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Applied and Environmental Microbiology, October 1998, p. 3989-3997, Vol. 64, No. 10
0099-2240/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
Molecular Diversity of Rhizobia Occurring on
Native Shrubby Legumes in Southeastern Australia
Bénédicte
Lafay* and
Jeremy J.
Burdon
Centre for Plant Biodiversity Research, CSIRO
Plant Industry, Canberra ACT 2601, Australia
Received 18 August 1997/Accepted 28 July 1998
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ABSTRACT |
The structure of rhizobial communities nodulating native shrubby
legumes in open eucalypt forest of southeastern Australia was
investigated by a molecular approach. Twenty-one genomic species were
characterized by small-subunit ribosomal DNA PCR-restriction fragment
length polymorphism and phylogenetic analyses, among 745 rhizobial
strains isolated from nodules sampled on 32 different legume host
species at 12 sites. Among these rhizobial genomic species, 16 belonged
to the Bradyrhizobium subgroup, 2 to the Rhizobium leguminosarum subgroup, and 3 to
the Mesorhizobium subgroup. Only one genomic species
corresponded to a known species (Rhizobium tropici). The
distribution of the various genomic species was highly unbalanced
among the 745 isolates, legume hosts, and sites. Bradyrhizobium species were by far the most abundant, and
Rhizobium tropici dominated among the Rhizobium
and Mesorhizobium isolates in the generally acid soils
where nodules were collected. Although a statistically significant
association occurred between the eight most common genomic
species and the 32 hosts, there was sufficient overlap in distributions
that no clear specificity between rhizobial genomic species and legume
taxa was observed. However, for three legume species, some preference
for particular genomic species was suggested. Similarly, no
geographical partitioning was found.
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INTRODUCTION |
The family Fabaceae is one of the
most successful families of angiosperms. It is the third largest, with
approximately 650 genera and 20,000 species (16), and is
most remarkable for its wide evolutionary diversification
(56) and cosmopolitan distribution (58). Many of
its members are of considerable agricultural or ecological importance,
generally reflecting their ability to develop symbiotic associations
with nitrogen-fixing soil bacteria, a feature which is widespread
within the family (1). The bacteria inducing nitrogen-fixing
nodules on leguminous plants correspond to five formally recognized
genera (34, 67); Rhizobium (23),
Sinorhizobium (9), Bradyrhizobium
(36), Azorhizobium (17), and
Mesorhizobium (34). These genera all belong to
the alpha subdivision of the proteobacteria but represent separate
lineages, relatively distant from one another and each more closely
related to nonnodulating taxa.
In Australia, the Fabaceae constitute a significant part of the
vascular flora, representing about 10% of the estimated 18,000 native
plant species (14). Several tribes (e.g., Mirbeliae and Bossiaeae) and a number of genera (e.g., Daviesia,
Bossiaea, and Pultenaea) of the family are
endemic. Native legumes are widely distributed throughout the
continent, occurring in all vegetation types except salt marshes and
marine aquatic communities (14). They are often a dominant
part of ecosystems in which they occur, whether this is measured in
terms of structural position, numbers, or overall biomass. This
dominance may reflect the advantage that legumes gain in soils of low
fertility (a characteristic feature of the majority of Australian
ecosystems) from symbiotic nitrogen-fixing associations with rhizobia.
In such situations, plant-microbial associations that help circumvent
nutrient deficiencies are likely to be of considerable significance in
determining the species and structural diversity of individual
ecosystems.
Paradoxically, relatively few studies have aimed to uncover the nature
of these bacterial symbionts in their native environments. A synthesis
of the work conducted in Australia over the past 40 years shows that
the comprehension of native rhizobia in the country is mainly based on
nodulation experiments and growth characteristics (14).
