, 42 from Ruace and 41 from Sussundenga.

PCR amplification of the repetitive (rep, i.e., BOX) genomic region.PCR amplification of the BOX region of the genomic DNA from each isolate resulted in distinctive banding patterns, ranging from one to six fragments per DNA profile. Most of the bands were very clear; however, a few faint bands were also observed. The cluster analysis (unweighted pair group method using average linkages [UPGMA] algorithm and Jaccard's similarity coefficient) differentiated the bacterial isolates into 17 clades at 50% Jaccard's similarity coefficient (Fig. 1). The number of isolates in each clade ranged from 2 to 17, with six isolates (TUTVU31, TUTVU-5, TUTVU-4, TUTVU-6, TUTVU-8, and TUTVU63) showing high genetic distinctiveness from the others at 0% Jaccard's similarity coefficient. Conversely, there was 100% similarity coefficient for 68 bacterial isolates (55%), and these were distributed within 9 of the 17 clades. Interestingly, among these, only isolate TUTVU26 showed 100% Jaccard's similarity coefficient with the commercial inoculant BIOFIX (TUTVC) (Fig. 1). There was no clear distinction based on isolate site of origin, as the rhizobial isolates from the three study sites were distributed across all the clades, except for clade XII, which consisted of bacterial isolates exclusively from the Ruace site (Table 1; Fig. 1). Isolates from Muriaze were distributed across 11 different clades, while those from Ruace and Sussundenga were found in 12 and 13 different clades, respectively.

FIG 1
FIG 1

Dendrogram based on BOX-PCR fingerprints of cowpea nodule isolates.

View this table:
TABLE 1

Origin, BOX-PCR, ITS (16S-23S rRNA) characterization, nodulation status and colony morphology of symbiotic bacteria isolated from the root nodules of cowpea from Mozambiquea

The BOX-PCR clustering data were used to analyze the diversity indices of the bacterial isolates from the three study sites. The types of bacterial isolates varied across sites (Fig. 1). The Shannon diversity index (H′) was highest (1.12) at Muriaze and lowest (0.09) at Sussundenga. The values for the Margalef index (R1), which is a measure of species richness, were similar across the three experimental sites, at 7.55, 8.64, and 8.07 for Muriaze, Ruace, and Sussundenga, respectively. The values for the Pielou index (E1), which measures species evenness, were similar at the Muriaze and Ruace experimental sites (1.01 and 0.92) but lowest (0.08) at Sussundenga.

Ninety-nine out of the 122 isolates (81%) effectively induced nodule formation in cowpea variety IT-18, the homologous host. Of this number, 28%, 32%, and 39% originated from Muriaze, Ruace, and Sussundenga, respectively.

The 99 effective rhizobial isolates differed significantly in morphological characteristics, with 94% of the isolates forming visible colonies on yeast mannitol agar (YMA) plates in 5 to10 days and only 6% forming visible colonies in 2 to 4 days. About 50% of the colonies were milky, 25% translucent, 37% gummy, and 21% secreting exopolysaccharide. Colony size varied from 0.8 to 2.4 mm in diameter.

PCR-RFLP analysis of the 16S-23S rRNA genomic region.The PCR-amplified products of the 16S-23S rRNA ITS region of rhizobial DNA showed single polymorphic bands, ranging from 258 to 1,350 bp in size, except for isolates TUTVU17, TUTVU54, TUTVU58, TUTVU65, TUTVU66, TUTVU98, and TUTVU99, which produced two fragments each (Table 1). The commercial inoculant strain (BIOFIX) also produced double bands and was distinct from the test isolates. Based on the differences in band size, the rhizobial isolates were grouped into 17 distinctive ITS types, numbered I to XVII (Table 1). The majority of rhizobial isolates from the three experimental sites grouped within ITS type IX with a band size of 1,000 bp (Table 1). Rhizobial isolates TUTVU37 and TUTVU49, however, failed to amplify.

Gel electrophoresis of products from the restriction endonuclease-digested PCR-amplified ITS genomic region indicated the presence of different enzyme recognition sites, as distinct polymorphic bands were observed. From the binary scoring, the HaeII restriction endonuclease enzyme was the most discriminatory, yielding 23 distinct fingerprint profiles (A to W), while the HindIII and HinfI restriction endonuclease enzymes yielded 19 (A to S) and 21 (A to U) distinct fingerprint profiles, respectively (Table 1).

The combined ITS-RFLP cluster analysis revealed six distinct ITS-RFLP clusters (I to VI) at 50% Jaccard's similarity coefficient (Fig. 2). Five of the rhizobial isolates (namely, TUTVU31, TUTVU2, TUTVU99, TUTVU33, and TUTVU54) were very distinct, as they showed no similarity with the other test isolates at 0 to 0.1% Jaccard's similarity coefficient. The commercial inoculant (BIOFIX) was highly distinct, at 0% Jaccard's similarity coefficient. Clade II had 57% of the rhizobial isolates, of which 51% originated from Sussundenga site. Seven groups of rhizobial isolates within clades I, II, and V, which comprised between 2 and 28 rhizobial isolates, each showed 100% Jaccard's similarity coefficient. Isolates from all the three study sites were represented in four of the seven groups. Clades III and VI each comprised two and four test isolates from the Ruace site only, while clade IV comprised four test isolates exclusively from Sussundenga. Based on cluster analysis, most of the isolates from the different experimental sites showed a high level of similarity at the ITS genomic level.

FIG 2
FIG 2

Dendrogram generated from ITS-RFLP banding pattern of rhizobial isolates nodulating cowpea in Mozambique.

Representative isolates from each cluster of the ITS-RFLP dendrogram were selected for further phylogenetic study. Following PCR amplification, full-length amplicons (1.5 kb) of the 16S rRNA gene were obtained for all 34 test rhizobial isolates. However, 11 gene sequences of poor quality were excluded from the 16S rRNA study. The BLASTn sequence analysis of 23 test rhizobial isolates showed that 74% of the isolates had very high sequence similarities to Bradyrhizobium, while 26% had high gene sequence similarities with Rhizobium type strains. The phylogenetic analysis was therefore computed separately for each group of isolates (i.e., Bradyrhizobium and Rhizobium).

The Bradyrhizobium lineage.The 16S rRNA consensus sequence (534 bp) comprised 449 conserved, 85 variable, 14 parsimony informative, and 71 singleton sites (see Table S2 in the supplemental material). The maximum likelihood phylogeny of the 16S rRNA genes placed the test isolates in three distinct clades (I to III) (Fig. 3A). In clade I, isolates TUTVU87, TUTVU99, TUTVU86, TUTVU81, TUTVU70, TUTVU63, TUTVU55, TUTVU39, TUTVU36, TUTVU22, TUTVU21, and TUTVU11 grouped with the reference type strains of the B. elkanii group (i.e., B. elkanii and B. pachyrhizi) and showed 100% identical sequences, with 80% bootstrap support. Clade II comprised isolates TUTVU44, TUTVU1, TUTVU13, and TUTVU7 with 61% bootstrap support, and all of them had sequences identical to those of the type strains of B. kavangense and B. subterraneum, except for isolate TUTVU44, with 99.8% sequence identity. Isolate TUTVU5 shared identical sequences with the reference type strains of B. denitrificans, B. huanghuaihaisense, B. ingae, B. iriomotense, and B. stylosanthis in clade III, with 50% bootstrap support.

FIG 3FIG 3
FIG 3

(A) Maximum likelihood based phylogenetic relationships between test isolates of cowpea root nodules and reference Bradyrhizobium type strains (NCBI) based on 16S rRNA gene sequences. Bootstrap values (1,000 replicates) of ≥50% are indicated at the nodes. (B) Maximum likelihood-based phylogenetic relationships between test isolates of cowpea root nodules and Rhizobium reference type strains (NCBI) based on 16S rRNA partial gene sequences. Bootstrap values are indicated at the nodes.

The Rhizobium lineage.The 569-bp length of 16S rRNA consensus sequence contained 457 conserved, 112 variable, 72 parsimony, and 40 singleton sites (Table S2). The 16S rRNA maximum likelihood phylogeny placed the test isolates in three (I to III) distinct clades (Fig. 3B). In clade I, isolates TUTVU50 and TUTVU33 grouped together with the type strains of R. hainanense, R. multihospitium, and R. miluonense, with 95% bootstrap support and 99.2 to 99.8% sequence identity, while isolate TUTVU31 formed an outgroup of clade I. Clade II comprised isolate TUTVU68 and the type strains of R. etli, R. fabae, R. pisi, R. binae, R. sophoriradicis, R. mongolense, and R. vallis, with 100% sequence identity. In clade III, isolate TUTVU40 had a sequence identical to that of the type strain of R. pusense, with 100% bootstrap support, while isolate TUTVU67 formed an outgroup of clade III.

Sequence and phylogenetic analysis of housekeeping genes.The PCR-amplified products of the gyrB, rpoB, glnII, and recA housekeeping genes yielded single fragments of approximately 750, 500, 650, and 700 bp, respectively. As done for the 16S rRNA phylogenetic analysis, the sequences of housekeeping genes were compared with published type reference strain sequences using the BLASTn program of the NCBI (GenBank). These genes had high sequence similarities to the reference type strains of two distinct rhizobial genera (Bradyrhizobium and Rhizobium). The phylogenetic analyses were again done separately for the two groups of rhizobial strains (i.e., Bradyrhizobium and Rhizobium).

Bradyrhizobium lineage gene sequence analysis.Due to the poor quality of some sequences obtained, only 24 glnII, 25 gyrB, 21 rpoB, and 26 recA housekeeping gene sequences of the test isolates were subjected to single-gene glnII (244 bp), gyrB (358 bp), rpoB (226 bp), and recA (301 bp) phylogenetic analysis. The gyrB housekeeping gene had the highest number (112) of parsimony informative sites, while the glnII housekeeping gene had the lowest number (18) of singleton sites (Table S2).