These two criteria are now held to be insufficient (28) and
can be misleading, e.g., slow-growing Mesorhizobium ciceri (51). The aim of the present study has been to improve our
knowledge of Australian native rhizobial diversity and to analyze the
influence of the nature of the associated host legume as well as
the geographic origin on the structure of the native rhizobial
communities. We tried to ensure that sampling was restricted to
rhizobial communities that were not invaded by strains exotic to
Australia (e.g., previously used as crop inoculants) by collecting
rhizobia at sites located in national parks or away from cropping
systems, and we focused on shrubby legumes composing the undercover in
woodland and forest ecosystems of southeastern Australia. A molecular
systematics approach combining small-subunit (SSU) ribosomal DNA (rDNA)
PCR-restriction fragment length polymorphism (RFLP) analysis
and sequencing was adopted to facilitate rapid identification of a
large number of strains.
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MATERIALS AND METHODS |
Nodule collection and isolation of bacterial strains.
Plants
of 32 legume species (Table 1) were
excavated at various field sites in southeastern Australia (Fig.
1) during spring and early summer for all
but the Island Bend site and for Gompholobium huegelii at
Lobs Hole, which were both sampled at the end of summer. At each site,
up to 10 individuals of a minimum of two legume species were sampled.
From these, segments of roots with attached nodules were excised and
transported in plastic bags to the laboratory, where bacterial strains
were isolated the following day. In the process, the nodules were
separated from the root, washed in distilled water, and then surface
sterilized following the technique of Cannon et al. (8) with
a nodule-sterilizing apparatus (25). The nodules were
crushed, and the exudate was streaked onto yeast-mannitol agar medium
(64). Pure cultures were obtained with one or more further
subculturing steps.
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FIG. 1.
Geographical location of the 12 sites where nodules were
collected: Ben Boyd National Park (BBNP), Black Mountain (BM), Boboyan
Road (BR), Gunning Road (GR), Island Bend (IB), Lowden Forest Park Road
(LFPR), Lobs Hole (LH), Mount Franklin (MF), Mundoonen Range (MR),
Tianjiara Falls (TF), Turpentine Road (TR), and Two Sticks Road (TSR).
Abbreviations: ACT, Australian Capital Territory; NSW, New South Wales;
VIC, Victoria.
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DNA preparation.
Bacterial DNA was prepared following the
method described by Sritharan and Barker (61). Bacteria were
grown on yeast-mannitol agar medium and colonies were collected,
suspended in 100 µl of 10 mM Tris (pH 8.0)-1 mM EDTA-1% Triton
X-100 solution, and boiled for 5 min. After a single chloroform
extraction, 5 µl of the supernatant was used in the amplification
reaction.
SSU rRNA gene amplification.
Primers corresponding to
positions 8 to 28 and 1492 to 1509 (39) in the
Escherichia coli SSU rRNA sequence (7) were used for amplification of the SSU rRNA genes by PCR. PCRs were carried out
in a 100-µl volume containing 5 µl of template DNA solution, 50 pmol of each of the two primers, 200 µM deoxynucleoside triphosphate (Boehringer Mannheim), and 2.5 U of Amplitaq DNA polymerase
(Perkin-Elmer) in Amplitaq DNA polymerase reaction buffer (10 mM
Tris-HCl [pH 8.3], 50 mM KCl, 1.5 mM MgCl2).
Amplifications were performed with a Hybaid Omnigene thermocycler with
the following temperature profile: an initial cycle consisting of a
denaturation step at 95°C for 5 min, an annealing step at 52°C for
120 s, and an extension step at 72°C for 90 s; 30 cycles of
denaturation at 94°C for 30 s, annealing at 52°C for 60 s, and extension at 72°C for 60 s; and a final extension step at
72°C for 5 min.
SSU rDNA PCR-RFLPs.
Aliquots of PCR products (10 µl) were
digested with restriction endonucleases as described by Laguerre et al.
(38). Nine restriction enzymes, AluI,
DdeI, HaeIII, HhaI, HinfI,
MspI, NdeII, RsaI, and TaqI
(New England Biolabs), were first used to screen a limited number of
bacterial strains. These had no greater discriminating power than a
combination of only four enzymes (HhaI, HinfI,
MspI, and RsaI) as observed by Laguerre et al.