The maximum likelihood phylogenies were generally congruent for three (glnII, gyrB, and rpoB) of the four housekeeping single-gene phylogenies, although there were some slight variations in the sequence identity of a few test isolates. In each tree topology, the test isolates grouped into four (I to IV) distinct clades (Fig. S1a, S2a, and S3a), with most of them clustering within similar clades across all three housekeeping genes. For example, clade I consisted of isolates TUTVU3, TUTVU11, TUTVU14, TUTVU22, TUTRU30, TUTVU36, TUTVU39, TUTVU47, TUTVU55, TUTVU59, TUTVU63, TUTVU66, TUTVU70, TUTVU77, TUTVU81, TUTVU86, TUTVU87, TUTVU92, and TUTVU99 in the four housekeeping gene phylogenies and had between 97.1 and 100% sequence identity to the type reference strains of B. pachyrhizi, B. elkanii, B. viridifuturi, and B. ferriligni. The only exception was isolate TUTVU66, which grouped with the type strain of B. arachidis, with 95% bootstrap support and 99.1% sequence identity in clade III of the glnII phylogeny, and isolate TUTVU21, which had a sequence identical to that of the type strain of B. elkanii in the rpoB gene but had a proximal relationship to the type strains of B. embrapense and B. viridifuturi, with 95.4% sequence identity and 57% bootstrap support in the glnII gene phylogeny.

In a similar manner, clade II, which comprised isolates TUTVU13, inoculant strain, and TUTVU44, showed a proximal relationship with the type strain of B. diazoefficiens, with 96.7 to 100% sequence identity in the gyrB and glnII phylogenies. But in the rpoB phylogeny, clade II comprised isolates TUTVU13, TUTVU1, and TUTVU44 together with the type strain of B. yuanmingense and shared 82% bootstrap support and 98.2 to 100% sequence identity. Isolate TUTVU1, however, showed discordance in the glnII phylogram. Isolates TUTVU54, TUTVU7, and TUTVU5 formed outgroups of clade II in the glnII phylogeny but not in the rpoB phylogeny, while isolate TUTVU5 grouped in clade III with the type strain of B. arachidis. Isolates TUTVU54 and TUTVU7 also grouped in clade IV with the type strain of B. liaoningense, with 52% and 62% bootstrap support, respectively. In the gyrB phylogeny, isolate TUTVU5 had 96.6% sequence identity with the type strain of B. stylosanthis, with 64% bootstrap support. Isolates TUTVU14, TUTVU39, TUTVU55, TUTVU63, and TUTVU87 were consistently grouped with the type strain of B. pachyrhizi in all three housekeeping gene phylogenies.

A discrepancy was, however, observed in the recA phylogeny (Fig. S4). For example, in clade II, isolates TUTVU7, TUTVU44, TUTVU13, and TUTVU54 had a proximal relationship with the type strain of B. subterraneum, while it showed a proximal relationship with the type strain of B. diazoefficiens in the gyrB and glnII phylogenies. Similarly, in clade III, isolate TUTVU63 grouped with the type strain of B. stylosanthis in this gene but also consistently showed a nearly identical relationship with the type strain of B. pachyrhizi in the other housekeeping gene phylogenies.

The Rhizobium lineage.The phylogenetic analysis was performed using sequences individually aligned with ClustalW for the glnII (358 bp), gyrB (577 bp), and rpoB (218 bp) housekeeping genes along with reference type strains obtained from the NCBI GenBank database.

The maximum likelihood phylogenetic analyses were congruent for the glnII, gyrB, and rpoB housekeeping genes (Fig. S1b, S2b, and S3b). For example, isolates TUTVU50 and TUTVU31 showed high genetic relatedness to type strains in the R. tropici group, with high bootstrap support. However, isolates TUTVU33 and TUTVU67 also clustered with the type strain of R. galegae, with 100% bootstrap support in the glnII and gyrB phylogenies, while in the rpoB phylogeny, isolate TUTVU33 grouped with the type strain of R. mesosinicum, with 86% bootstrap support. In contrast, isolate TUTVU40 showed genetic relatedness to Neorhizobium pusense in all three housekeeping gene phylogenies.

Interestingly, the gyrB nucleotide sequence of isolate TUTVU7 showed variation (gaps) in comparison to test and type bradyrhizobial strains. The 60-bp natural deletion was observed in the gyrB genomic region of isolate TUTVU7 (Fig. 4A). The annotation result suggested the occurrence of a deletion in the coding region (Fig. 4B).

FIG 4
FIG 4

Sequence alignment of amplicons of gyrB gene of test cowpea isolates. (A) Nucleotide sequence alignment showing 60-bp deletion regions; (B) amino acid sequence alignment regions showing 20-amino-acid deletion regions.

Concatenated gene phylogenies.A concatenated gene sequence analysis was performed to refine the phylogenetic positions of the test isolates within the Bradyrhizobium and Rhizobium lineages. A maximum likelihood phylogeny was constructed using concatenated aligned sequences of the test isolates and type reference strains common across the four housekeeping genes.

The Bradyrhizobium lineage.For the Bradyrhizobium lineage, 15 test isolates were included in the concatenated (glnII plus gyrB plus recA plus rpoB) gene sequence analysis. The consensus sequence of 1,128 bp comprised 648 conserved, 480 variable, 290 parsimony, and 190 singleton sites (Table S2). In the concatenated tree topology, the test isolates grouped into two distinct (I and II) clades (Fig. 5). Ten isolates (namely, TUTVU86, TUTVU21, TUTVU22, TUTVU70, TUTVU81, TUTVU11, TUTVU55, TUTVU39, TUTVU63, and TUTVU87) clustered in clade I, with 99% bootstrap support. In this clade, test isolates TUTVU21 and TUTVU22 had between 98.1 and 98.7% sequence identities with the type strain of B. pachyrhizi. In clade II, isolates TUTVU7, TUTVU5, TUTVU1, TUTVU13, and TUTVU44 grouped together without any type reference strains.

FIG 5
FIG 5

Maximum likelihood-based concatenated (glnII plus gyrB plus recA plus rpoB) phylogenetic tree of test Bradyrhizobium isolates and reference type sequences (GenBank). Bootstrap values are indicated at the nodes.

The Rhizobium lineage.For the Rhizobium lineage, concatenated housekeeping gene analysis was performed in two subsets (glnII plus gyrB) and (glnII plus gyrB plus rpoB) to account for inconsistencies in the number of test isolates in the individual phylogenetic trees (Fig. 6A and B). The consensus analysis of the glnII plus gyrB concatenated gene sequences of 976 bp consisted of 558 conserved and 418 variable sites, while the concatenated gene sequences (glnII plus gyrB plus rpoB) of 1,194 bp comprised 531 variable and 361 parsimony informative sites (Table S2).

FIG 6
FIG 6

(A) Maximum likelihood-based concatenated (glnII plus gyrB) phylogenetic tree of test Rhizobium isolates and reference type sequences (GenBank). Bootstrap values are indicated at the nodes. (B) Maximum likelihood-based concatenated (glnII plus gyrB plus rpoB) phylogenetic tree of test Rhizobium isolates and reference type sequences (GenBank). Bootstrap values are indicated at the nodes.

In the maximum likelihood phylogeny of both concatenated housekeeping gene subsets, isolates TUTVU50 and TUTVU31 were closely related (99.6% and 97.7% sequence identity) to the type strain of R. tropici, while isolates TUTVU33 and TUTVU67 both showed low sequence identities (94.7% and 95.2%) with the type strain of R. galegae, with 100% bootstrap support (Fig. 6). Isolate TUTVU40 in clade III had a 96.8% sequence similarity with the type strain of Neorhizobium pusense, with 100% bootstrap support in both concatenated housekeeping gene subsets.

Phylogenetic analysis of the nifH symbiotic gene sequences.The PCR-amplified products of the nifH gene yielded single polymorphic fragments 800 bp in length. In the maximum likelihood phylogeny, the test isolates grouped into eight distinct clades (I to VIII) (Fig. 7). Clades III and I comprised test isolates which previously clustered with the B. elkanii lineage in the housekeeping gene phylogenies. All the isolates in clade III, however, formed a monophyletic group, with no close relatedness to any known type strains. Clades II, IV, V, and VI each comprised isolates TUTVU77, TUTVU5, TUTVU7, and TUTVU13, which, respectively, had 99.5%, 97.9%, 99.1%, and 99.5% sequence similarities with the type strains of B. viridifuturi, B. arachidis, B. subterraneum, and B. yuanmingense. In clade VIII, isolate TUTVU99 had identical sequence similarity with reference type strains of B. japonicum, B. diazoefficiens, B. liaoningense, B. huanghuaihaiense, B. daqingense, B. ottawaense, and B. lupini, with 100% bootstrap support. However, test isolates TUTVU1, TUTVU44, TUTVU54, and TUTVU66 in clade VII grouped together but not with any known reference type strain.

FIG 7
FIG 7

Maximum likelihood-based phylogenetic relationships between test isolates of cowpea root nodules and reference Bradyrhizobium type strains (GenBank) based on nifH gene sequences. Bootstrap values are indicated at the nodes.

16S rRNA metagenomic analysis.A total of 127,013 reads were observed, and each sample recorded >10,000 reads. The observed reads were assigned to high taxonomic ranks. The analysis based on occurrence and relative abundance of different phyla suggested strong dissimilarities among bacterial communities in all the test rhizosphere soil samples. The results showed that at Sussundenga the rhizosphere of the two cowpea varieties attracted the highest number (18) of bacterial phyla, followed by Muriaze and then Ruace. We also found bacteria of unknown genera in the rhizosphere soils, and their numbers followed the same pattern as the identifiable phyla, in that these unidentified bacteria occurred in greater abundance in cowpea rhizosphere at Sussundenga (45.05 to 53.34%), followed by Muriaze and Ruace. However, of the betaproteobacteria found, the rhizosphere of IT-18 at Sussundenga and IT-1263 at Muriaze revealed the highest number of bradyrhizobial populations (data not shown).