(38). Restricted fragments were separated by electrophoresis
on 3% NuSieve 3:1 agarose gels at 80 V for 5 h and visualized by
ethidium bromide staining.
Nodulation tests.
The nodulation ability of the
representative strains of each PCR-RFLP genotype was verified by
inoculation onto the young plants of the original host grown from seeds
which had been collected at their corresponding site at the end of
summer. The seeds were dried over silica gel and surface sterilized in
concentrated H2SO4 for 20 min. After the acid
was drained, the seeds were washed thoroughly with 10 changes of
sterile water and placed in moistened steam-sterilized sand to
germinate at 22°C for 7 days. The seedlings were then planted in a
steam-sterilized sand-vermiculite mix in 12-cm-diameter pots and left
to establish in the greenhouse at 25°C. After 2 days, they were
inoculated at the base of the stem with 1 ml of the appropriate
bacterial inoculum (heavy suspension of the log-phase culture on
yeast-mannitol agar in 10 ml of N-free Jensen solution, pH 6.8). The
surface was covered with polyurethane beads to prevent evaporation and
contamination. The plants were grown in the greenhouse at 25°C and
watered with a sterile N-free nutrient solution twice each week. After
10 weeks of growth in the greenhouse, nodules were observed on the
seedling roots for all genomic species. Control uninoculated
plants were unnodulated.
PCR product sequencing.
Representative examples of isolates
possessing each of the distinct PCR-RFLP genotypes detected, with a
minimum of two for the most frequent genotypes, were used in a
subsequent sequence comparison. SSU rDNA PCR products were purified
with a Wizzard PCR Preps DNA purification system (Promega) as specified
by the manufacturer. The sequencing reaction was performed with the ABI PRISM dye terminator cycle-sequencing ready-reaction kit with Amplitaq
DNA polymerase FS as specified by the manufacturer, on an FTS-1 thermal
cycler (Corbett). Sequencing products were analyzed with an ABI
automatic sequencer model 377. Sense and antisense synthetic primers
complementary to conserved eubacterial domains corresponding to
positions 100 to 120, 243 to 263, 343 to 357, 518 to 536, 685 to 704, 787 to 803, 907 to 926, 1100 to 1115, 1224 to 1241, and 1385 to 1401 in
the E. coli SSU rRNA sequence (7) were used to
sequence both strands of the SSU rRNA gene.
Sequence analysis.
The SSU rRNA gene sequences were aligned
manually by comparison with a database of alpha proteobacteria SSU rRNA
sequences aligned on the basis of their phylogenetic relationships by
using the program VSM 4.0 for SSU rRNA sequence database management (11). All of the sites were included in the phylogenetic
analysis, except in the case of Bradyrhizobium species
analysis, for which a short stretch of the sequences (positions 997 to
1041 in the E. coli SSU rRNA sequence [7])
was excluded. Phylogenetic analyses were performed by the
neighbor-joining method (59) with the program NEIGHBOR in
PHYLIP version 3.5c (21). Distances were computed with
DNADIST under the Jin and Nei distance (35). One thousand
bootstrap replications were performed with SEQBOOT. The graphic
manipulation of the tree was realized with NJplot (55).
Nucleotide sequence accession numbers.