DISCUSSION

The benefits of legume N2 fixation can potentially be better exploited by exploring new biogeographic regions for novel rhizobia and legume germplasm, both of which can lead to the discovery of novel symbioses and elite microsymbionts in support of agriculture. Cowpea (Vigna unguiculata [L.] Walp.) is an important food legume in Mozambique, but its rhizobial diversity and phylogeny are still little understood.

In this study, the diversity of root nodule bacteria isolated from cowpea was assessed using PCR-based fingerprinting techniques. The BOXA1R subunit of the BOX repetitive elements in the bacterial genome is highly conserved; therefore, PCR amplification analysis of this region can be used to differentiate bacterial isolates (26). Although the region is highly conserved, genetic distances between repetitive sequences can vary in unrelated organisms (26). The technique has therefore been successfully employed in diversity studies of rhizobia to identify interstrain variations (27, 28).

The 122 cowpea root-nodule bacteria used in this study were grouped into 17 distinct clusters following analysis of their BOX-PCR-amplified products. However, within these 17 clades, there was 100% Jaccard's similarity coefficient for over half (68 out of 123) of the isolates, indicating a high level of relatedness among the isolates. The BOX-PCR results showed high genetic distance (0% Jaccard's similarity coefficient) in six isolates (namely, TUTVU31, TUTVU-5, TUTVU-4, TUTVU-6, TUTVU-8, and TUTVU63), suggesting that they are genetically distinct from the rest (0% coefficient).

The nodulation assay showed that 81% (99 out of 122) of the isolates could elicit nodule formation in cowpea (IT-18), in fulfilment of Koch's postulates (29). The 23 nonnodulating isolates are likely to be endophytes, as the presence of nodule endophytes has been reported for Vigna species (30). According to Peix et al. (9), endophytic bacteria can coexist with rhizobial strains in nodules without causing any visible harm, and sometimes they even play a supportive role in plant growth and N2 fixation. In fact, nodules may actually be an important ecological niche in the life cycle of these endophytes (31), since they are sheltered from environmental stress as well as competition for resources in the rhizosphere (32). In return, endophytic bacteria can support the growth of their host through secretion of metabolites such as indole-3-acetic acid (IAA) (33), lumicrome, and riboflavin (34).

The use of morphological traits in this study revealed considerable variations among the isolates. About 94% of them that were proven by Koch's postulates to be nodule-forming rhizobia took 5 to 10 days to grow on YMA plates, while 6% took 2 to 4 days. This could suggest the presence of fast-growing (Rhizobium) and slow-growing (Bradyrhizobium) rhizobial species among the isolates. Various studies have reported the ability of both Rhizobium and Bradyrhizobium to nodulate cowpea (35, 36). The results of this study are therefore consistent with those reports. The results of this study also further confirmed the nodulation promiscuity of cowpea as a host plant (37).

Multiple alleles in the rRNA operon of bacteria account for spacer variations between and among species and can be used for typing and identification of rhizobial species (38). The ITS ribosomal genes have been shown to be good markers for assessing rhizobial diversity (16). In this study, ITS-PCR analysis grouped the 99 rhizobial isolates into 17 distinct ITS types, which showed very high polymorphism (258 to 1,350 bp) within the ITS region of the rhizobial genomic DNA. These results have therefore confirmed the discriminatory power of ITS tools in rhizobial diversity studies (16, 39, 40). The differences in product size have been attributed to variations in the number, length, and composition of the ITS spacer regions of various bacteria (38), partly explained by the insertion and/or deletion of tRNA genes in the ITS regions (41). Furthermore, the nonamplification of the ITS regions of isolates TUTVU37 and TUTVU49 may be explained by the possibility of an absence of tRNA genes within their ITS regions and/or primer and thermal cycling incompatibility. It has been suggested that the presence of the double bands observed following gel electrophoresis of the ITS-PCR-amplified products in isolates TUTVU54, TUTVU58, TUTVU65, TUTVU66, TUTVU17, TUTVU98, and TUTVU99 could be due to variations in length and/or the formation of heteroduplex DNA structures and single-stranded DNA during PCR amplification (42, 43). However, this could also be attributed to base deletions in a single strand of the DNA duplex which resulted in differences in electrophoretic mobility (44).

The ITS-RFLP analysis delineates bacteria based on the differences in the recognition sites for restriction endonuclease enzymes within the ITS regions of the genomic DNA. This has been shown to be applicable to rhizobial DNA fingerprinting (16, 45, 46). In this study, ITS-RFLP analysis of 99 rhizobial isolates yielded between 19 and 23 different fingerprint profiles for each endonuclease enzyme, suggesting the presence of different recognition sites for the endonuclease enzymes. Five isolates (namely, TUTVU33, TUTVU42, TUTVU47, TUTVU50, and TUTVU62), however, appeared not to have the recognition sites (GCGC) within their ITS regions for the four base-cutting HaeII restriction endonuclease, as they failed to digest with this enzyme. This is similar to the findings of Ngo Nkot et al. (47).

In the combined ITS-RFLP cluster analysis, 100% Jaccard's similarity coefficient was observed in 68 out of the 99 rhizobial isolates located in different clusters, suggesting that despite the differences observed for each restriction endonuclease enzyme when analyzed singly, most of the isolates had a similar number of enzyme recognition sites for the three restriction endonuclease enzymes. This could be attributed to intraspecific differences between and among the isolates (47). However, unlike other studies, where a correlation was found between rhizobial diversity and either the land use system (47) or the environment (48), the rhizobial isolates in this study were distributed across all clusters irrespective of geographic origin or cowpea variety (Table 1). For example, in clade II, which contained more than half (57%) of the isolates, 28%, 21%, and 51% were, respectively, isolated from root nodules collected at Muriaze, Ruace, and Sussundenga. Similarly, 49% and 51% of microsymbionts were isolated from cowpea varieties IT-18 and IT-1263, respectively. However, there was a 100% Jaccard's similarity among the rhizobial isolates in clades I and II, thus suggesting the presence of similar recognition sites for all three endonuclease enzymes within the ITS genomic regions of these isolates. The highest heterogeneity was found among five rhizobial isolates (namely, TUTVU31, TUTVU2, TUTVU99, TUTVU33, and TUTVU54) which showed marked distinctiveness based on their endonuclease enzyme recognition sites. This distinctiveness was also observed in the commercial strain BIOFIX, which suggests that this rhizobial strain is very different from the resident rhizobial populations of the experimental sites.

In the BOX-PCR analysis, the BIOFIX inoculant genome showed 100% similarity with only one isolate (TUTVU26), which suggests that nodule occupancy by the inoculant strain was very low. This could be attributed to poor competitiveness of the introduced strain compared to the resident native cowpea rhizobia in the soil, or incompatibility between the inoculant strain and the cowpea varieties used.

Within Africa, where the crop is indigenous, studies of cowpea rhizobia have found nodulation of the legume by species belonging to the genus Bradyrhizobium in Senegal, Botswana, Ghana, South Africa, Angola, and Namibia (7, 36, 37, 48) and to the genus Rhizobium based on rhizobial growth rate (35). However, beyond the African continent, cowpea nodulation by species of Rhizobium, Sinorhizobium, Ralstonia, Achromobacter, and Microvirga has also been reported (15, 23, 25, 49–51). In this study, the evolutionary relationships of cowpea rhizobia originating from three agroecological zones of Mozambique were determined using sequence analysis of their ribosomal, symbiotic, and housekeeping genes in order to ascertain the microsymbionts nodulating cowpea in that country.

Nucleotide sequence analysis of the 16S rRNA gene is universally accepted as a way to determine the phylogenetic relationships of all microorganisms on Earth (9). In this study, phylogenetic analysis of the 16S rRNA gene revealed the nodulation of cowpea by diverse Rhizobium and Bradyrhizobium species in Mozambican soils.

Housekeeping genes have been extensively used in Bradyrhizobium phylogeny in order to properly delineate closely related species (52–54). In this study, the single-gene phylogenies were highly congruent between and among the four housekeeping genes (gyrB, glnII, recA, and rpoB). This congruence was, however, predominant in the test isolates that were closely related to the B. elkanii lineage, a finding consistent with the 16S rRNA phylogeny. Beyond this, however, the test isolates showed various degrees of incongruence in the four housekeeping gene phylogenies. For example, although isolate TUTVU1 was closely related to B. diazoefficiens in the gyrB phylogeny, it showed a close relationship with B. yuanmingense in both the glnII and rpoB phylogenies. Similarly, isolates TUTVU13 and TUTVU44, which were closely related to B. yuanmingense in the rpoB phylogeny, grouped with B. diazoefficiens in the gyrB and glnII phylogenies. Like this in recA phylogeny, isolate TUTVU63, which grouped with the type strain of B. stylosanthis, consistently showed a proximal relationship with B. pachyrhizi in the other housekeeping gene phylogenies. These inconsistencies with housekeeping gene phylogenies could be attributed to horizontal gene transfer, subsequent recombination events, and/or differences in the evolutionary history of the gene, (55–58).

Single housekeeping gene phylogenies may therefore not always reflect organismal phylogeny, as they can be sensitive to unequal evolutionary rates among taxa and nucleotide sites within a single gene (59) and could lack adequate phylogenetic information for the resolution of all relationships (60). Phylogenetic information is therefore usually deduced from more than one locus in constructing evolutionary trees (52, 60). In this study, concatenated phylogeny clearly grouped the test isolates into two different major clades, thus clarifying the discrepancies that were found between single-gene phylogenies. Within the B. elknaii lineage (clade I), all the test isolates revealed less than 1% sequence divergence with B. pachyrhizi. Recently, B. pachyrhizi was identified as a cowpea microsymbiont in Angola (36) and Spain (24), although it was originally reported as the microsymbiont nodulating yam bean, Pachyrhizus erosus (61). Likewise, the nodulation of cowpea by B. arachidis, B. elkanii, and B. yuanmingense has been previously reported (24, 62).

Isolates TUTVU1, TUTVU5, TUTVU7, TUTVU13, and TUTVU44 obtained from root nodules of cowpea varieties IT-18 and IT-1263 at Muriaze and Ruace grouped together without any type reference strains and could therefore belong to a novel bradyrhizobial species in Mozambican soils.

This study further found that along with Bradyrhizobium species, fast-growing isolates belonging to Rhizobium also nodulated cowpea in Mozambican soils, a finding consistent with reports from Zimbabwe (35), Brazil (49), and China (22). But Rhizobium isolates could also form effective root nodules on cowpea variety IT-18 in this study and had identical to nearly identical sequences with three different Rhizobium type strains. This is in contrast to the report by Gronemeyer et al. (36) that fast-growing bacteria isolated from cowpea nodules in Namibia failed to induce nodulation in cowpea.

In this study, single housekeeping gene phylogenies showed a high degree of congruence between the gyrB and glnII phylogenies for all the test isolates, although there was a small variation in sequence identity with reference type strains. In the concatenated (gyrB plus glnII) phylogeny, isolates TUTVU50 and TUTVU31 clustered with R. tropici, with 100% bootstrap support, while isolates TUTVU33 and TUTVU67 were closely related to Neorhizobium galegae. Interestingly, Neorhizobium galegae, which was originally isolated from Galega officinalis (63) and has a worldwide distribution, has also been reported to be a microsymbiont of another Vigna species, Vigna radiata (64). Although isolate TUTVU40 consistently clustered with R. pusense in both subsets of the concatenated gene phylogenies, this reference type strain (R. pusense) was originally isolated from the rhizosphere of Cicer arientinum (65) and has not been reported as a nodulating bacterial symbiont of cowpea.

The gyrB product amino acid sequence length variation between TUTVU7 and other test isolates as well as reference type bradyrhizobial strains suggests some variation in their protein structure which might have affected stability and function of the protein. In an earlier study, aberration in the gyrB gene caused resistance to the antibiotic novobiocin in Escherichia coli (66).

Maximum likelihood phylogeny of the symbiotic nifH gene showed inconsistency with other core housekeeping gene phylogenies (Fig. 7). For example, all the isolates in clade III formed a monophyletic group without any reference type strain in the nifH phylogeny but clustered with B. elkanii in the core housekeeping genes. This suggests that there might be differences in the evolutionary history of chromosomal and symbiotic genes. The inconsistency of nifH and housekeeping gene phylogenies could be due to interstrain gene transfer and/or recombination of nifH sequences. In fact, the phylogenies of symbiotic genes are reported to differ from those of the core housekeeping genes (55, 67), an argument that is strongly supported by the nifH and housekeeping gene phylogenies obtained in this study. The clustering of isolates in clade VII without any type reference strain in nifH phylogeny probably indicates that the nifH gene of this clade has an origin different from that of known bradyrhizobial strains.

Taken together, the results of this study have identified the microsymboints nodulating cowpea in Mozambique. From the literature, this is the pioneer report of nodule occupancy by Rhizobium species in Africa, which indicates a wider rhizobial species boundary of the crop in its center of origin. The possibility of greater rhizobial diversity, however, cannot be ruled out, as this study covered only one locality in 3 out of the 10 agroecological zones of Mozambique. This study has contributed to the global database on the distribution of rhizobial species and reported the presence of diverse Bradyrhizobium spp. (B. pachyrhizi, B. elkanii, and B. yuanmingense) and a novel Bradyrhizobium sp., as well as R. tropici, R. pusense, and Neorhizobium galegae, as nodulating microsymbionts of cowpea in Mozambique. Given the large diversity of rhizobia nodulating cowpea in Mozambique, it could prove useful to evaluate these test isolates for symbiotic efficiency and competitiveness as a first step toward identifying superfixers for inoculant production in order to increase food and nutritional security. Furthermore, isolate TUTVU7 produced novel sequences of unknown origin, and this needs functional confirmation using gyrase DNA (gyrB) during replication and transcription of biological processes.

MATERIALS AND METHODS

Description of experimental sites.Two farmer-adopted improved cowpea varieties were used. These include IT-1263, which is semierect, with indeterminate growth, drought tolerant, and high yielding and produces dark brown large seeds, and IT-18, which is erect, with determinate growth, and high yielding and produces small light brown seeds. The field experiments were set up at three sites during the 2014-2015 cropping seasons: at Muriaze in Nampula Province of northern Mozambique, Ruace in Zambezia Province of central Mozambique, and Sussundenga in Manica Province of central Mozambique (Fig. 8).

FIG 8
FIG 8

Three test locations in different provinces in Mozambique.

The three sites are located in different agroecological zones (AEZs). Muriaze, in Nampula Province, is in AEZ 7, which is a vast region in northern Mozambique (68) with elevations ranging from 200 to 1,000 m above sea level and annual rainfall of 1,000 to 1,400 mm. Its soil types vary, with utisols and oxisols as the most predominant. Ruace, in Zambezia Province, is in AEZ 10, a high-elevation region, situated 1,000 m above sea level. The annual rainfall exceeds 1,200 mm, and the soils have a predominance of high clay utisols. Sussundenga, in Manica Province, is in AEZ 4, which lies 200 to 1,000 m above sea level. It has an annual rainfall of 1,000 to 1,200 mm and soils dominated by oxisols. The geographical and soil details of test locations are provided in Table 2.

View this table:
TABLE 2

Geographical and soil information for test locations

Two rhizobial treatments were used (inoculated and uninoculated) for the two test varieties at all locations. All seeds were preinoculated in the shade with a commercial cowpea inoculant (BIOFIX; MEA, Kenya), according to the manufacturer's instructions.

Root nodule sampling and isolation of bacteria.Dark green healthy plants were sampled for nodules at 50% flowering to early podding stage. The root nodules were carefully detached and stored in vials containing silica gel at 4°C prior to molecular analysis. The nodules were surface sterilized using standard methods (29, 69). First, the nodules were rehydrated by soaking in sterile distilled water for a few hours and then placed in 95% ethanol for 10 s to break their surface tension. The nodules were surface sterilized in 3% (vol/vol) sodium hypochlorite for 4 min, rinsed in six changes of sterile distilled water, and transferred aseptically into sterile petri dishes for bacterial isolation. A loopful of last washed water was streaked on a yeast mannitol agar (YMA) plate to see if the nodule surface was well sterilized. Each sterilized root nodule was crushed aseptically in a drop of sterile distilled water, using sterile blunt forceps. A loopful of the macerate was streaked on YMA plates. The plates were incubated at 28°C and observed for bacterial growth up to 14 days. Visible colonies were purified by reculturing on YMA plates until single homogenous colonies were obtained. Based on their colony appearance on YMA, isolates were grouped as slow (>5 days) and fast (<4 days) growers. In addition, colony characteristics such as shape, color, texture, and size were scored for each rhizobial isolate (70).

Pure single colonies were maintained as working cultures on YMA slants in sterile McCartney bottles at 4°C, and another batch was stored at −20°C in 40% glycerol as stock culture.

Nodulation assay.An authentication assay was done for nodule formation on the homologous host by each bacterial isolate. For this, a loopful of each pure single colony isolate was cultured in 6 ml of YM broth to exponential growth phase (≈1 × 107 to 1 × 108 cells · ml−1) and used as inoculum. Seedlings of cowpea variety IT-18 were raised aseptically as described by Somasegaran and Hoben (69) and Woomer et al. (70) using beach sand as growth medium. The beach sand was washed, placed in 1-liter pots, covered with cotton and aluminum foil, and autoclaved at 121°C for 20 min. The experiment was set up in triplicates, and three seeds were sown per pot. The seedlings were thinned down to one plant per pot 3 days after germination. The bacterial inoculation used 2 ml of broth culture per seedling and was done aseptically under a laminar-flow hood 1 day after thinning. Uninoculated seedlings were included as negative controls, while 0.5 mM KNO3-fed seedlings served as positive controls. The pots were arranged in a randomized complete block design in the greenhouse. Watering of plants with sterile distilled water was done when necessary and was alternated with the supply of Broughton & Dilworth (71) nitrogen-free nutrient solution. Plant harvesting was done 3 weeks after inoculation for nodulation assessment.

Total genomic DNA extraction and BOX-PCR fingerprinting.Total genomic DNA was extracted using a GenElute bacterial DNA isolation kit according to the manufacturer's instructions (Sigma-Aldrich, USA). The integrity of the extracted DNA was evaluated on a 1% agarose gel stained with ethidium bromide. The genomic DNA was amplified using BOXA1R primer (26) to generate fingerprint patterns. DNA amplification was performed in a final reaction volume of 25 μl comprising 1 μl (50 to 80 ng) of genomic DNA, 3 μl (5×) of buffer, 1 μl (10 pM) of primer, 0.1 μl (5 U) of Taq polymerase (Bioline, USA), and 18.9 μl of sterile distilled water. The reaction mixture was incubated in a thermal cycler (T100; Bio-Rad, USA) at standard temperature profiles (Table 3). The PCR-amplified products (25 μl) were mixed with 5 μl (6×) of loading dye and placed in wells of 1.2% (wt/vol) agarose gel containing 1× Tris-acetate-EDTA (TAE) stained with ethidium bromide for gel electrophoresis at 95 V for 3 h. Gel imaging and documentation were done using a GEL Doc XR+ molecular imager (Bio-Rad). The data were analyzed using Bio-Numerics software (temporary license from Applied Maths, Belgium; permission received to publish). Similarity matrices were calculated using Jaccard's coefficient (72), and cluster analyses were generated using the unweighted pair group method with arithmetic averages (UPGMA) algorithm (73). Cluster analysis was done at 50% Jaccard's similarity coefficient, and distinct clades were considered to represent diverse groups.