The SSU rRNA gene
sequences corresponding to the rhizobial genomic species
identified have been deposited in the EMBL nucleotide database under
accession no. Z94803 to Z94823. The accession numbers of the nucleotide
sequences of the SSU rRNA genes of the Rhizobiaceae and
related alpha proteobacteria used for comparison are as follows:
Afipia clevelandensis, M69186; Afipia felis, M65248; Agrobacterium rhizogenes LMG152, X67224;
Agrobacterium tumefaciens Ch-Ag-4, D14505; Agromonas
oligotrophica, D78366; Blastobacter denitrificans,
X66025; Bradyrhizobium elkanii, U35000; Bradyrhizobium
japonicum USDA 6T, U69638, and USDA 110, D13430;
Bradyrhizobium spp. 129, D14508; 55S, D14507; LMG 9514, X70401; LMG 9520, X70403; LMG 9580, X70404; LMG 9966, X70403; LMG
10698, X70405; Brucella melitensis, L26166;
Mesorhizobium ciceri, U07934; Mesorhizobium
huakuii, D12797; Mesorhizobium loti A, X67229, and B,
X67230; Mesorhizobium mediterraneum, L38825;
Mesorhizobium spp. LMG7836, XZ68389, and LMG7854, X68391;
Mesorhizobium tianshanense, U71079; Mycoplana
dimorpha, D12786; Ochrobactrum anthropi, D12794;
"Photorhizobium" thompsonianum, L23405;
Phyllobacterium myrsinacearum, D12789; Phyllobacterium
rubiacearum, D12790; Rhizobium etli, U28939;
Rhizobium gallicum, U86343; Rhizobium hainanense,
U71078; Rhizobium leguminosarum, X67227; Rhizobium tropici, D12798; Rhizobium sp. LMG 9509, X67232;
Rhodopseudomonas palustris, D25312; and Zoogloea
ramigera, X74915.
Statistical analyses.
Association between rhizobium
genomic species and hosts, or between rhizobium genomic
species and sites, was tested by using the log-likelihood ratio
statistic (15). This test has been shown to perform well for
large sample sizes, such as those in our study (37), even
when there are low expected numbers for some combinations. The nature
of the associations was then examined by correspondence analysis
(29).
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RESULTS |
SSU rDNA PCR-RFLP study.
A total of 745 strains, representing
all the strains isolated from all the hosts, were included in the RFLP
study. Gel electrophoresis of the PCR products revealed that the
amplification reaction produced a single DNA molecule slightly
less than 1.5 kb long for all strains. Four restriction endonucleases,
HhaI, HinfI, MspI, and
RsaI, were used to characterize the whole collection. From 4 to 10 distinct restriction patterns were detected with each of these
enzymes. The combination of the four patterns identified 21 SSU rDNA
types that we arbitrarily named A to U.
SSU rDNA sequence analyses.
SSU rRNA gene sequences of the 21 rRNA genomic species were aligned by comparison with a database
containing about 500 aligned SSU rRNA sequences of alpha
proteobacteria. Phylogenetic analyses including representatives of all
rhizobial genera and related alpha proteobacteria revealed that all 21 SSU rRNA genomic species detected belonged to the
Rhizobium-Agrobacterium group as defined in the Ribosomal
Database Project (45). Sixteen of the SSU rRNA genomic species clustered within the
Bradyrhizobium subgroup, and three (genomic
species S, T, and U) grouped within the Mesorhizobium subgroup, while the remaining two (genomic species Q and R)
grouped within the R. leguminosarum subgroup.
Genomic species were further studied according to the subgroup to which
they were related. In each case, outgroups were chosen
as the most
closely related species in accordance with the Ribosomal
Database
Project general phylogeny of procaryotes. Within the
R. leguminosarum subgroup, genomic species Q and R clustered
with
R. tropici (Fig.
2A), Q being much more closely related to
R. tropici (with a difference of one base between their
SSU rDNA
sequences) than to genomic species R (with a
difference of 40
bases). Genomic species S and species T and U
formed two individualized
lineages which were each clearly affiliated
with one of the two
major groups within the
Mesorhizobium
subgroup (Fig.