View this table:
TABLE 3

Primers and PCR amplification setup for each gene in the present study

Determination of diversity index.To determine the genetic diversity at each experimental site, diversity indices were estimated based on the representative groupings in the dendrogram. Species diversity was computed using the Shannon index (74) as H′ = −Σ [(n1/N) ln (n1/N)], where N is the total number of all isolates from each experimental site and n1 is the number of isolates belonging to a particular clade at each experimental site. The bacterial species richness was computed using the Margalef index (75) as R1 = [(S − 1)/ln (n)], where S is the total number of representative clades at each experimental site and n is the total number of isolates in all the clades at each experimental site.

The species evenness was measured using the Pielou index (76) as E1 = H′/ln(S), where H′ is the Shannon index and S is the total number of representative clades at each experimental site.

PCR amplification of the 16S-23S rRNA ITS region.The intergenic transcribed spacer (ITS; 16S-23S rRNA) region of the bacterial genomic DNA was amplified using the respective primer pairs to generate fingerprint patterns. DNA amplification was performed as described for BOX-PCR fingerprinting; a list of primers and thermal cycling conditions are presented in Table 3.

RFLP analysis of the PCR-amplified products of the 16S-23S rRNA (ITS) region.The PCR-amplified ITS products were digested using 6-bp-cutting (HindIII), 4-bp-cutting (HaeII), and 5-bp-cutting (HinfI) restriction endonuclease enzymes. The enzyme digestion was carried out by following the manufacturer's instructions (Thermo Scientific, Lithuania). Gel electrophoresis of the digested products was done on a 3% agarose gel at 85 V for 2.5 h. Band sizes were scored against a 100-bp ladder (GeneRuler).

The fingerprint banding patterns generated were analyzed by designating an alphabet to each distinct pattern. The different patterns were then scored based on the presence (1) or absence (0) of the three test endonuclease enzymes' digestion site for each isolate, thus generating a binary matrix. The Jaccard's similarity coefficient (72) was calculated using the similarity matrices, followed by cluster analysis using the UPGMA algorithm (73) with NTSYS pc 2.1 software (USA) (77).

PCR amplification of the 16S rRNA, nifH, and housekeeping genes (glnII, rpoB, recA, and gyrB).The target genes from the rhizobial genomic DNA of each test isolate were amplified using suitable primer pairs and thermal cycling conditions as described for BOX-PCR fingerprinting. All PCR-amplified products (16S rRNA, nifH, glnII, rpoB, recA, and gyrB regions) were purified with a PCR cleanup kit (New England BioLabs, USA), followed by gene sequencing (Macrogen, The Netherlands).

Sequence and phylogenetic analysis.The integrity of each gene sequence was verified using BioEdit 7.0.0 software (78), followed by the determination of their relatedness to type reference strain sequences in the GenBank database (https://www.ncbi.nlm.nih.gov/genbank/) using the BLASTn program. Each isolate sequence was aligned with type reference strain (GenBank database) using MUSCLE (79). Phylogenetic trees were generated from the aligned sequences using the MEGA 6 software program (80), which calculates evolutionary distances using the Kimura 2-parameter model (81). The evolutionary history was inferred using the maximum likelihood method algorithm with 1,000 bootstraps (82). The pairwise sequence identities of single and concatenated gene sequences were calculated using the Bio-edit sequence identity matrix. The conserved, variable, and parsimony nucleotide informative sites were determined for the genes under study (MEGA 6). The sequences were then submitted to GenBank to obtain accession numbers (Table S1).

Metagenomic analysis of 16S rRNA.Rhizosphere soil samples of two test cowpea varieties grown at the experimental sites (Muriaze, Ruace, and Sussundenga) were collected and stored at −20°C prior to DNA isolation. Bacterial genomic DNA was extracted from 0.5 g of rhizosphere soil using PowerSoil DNA isolation kit according to the manufacturer's instructions (Mo Bio, USA). Each DNA sample was subjected to PCR amplification using primer pairs that contained adapter-ligated fragments 5′TCGTCGGCAGCGTCAGATGTGTATAAGAGACAGCCTACGGGNGGCWGCAG3′ and 5′GTCTCGTGGGCTCGGAGATGTGTATAAGAGACAGGACTACHVGGGTATCTAATCC3′, thus targeting variable regions V3 and V4 of the 16S rRNA gene (83). The PCR was carried out with 10 to 15 ng DNA in 25-μl reaction volume containing 12.5 μl of 2× KAPA HiFi hotstart ready mix and 5 μl of each primer (1 μM), with the following temperature profile: 95°C for 30 s, 25 cycles of 95°C for 30 s, 55°C for 30 s, and 72°C for 30 s, and then 72°C for 5 min. The PCR-amplified samples were sent to Macrogen, South Korea, for Miseq Illumina paired-end sequencing and analysis.

Accession number(s).The sequences of cowpea-nodulating rhizobial isolates used in this study are available in the GenBank database under accession numbers KY941240 through KY941398.

ACKNOWLEDGMENTS

This study was supported with grants from the Bill and Melinda Gates Foundation Project on Capacity Building in Legume Sciences in Africa, the South African Department of Science and Technology, the Tshwane University of Technology, the National Research Foundation in Pretoria, and the South African Research Chair in Agrochemurgy and Plant Symbioses. I.N.C. is grateful for a competitive doctoral fellowship obtained from the Bill and Melinda Gates Foundation Project on Capacity Building in Legume Sciences in Africa.

FOOTNOTES

    • Received 15 August 2017.
    • Accepted 19 October 2017.
    • Accepted manuscript posted online 3 November 2017.

  • Supplemental material for this article may be found at https://doi.org/10.1128/AEM.01712-17.

This is an open-access article distributed under the terms of the Creative Commons Attribution 4.0 International license.

REFERENCES

  1. 1.↵
    1. Walker T,
    2. Cunguara B,
    3. Donovan C
    . 2016. Assessing directions for cowpea R&D in USAID'S feed the future program in Mozambique. University of Illinois at Urbana-Champaign, Urbana, IL.
  2. 2.↵
    1. Belane AK,
    2. Dakora FD
    . 2009. Measurement of N2 fixation in 30 cowpea (Vigna unguiculata L. Walp.) genotypes under field conditions in Ghana, using the 15N natural abundance technique. Symbiosis 48:47–56. doi:10.1007/BF03179984.
    OpenUrlCrossRef
  3. 3.↵
    1. Gonçalves A,
    2. Goufo P,
    3. Barros A,
    4. Domínguez-Perles R,
    5. Trindade H,
    6. Rosa EA,
    7. Ferreira L,
    8. Rodrigues M
    . 2016. Cowpea (Vigna unguiculata L. Walp), a renewed multipurpose crop for a more sustainable agri-food system: nutritional advantages and constraints. J Sci Food Agricul 96:2941–2951. doi:10.1002/jsfa.7644.
    OpenUrlCrossRef
  4. 4.↵
    1. Masson-Boivin C,
    2. Giraud E,
    3. Perret X,
    4. Batut J
    . 2009. Establishing nitrogen-fixing symbiosis with legumes: how many rhizobium recipes? Trends Microbiol 17:458–466. doi:10.1016/j.tim.2009.07.004.
    OpenUrlCrossRefPubMedWeb of Science
  5. 5.↵
    1. Berrada H,
    2. Fikri-Benbrahim K
    . 2014. Taxonomy of the rhizobia: current perspectives. Br Microbiol Res J 4(6):616. doi:10.9734/BMRJ/2014/5635.
    OpenUrlCrossRef
  6. 6.↵
    1. Peoples MB,
    2. Brockwell J,
    3. Herridge DF,
    4. Rochester IJ,
    5. Alves BJR,
    6. Urquiaga S,
    7. Boddey RM,
    8. Dakora FD,
    9. Bhattarai S,
    10. Maskey SL,
    11. Sampet C
    . 2009. The contributions of nitrogen-fixing crop legumes to the productivity of agricultural systems. Symbiosis 48:1–17. doi:10.1007/BF03179980.
    OpenUrlCrossRefWeb of Science
  7. 7.↵
    1. Pule-Meulenberg F,
    2. Belane AK,
    3. Krasova-Wade T,
    4. Dakora FD
    . 2010. Symbiotic functioning and bradyrhizobial biodiversity of cowpea (Vigna unguiculata L. Walp.) in Africa. BMC Microbiol 10:89. doi:10.1186/1471-2180-10-89.
    OpenUrlCrossRefPubMed
  8. 8.↵
    1. Msimbira LA,
    2. Jaiswal SK,
    3. Dakora FD
    . 2016. Identification and characterization of phages parasitic on bradyrhizobia nodulating groundnut (Arachis hypogaea L.) in South Africa. Appl Soil Ecol 108:334–340. doi:10.1016/j.apsoil.2016.09.010.
    OpenUrlCrossRef
  9. 9.↵
    1. Peix A,
    2. Ramírez-Bahena MH,
    3. Velazquez E,
    4. Bedmar EJ
    . 2015. Bacterial associations with legumes. Crit Rev Plant Sci 34:17–42. doi:10.1080/07352689.2014.897899.
    OpenUrlCrossRef
  10. 10.↵
    1. Suneja P,
    2. Dudeja SS,
    3. Dahiya P
    . 2016. Deciphering the phylogenetic relationships among rhizobia nodulating chickpea: a review. J Appl Biol Biotechnol 4:61–70. doi:10.7324/JABB.2016.40310.
    OpenUrlCrossRef
  11. 11.↵
    1. Rashid MHO,
    2. Young JPW,
    3. Everall I,
    4. Clercx P,
    5. Willems A,
    6. Braun MS,
    7. Wink M
    . 2015. Average nucleotide identity of genome sequences supports the description of Rhizobium lentis sp. nov., Rhizobium bangladeshense sp. nov. and Rhizobium binae sp. nov. from lentil (Lens culinaris) nodules. Int J Syst Evol Microbiol 65:3037–3045. doi:10.1099/ijs.0.000373.
    OpenUrlCrossRef
  12. 12.↵
    1. Ardley JK,
    2. Parker MA,
    3. de Meyer SE,
    4. Trengove RD,
    5. O'Hara GW,
    6. Reeve WG,
    7. Yates RJ,
    8. Dilworth MJ,
    9. Willems A,
    10. Howieson JG
    . 2012. Microvirga lupini sp. nov., Microvirga lotononidis sp. nov. and Microvirga zambiensis sp. nov. are alpha proteobacterial root-nodule bacteria that specifically nodulate and fix nitrogen with geographically and taxonomically separate legume hosts. Int J Syst Evol Microbiol 62:2579–2588. doi:10.1099/ijs.0.035097-0.
    OpenUrlCrossRefPubMedWeb of Science
  13. 13.↵
    1. Maynaud G,
    2. Willems A,
    3. Soussou S,
    4. Vidal C,
    5. Maure L,
    6. Moulin L,
    7. Cleyet-Marel JC,
    8. Brunel B
    . 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. Syst Appl Microbiol 35:65–72. doi:10.1016/j.syapm.2011.11.002.
    OpenUrlCrossRefPubMed
  14. 14.↵
    1. Bournaud C,
    2. de Faria SM,
    3. dos Santos JM,
    4. Tisseyre P,
    5. Silva M,
    6. Chaintreuil C,
    7. Gross E,
    8. James EK,
    9. Prin Y,
    10. Moulin L
    . 2013. Burkholderia species are the most common and preferred nodulating symbionts of the Piptadenia group (tribe Mimoseae). PLoS One 8:e63478. doi:10.1371/journal.pone.0063478.
    OpenUrlCrossRef
  15. 15.↵
    1. Radl V,
    2. Simoes-Araujo JL,
    3. Leite J,
    4. Passos SR,
    5. Martins LM,
    6. Xavier GR,
    7. Rumjanek NG,
    8. Baldani JI,
    9. Zilli JE
    . 2014. Microvirga vignae sp. nov., a root nodule symbiotic bacterium isolated from cowpea grown in semi-arid Brazil. Int J Syst Evol Microbiol 64:725–730. doi:10.1099/ijs.0.053082-0.
    OpenUrlCrossRefPubMed
  16. 16.↵
    1. Jaiswal SK,
    2. Msimbira LA,
    3. Dakora FD
    . 7 March 2017. Phylogenetically diverse group of native bacterial symbionts isolated from root nodules of groundnut (Arachis hypogaea L.) in South Africa. Syst Appl Microbiol. doi:10.1016/j.syapm.2017.02.002.
    OpenUrlCrossRef
  17. 17.↵
    1. Binde DR,
    2. Menna P,
    3. Bangel EV,
    4. Barcellos FG,
    5. Hungria M
    . 2009. rep-PCR fingerprinting and taxonomy based on the sequencing of the 16S rRNA gene of 54 elite commercial rhizobial strains. Appl Microbiol Biotechnol 83:897–908. doi:10.1007/s00253-009-1927-6.
    OpenUrlCrossRefPubMed
  18. 18.↵
    1. Van Berkum P,
    2. Terefework Z,
    3. Paulin L,
    4. Suomalainen S,
    5. Lindström K,
    6. Eardly BD
    . 2003. Discordant phylogenies within the rrn loci of rhizobia. J Bacteriol 185:2988–2998. doi:10.1128/JB.185.10.2988-2998.2003.
    OpenUrlAbstract/FREE Full Text
  19. 19.↵
    1. Saïdi S,
    2. Chebil S,
    3. Gtari M,
    4. Mhamdi R
    . 2013. Characterization of root-nodule bacteria isolated from Vicia faba and selection of plant growth promoting isolates. World J Microbiol Biotechnol 29:1099–1106. doi:10.1007/s11274-013-1278-4.
    OpenUrlCrossRefPubMed
  20. 20.↵
    1. Martens M,
    2. Dawyndt P,
    3. Coopman R,
    4. Gillis M,
    5. De Vos P,
    6. Willems A
    . 2008. Advantages of multilocus sequence analysis for taxonomic studies: a case study using 10 housekeeping genes in the genus Ensifer (including former Sinorhizobium). Int J Syst Evol Microbiol 58:200–214. doi:10.1099/ijs.0.65392-0.
    OpenUrlCrossRefPubMedWeb of Science
  21. 21.↵
    1. Germano MG,
    2. Menna P,
    3. Mostasso FL,
    4. Hungria M
    . 2006. RFLP analysis of the rRNA operon of a Brazilian collection of bradyrhizobial strains from 33 legume species. Int J Syst Evol Microbiol 56:217–229. doi:10.1099/ijs.0.02917-0.
    OpenUrlCrossRefPubMedWeb of Science
  22. 22.↵
    1. Zhang WT,
    2. Yang JK,
    3. Yuan TY,
    4. Zhou JC
    . 2007. Genetic diversity and phylogeny of indigenous rhizobia from cowpea [Vigna unguiculata (L.) Walp.]. Biol Fertil Soils 44:201–210. doi:10.1007/s00374-007-0196-8.
    OpenUrlCrossRef
  23. 23.↵
    1. Guimaraes AA,
    2. Jaramillo PMD,
    3. Nobrega RSA,
    4. Florentino LA,
    5. Silva KB,
    6. de Souza Moreira FM
    . 2012. Genetic and symbiotic diversity of nitrogen-fixing bacteria isolated from agricultural soils in the Western Amazon by using cowpea as the trap plant. Appl Environ Microbiol 78:6726–6733. doi:10.1128/AEM.01303-12.
    OpenUrlAbstract/FREE Full Text
  24. 24.↵
    1. Bejarano A,
    2. Ramírez-Bahena MH,
    3. Velazquez E,
    4. Peix A
    . 2014. Vigna unguiculata is nodulated in Spain by endosymbionts of Genisteae legumes and by a new symbiovar (vignae) of the genus Bradyrhizobium. Syst Appl Microbiol 37:533–540. doi:10.1016/j.syapm.2014.04.003.
    OpenUrlCrossRef
  25. 25.↵
    1. Sarr PS,
    2. Yamakawa T,
    3. Saeki Y,
    4. Guissé A
    . 2011. Phylogenetic diversity of indigenous cowpea bradyrhizobia from soils in Japan based on sequence analysis of the 16S–23S rRNA internal transcribed spacer (ITS). Syst Appl Microbiol 34:285–292. doi:10.1016/j.syapm.2010.11.021.
    OpenUrlCrossRefPubMed
  26. 26.↵
    1. Versalovic J,
    2. Schneider M,
    3. de Bruijn FJ,
    4. Lupski JR
    . 1994. Genomic fingerprinting of bacteria using repetitive sequence-based polymerase chain reaction. Methods Mol Cell Biol 5:25–40.
    OpenUrl
  27. 27.↵
    1. Kaschuk G,
    2. Hungria M,
    3. Andrade DS,
    4. Campo RJ
    . 2006. Genetic diversity of rhizobia associated with common bean (Phaseolus vulgaris L.) grown under no-tillage and conventional systems in Southern Brazil. Appl Soil Ecol 32:210–220. doi:10.1016/j.apsoil.2005.06.008.
    OpenUrlCrossRefWeb of Science
  28. 28.↵
    1. Menna P,
    2. Pereira AA,
    3. Bangel EV,
    4. Hungria M
    . 2009. rep-PCR of tropical rhizobia for strain fingerprinting, biodiversity appraisal and as a taxonomic and phylogenetic tool. Symbiosis 48:120–130. doi:10.1007/BF03179991.
    OpenUrlCrossRef
  29. 29.↵
    1. Vincent JM
    . 1970. A manual for the practical study of root nodule bacteria. Blackwell Scientific, Oxford, United Kingdom.
  30. 30.↵
    1. Pandya M,
    2. Naresh Kumar G,
    3. Rajkumar S
    . 2013. Invasion of rhizobial infection thread by non rhizobia for colonization of Vigna radiata root nodules. FEMS Microbiol Lett 348:58–65. doi:10.1111/1574-6968.12245.
    OpenUrlCrossRefPubMedWeb of Science
  31. 31.↵
    1. Deng ZS,
    2. Zhao LF,
    3. Kong ZY,
    4. Yang WQ,
    5. Lindstrom K,
    6. Wang ET,
    7. Wei GH
    . 2011. Diversity of endophytic bacteria within nodules of the Sphaerophysa salsula in different regions of Loess Plateau in China. FEMS Microbiol Ecol 76:463–475. doi:10.1111/j.1574-6941.2011.01063.x.
    OpenUrlCrossRefPubMed
  32. 32.↵
    1. Kobayashi DY,
    2. Palumbo JD
    . 2000. Bacterial endophytes and their effects on plants and uses in agriculture, p 199. In Bacon CW, White JF, Jr (ed), Microbial endophytes. Marcel Dekker, New York, NY.
  33. 33.↵
    1. Li JH,
    2. Wang ET,
    3. Chen WF,
    4. Chen WX
    . 2008. Genetic diversity and potential for promotion of plant growth detected in nodule endophytic bacteria of soybean grown in Heilongjiang province of China. Soil Biol Biochem 40:238–246. doi:10.1016/j.soilbio.2007.08.014.
    OpenUrlCrossRef
  34. 34.↵
    1. Dakora FD
    . 2015. Lumichrome: a bacterial signal molecule influencing plant growth, p 389. In de Bruijn FJ (ed), Biological nitrogen fixation. John Wiley & Sons, Inc, Hoboken, NJ.
  35. 35.↵
    1. Mpepereki S,
    2. Wollum AG,
    3. Makonese F
    . 1996. Diversity in symbiotic specificity of cowpea rhizobia indigenous to Zimbabwean soils. Plant Soil 186:167–171. doi:10.1007/BF00035071.
    OpenUrlCrossRef
  36. 36.↵
    1. Gronemeyer JL,
    2. Kulkarni A,
    3. Berkelmann D,
    4. Hurek T,
    5. Reinhold-Hurek B
    . 2014. Rhizobia indigenous to the Okavango region in sub-Saharan Africa: diversity, adaptations, and host specificity. Appl Environ Microbiol 80:7244–7257. doi:10.1128/AEM.02417-14.
    OpenUrlAbstract/FREE Full Text
  37. 37.↵
    1. Steenkamp ET,
    2. Stêpkowski T,
    3. Przymusiak A,
    4. Botha WJ,
    5. Law IJ
    . 2008. Cowpea and groundnut in southern Africa are nodulated by diverse Bradyrhizobium strains harbouring nodulation genes that belong to the large pantropical clade common in Africa. Mol Phylogenet Evol 48:1131–1144. doi:10.1016/j.ympev.2008.04.032.
    OpenUrlCrossRefPubMedWeb of Science
  38. 38.↵
    1. Gürtler V,
    2. Stanisich VA
    . 1996. New approaches to typing and identification of bacteria using the 16S–23S rDNA spacer region. Microbiology 142:3–16. doi:10.1099/13500872-142-1-3.
    OpenUrlCrossRefPubMedWeb of Science
  39. 39.↵
    1. Willems A,
    2. Munive A,
    3. de Lajudie P,
    4. Gillis M
    . 2003. In most Bradyrhizobium groups sequence comparison of 16S-23S rDNA internal transcribed spacer regions corroborates DNA-DNA hybridizations. Syst Appl Microbiol 26:203–210. doi:10.1078/072320203322346056.
    OpenUrlCrossRefPubMedWeb of Science
  40. 40.↵
    1. Jaiswal SK,
    2. Beyan SM,
    3. Dakora FD
    . 20 April 2016. Distribution, diversity and population composition of soybean-nodulating bradyrhizobia from different agro-climatic regions in Ethiopia. Biol Fertil Soils doi:10.1007/s00374-016-1108-6.
    OpenUrlCrossRef
  41. 41.↵
    1. Jensen MA,
    2. Webster JA,
    3. Straus N
    . 1993. Rapid identification of bacteria on the basis of polymerase chain reaction-amplified ribosomal DNA spacer polymorphisms. Appl Environ Microbiol 59:945–952.
    OpenUrlAbstract/FREE Full Text
  42. 42.↵
    1. Jensen MA,
    2. Straus N
    . 1993. Effects of PCR conditions on the formation of heteroduplex and single-stranded DNA products in the amplification of bacterial ribosomal DNA spacer regions. PCR Methods Appl 3:186–194. doi:10.1101/gr.3.3.186.
    OpenUrlCrossRefPubMedWeb of Science
  43. 43.↵
    1. Laguerre G,
    2. Mavingui P,
    3. Allard MR,
    4. Charnay MP,
    5. Louvrier P,
    6. Mazurier SI,
    7. Amarger N
    . 1996. Typing of rhizobia by PCR DNA fingerprinting and PCR-restriction fragment length polymorphism analysis of chromosomal and symbiotic gene regions: application to Rhizobium leguminosarum and its different biovars. Appl Environ Microbiol 62:2029–2036.
    OpenUrlAbstract/FREE Full Text
  44. 44.↵
    1. Ruano G,
    2. Kidd KK
    . 1992. Modeling of heteroduplex formation during PCR from mixtures of DNA templates. Genome Res 2:112–116. doi:10.1101/gr.2.2.112.
    OpenUrlAbstract/FREE Full Text
  45. 45.↵
    1. Sikora S,
    2. Redzepovic S
    . 2003. Genotypic characterization of indigenous soybean rhizobia by PCR-RFLP of 16S rDNA, rep-PCR and RAPD analysis. Food Technol Biotechnol 41:61–67.
    OpenUrl
  46. 46.↵
    1. Jaiswal SK,
    2. Anand A,
    3. Dhar B,
    4. Vaishampayan A
    . 2012. Genotypic characterization of phage-typed indigenous soybean bradyrhizobia and their host range symbiotic effectiveness. Microb Ecol 63:116–126. doi:10.1007/s00248-011-9950-4.
    OpenUrlCrossRefPubMed
  47. 47.↵
    1. Ngo Nkot L,
    2. Krasova-Wade T,
    3. Etoa FX,
    4. Sylla SN,
    5. Nwaga D
    . 2008. Genetic diversity of rhizobia nodulating Arachis hypogaea L. in diverse land use systems of humid forest zone in Cameroon. Appl Soil Ecol 40:411–416. doi:10.1016/j.apsoil.2008.06.007.
    OpenUrlCrossRef
  48. 48.↵
    1. Krasova-Wade T,
    2. Ndoye I,
    3. Braconnier S,
    4. Sarr B,
    5. de Lajudie P,
    6. Neyra M
    . 2003. Diversity of indigeneous bradyrhizobia associated with three cowpea cultivars (Vigna unguiculata (L.) Walp.) grown under limited and favorable water conditions in Senegal (West Africa). Afr J Biotechnol 2:13–22. doi:10.5897/AJB2003.000-1003.
    OpenUrlCrossRef
  49. 49.↵
    1. Martins LMV,
    2. Neves MCP,
    3. Rumjanek NG
    . 1997. Growth characteristics and symbiotic efficiency of rhizobia isolated from cowpea nodules of the north-east region of Brazil. Soil Biol Biochem 29:1005–1010. doi:10.1016/S0038-0717(96)00215-5.
    OpenUrlCrossRef
  50. 50.↵
    1. Zhang YF,
    2. Wang ET,
    3. Tian CF,
    4. Wang FQ,
    5. Han LL,
    6. Chen WF,
    7. Chen WX
    . 2008. Bradyrhizobium elkanii, Bradyrhizobium yuanmingense and Bradyrhizobium japonicum are the main rhizobia associated with Vigna unguiculata and Vigna radiata in the subtropical region of China. FEMS Microbiol Lett 285:146–154. doi:10.1111/j.1574-6968.2008.01169.x.
    OpenUrlCrossRefPubMed
  51. 51.↵
    1. Leite J,
    2. Seido SL,
    3. Passos SR,
    4. Xavier GR,
    5. Rumjanek NG,
    6. Martins LMV
    . 2009. Biodiversity of rhizobia associated with cowpea cultivars in soils of the lower half of the São Francisco River Valley. R Bras Ci Solo 33:1215–1226. doi:10.1590/S0100-06832009000500015.
    OpenUrlCrossRef
  52. 52.↵
    1. Vinuesa P,
    2. Rojas-Jiménez K,
    3. Contreras-Moreira B,
    4. Mahna SK,
    5. Prasad BN,
    6. Moe H,
    7. Selvaraju SB,
    8. Thierfelder H,
    9. Werner D
    . 2008. Multilocus sequence analysis for assessment of the biogeography and evolutionary genetics of four Bradyrhizobium species that nodulate soybeans on the Asiatic continent. Appl Environ Microbiol 74:6987–6996. doi:10.1128/AEM.00875-08.
    OpenUrlAbstract/FREE Full Text
  53. 53.↵
    1. Nzoué A,
    2. Miché L,
    3. Klonowska A,
    4. Laguerre G,
    5. de Lajudie P,
    6. Moulin L
    . 2009. Multilocus sequence analysis of bradyrhizobia isolated from Aeschynomene species in Senegal. Syst Appl Microbiol 32:400–412. doi:10.1016/j.syapm.2009.06.002.
    OpenUrlCrossRefPubMed
  54. 54.↵
    1. Rivas R,
    2. García-Fraile P,
    3. Velazquez E
    . 2009. Taxonomy of bacteria nodulating legumes. Rev Microbiol Insight 2:251–269.
    OpenUrl
  55. 55.↵
    1. Zinga MK,
    2. Jaiswal SK,
    3. Dakora FD
    . 2017. Presence of diverse rhizobial communities responsible for nodulation of common bean (Phaseolus vulgaris) in South African and Mozambican soils. FEMS Microbiol Ecol 93:fiw236. doi:10.1093/femsec/fiw236.
    OpenUrlCrossRefPubMed
  56. 56.↵
    1. Zhang XX,
    2. Guo HJ,
    3. Wang R,
    4. Sui XH,
    5. Zhang YM,
    6. Wang ET,
    7. Tian CF,
    8. Chen WX
    . 2014. Genetic divergence of Bradyrhizobium strains nodulating soybeans as revealed by multilocus sequence analysis of genes inside and outside the symbiosis island. Appl Environ Microbiol 80:3181–3190. doi:10.1128/AEM.00044-14.
    OpenUrlAbstract/FREE Full Text
  57. 57.↵
    1. Zhao L,
    2. Fan M,
    3. Zhang D,
    4. Yang R,
    5. Zhang F,
    6. Xu L,
    7. Wei X,
    8. Shen Y,
    9. Wei G
    . 2014. Distribution and diversity of rhizobia associated with wild soybean (Glycine soja Sieb. & Zucc.) in Northwest China. Syst Appl Microbiol 37:449–456. doi:10.1016/j.syapm.2014.05.011.
    OpenUrlCrossRef
  58. 58.↵
    1. Naamala J,
    2. Jaiswal SK,
    3. Dakora FD
    . 2016. Microsymbiont diversity and phylogeny of native bradyrhizobia associated with soybean (Glycine max L. Merr.) nodulation in South African soils. Syst Appl Microbiol 39:336–344. doi:10.1016/j.syapm.2016.05.009.
    OpenUrlCrossRef
  59. 59.↵
    1. Nichols R
    . 2001. Gene trees and species trees are not the same. Trends Ecol Evol 16:358–364. doi:10.1016/S0169-5347(01)02203-0.
    OpenUrlCrossRefPubMedWeb of Science
  60. 60.↵
    1. Gadagkar SR,
    2. Rosenberg MS,
    3. Kumar S
    . 2005. Inferring species phylogenies from multiple genes: concatenated sequence tree versus consensus gene tree. J Exp Zool B Mol Dev Evol 304B:64–74. doi:10.1002/jez.b.21026.
    OpenUrlCrossRefPubMed
  61. 61.↵
    1. Ramírez-Bahena MH,
    2. Peix A,
    3. Rivas R,
    4. Camacho M,
    5. Rodríguez-Navarro DN,
    6. Mateos PF,
    7. Martínez-Molina E,
    8. Willems A,
    9. Velázquez E
    . 2009. Bradyrhizobium pachyrhizi sp. nov. and Bradyrhizobium jicamae sp. nov., isolated from effective nodules of Pachyrhizus erosus. Int J Syst Evol Microbiol 59:1929–1934. doi:10.1099/ijs.0.006320-0.
    OpenUrlCrossRefPubMedWeb of Science
  62. 62.↵
    1. Krasova-Wade T,
    2. Le Quéréb A,
    3. Laguerrec G,
    4. N′Zouéb A,
    5. Ndioned J-A,
    6. do Regoa F,
    7. Sadioe O,
    8. Ndoyef I,
    9. Neyra M
    . 2014. Eco-geographical diversity of cowpea bradyrhizobia in Senegal is marked by dominance of two genetic types. Syst Appl Microbiol 37:129–139. doi:10.1016/j.syapm.2013.10.002.
    OpenUrlCrossRef
  63. 63.↵
    1. Lindstrom K
    . 1989. Rhizobium galegae, a new species of legume root nodule bacteria. Int J Syst Bacteriol 39:365–367. doi:10.1099/00207713-39-3-365.
    OpenUrlCrossRef
  64. 64.↵
    1. Mousavi SA,
    2. Österman J,
    3. Wahlberg N,
    4. Nesme X,
    5. Lavire C,
    6. Vial L,
    7. Paulin L,
    8. de Lajudie P,
    9. Lindström K
    . 2014. Phylogeny of the Rhizobium-Allorhizobium-Agrobacterium clade supports the delineation of Neorhizobium gen. nov. Syst Appl Microbiol 37:208–215. doi:10.1016/j.syapm.2013.12.007.
    OpenUrlCrossRef
  65. 65.↵
    1. Panday D,
    2. Schumann P,
    3. Das SK
    . 2011. Rhizobium pusense sp. nov., isolated from the rhizosphere of chickpea (Cicer arietinum L.). Int J Syst Evol Microbiol 61:2632–2639. doi:10.1099/ijs.0.028407-0.
    OpenUrlCrossRefPubMedWeb of Science
  66. 66.↵
    1. Huang WM
    . 1996. Bacterial diversity based on type II DNA topoisomerase genes. Annu Rev Genet 30:79–107. doi:10.1146/annurev.genet.30.1.79.
    OpenUrlCrossRefPubMedWeb of Science
  67. 67.↵
    1. Stepkowski T,
    2. Watkin E,
    3. McInnes A,
    4. Gurda D,
    5. Gracz J,
    6. Steenkamp ET
    . 2012. Distinct Bradyrhizbium communities nodulate legumes native to temperate and tropical monsoon Australia. Mol Phylogenet Evol 63:265–277. doi:10.1016/j.ympev.2011.12.020.
    OpenUrlCrossRefPubMed
  68. 68.↵
    1. Maria RM,
    2. Yost R
    . 2006. A survey of soil fertility status of four agroecological zones of Mozambique. Soil Sci 171:902–914. doi:10.1097/01.ss.0000228058.38581.de.
    OpenUrlCrossRef
  69. 69.↵
    1. Somasegaran P,
    2. Hoben HJ
    . 1994. Handbook for rhizobia. Springer-Verlag, New York, NY.
  70. 70.↵
    1. Woomer PL,
    2. Karanja N,
    3. Kisamuli SM,
    4. Murwira M,
    5. Bala A
    . 2011. A revised manual for rhizobium methods and standard protocols. http://www.n2africa.org.
  71. 71.↵
    1. Broughton W,
    2. Dilworth M
    . 1971. Control of leghaemoglobin synthesis in snake beans. Biochem J 125:1075–1080. doi:10.1042/bj1251075.
    OpenUrlAbstract/FREE Full Text
  72. 72.↵
    1. Jaccard P
    . 1912. The distribution of flora in the alpine zone. New Phytol 11:37–50. doi:10.1111/j.1469-8137.1912.tb05611.x.
    OpenUrlCrossRef
  73. 73.↵
    1. Sneath PHA,
    2. Sokal RR
    . 1973. Numerical taxonomy. WH Freeman and Co, San Francisco, CA.
  74. 74.↵
    1. Shannon CE,
    2. Weaver W
    . 1949. The mathematical theory of communication. University Illinois Press, Urbana, IL.
  75. 75.↵
    1. Margalef DR
    . 1958. Information theory in ecology. Gen Syst 3:36–71.
    OpenUrl
  76. 76.↵
    1. Pielou EC
    . 1977. Mathematical ecology. Wiley, New York, NY.
  77. 77.↵
    1. Rohlf FJ
    . 2009. NTSYS-pc Numerical Taxonomy System version 2.21c. Exeter Software, New York, NY.
  78. 78.↵
    1. Hall T
    . 2004. BioEdit version 7.0. http://www.mBio.ncsu.edu/BioEdit/bioedit.html.
  79. 79.↵
    1. Edgar RC
    . 2004. MUSCLE: multiple sequence alignment with high accuracy and high throughput. Nucleic Acids Res 32:1792–1797. doi:10.1093/nar/gkh340.
    OpenUrlCrossRefPubMedWeb of Science
  80. 80.↵
    1. Tamura K,
    2. Stecher G,
    3. Peterson D,
    4. Filipski A,
    5. Kumar S
    . 2013. MEGA6: Molecular Evolutionary Genetics Analysis version 6.0. Mol Biol Evol 30:2725–2729. doi:10.1093/molbev/mst197.
    OpenUrlCrossRefPubMedWeb of Science
  81. 81.↵
    1. Kimura M
    . 1980. A simple method for estimating evolutionary rate of base substitution through comparative studies of nucleotide sequences. J Mol Evol 16:111–120. doi:10.1007/BF01731581.
    OpenUrlCrossRefPubMedWeb of Science
  82. 82.↵
    1. Felsenstein J
    . 1985. Confidence limits on phylogenies: an approach using the bootstrap. Evolution 39:783–791.
    OpenUrlCrossRefPubMedWeb of Science
  83. 83.↵
    1. Klindworth A,
    2. Pruesse E,
    3. Schweer T,
    4. Peplies J,
    5. Quast C,
    6. Horn M,
    7. Glöckner FO
    . 2013. Evaluation of general 16S ribosomal RNA gene PCR primers for classical and next-generation sequencing-based diversity studies. Nucleic Acids Res 41(1):e1. doi:10.1093/nar/gks808.
    OpenUrlCrossRefPubMed
  84. 84.
    1. Appunu C,
    2. Sasirekha N,
    3. Prabavathy RV,
    4. Nair S
    . 2009. A significant proportion of indigenous rhizobia from India associated with soybean (Glycine max L.) distinctly belong to Bradyrhizobium and Ensifer genera. Biol Fertil Soils 46:57–63. doi:10.1007/s00374-009-0405-8.
    OpenUrlCrossRef
  85. 85.
    1. Weisburg WG,
    2. Barns SM,
    3. Pelletier DA,
    4. Lane DJ
    . 1991. 16S ribosomal DNA amplification for phylogenetic study. J Bacteriol 173:697–703. doi:10.1128/jb.173.2.697-703.1991.
    OpenUrlAbstract/FREE Full Text
  86. 86.
    1. Marek-Kozaczuk M,
    2. Leszcz A,
    3. Wielbo J,
    4. Wdowiak-Wróbel S,
    5. Skorupska A
    . 2013. Rhizobium pisi sv. trifolii K3.22 harboring nod genes of the Rhizobium leguminosarum sv. trifolii cluster. Syst Appl Microbiol 36:252–258. doi:10.1016/j.syapm.2013.01.005.
    OpenUrlCrossRef
  87. 87.
    1. Stêpkowski T,
    2. Żak M,
    3. Moulin L,
    4. Króliczak J,
    5. Golińska B,
    6. Narożna D,
    7. Mądrzak CJ
    . 2011. Bradyrhizobium canariense and Bradyrhizobium japonicum are the two dominant Rhizobium species in root nodules of lupin and serradella plants growing in Europe. Syst Appl Microbiol 34:368–375. doi:10.1016/j.syapm.2011.03.002.
    OpenUrlCrossRefPubMed
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KEYWORDS

BOX-PCR
ITS
phylogeny
nodulation
horizontal gene transfer
metagenome
Bradyrhizobium
rhizobium
agroecology

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