2B). Genomic
species S was closely related to the
M. loti-M. ciceri cluster,
its SSU rDNA differing from
M. loti A and B sequences by four
and three bases,
respectively, and differing from
M. ciceri SSU
rRNA by six
bases. The lineage formed by genomic species T and
U clustered
with
M. huakuii, from which they differed by four
and two
bases, respectively, at the SSU rDNA level. Their SSU
rDNA sequences
differed from each other by only two bases. In
both analyses, internal
branches linking the genomic species to
known rhizobial species
were supported above the 70% level by
1,000 bootstrap replications and
are thus expected to represent
true clades according to Hillis and Bull
(
32).
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FIG. 2.
Phylogenetic relationships among genomic species
belonging to the genera Rhizobium and
Mesorhizobium characterized by SSU rDNA PCR-RFLPs. The
phylogenetic trees were constructed by the neighbor-joining
method. The numbers correspond to the percentage of bootstrap
support for internal branches, based on 1,000 replications. The scale
bar corresponds to 0.005 substitution per site. (A) Phylogenetic
positions of genomic species Q and R within the
R. leguminosarum subgroup. (B) Phylogenetic positions
of genomic species S, T, and U within the
Mesorhizobium subgroup.
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The phylogenetic positions of 16 genomic species within the
Bradyrhizobium subgroup were investigated by comparing their
SSU
rDNA sequences to those of representatives of the various genera
within this group. Numerous sequences are available in the DNA
sequence databases for
B. japonicum; the choice of
representative
SSU rRNA sequences for this species was made according
to the
results of Barrera et al. (
6). The resulting
phylogenetic tree
(Fig.
3) exhibited a
poor level of resolution for some of the
internal branches, due to the
small divergence between the various
sequences. Consequently, the
phylogenetic position of the rhizobial
genomic species
identified in the PCR-RFLP study, in particular
for genomic
species E, G, and H, as well as the branching order
of the various
groupings, could not be resolved. With the exception
of L and P,
none of the genomic species identified in this study
clustered
with any of the known species of this group at a significant
level by
1,000 bootstrap replications. Eleven of them (A, B, C,
D, F, I, J, K,
M, N, and O) formed a group of closely related
species, relatively
distant from any known species. The sequences
of any two of these
genomic species exhibited a very high degree
of similarity: the
minimum difference was one base (genomic species
A and M); the
maximum was 18 bases (genomic species B and F),
which
represents less than 4% of the sequence. Genomic species
L and P
clustered with the
B. elkanii cluster, which is the only
one
supported at a significant level by 1,000 bootstrap replications.
SSU
rDNAs of all the species included in this cluster present
a
characteristic sequence from positions 997 to 1041, according
to
E. coli SSU rRNA sequence numbering (
7), which
was also
found in the genomic species L and P sequences. This
part of the
SSU rDNA was not included in the construction of the
phylogeny
presented in Fig.
3 (see Discussion). The other three
genomic
species, E, G, and H, did not show a significant
phylogenetic
affinity for any particular branch and thus are likely to
constitute
separate lineages within the
Bradyrhizobium
subgroup.
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FIG. 3.
Phylogenetic relationships among genomic species
belonging to the genus Bradyrhizobium characterized by SSU
rDNA PCR-RFLPs. The phylogenetic tree was constructed by the
neighbor-joining method. The numbers correspond to the percentage of
bootstrap support for internal branches, based on 1,000 replications.
The scale bar corresponds to 0.002 substitution per site.
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Analysis of rhizobial diversity.
The distribution of the 745 strains among the various rhizobial genomic species was highly
unbalanced (Table 2). Most of them
(94.3%) were Bradyrhizobium species, with one
genomic species, A, representing more than half of the total
number of strains (57.6%). Among the 21 genomic species, only
8 constituted 97.05% of the entire collection. The remaining 13 genomic species each represented less than 1% of the strains
and, in most cases, were only isolated once. Various combinations
of rhizobial genomic species occurring on the same host plant
were observed, most of them logically involving genomic species
A strains. Rhizobium and Mesorhizobium
genomic species were found on 14% of the plants hosting a
minimum of two nodules and always co-occurred with various Bradyrhizobium species.
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TABLE 2.
Distribution of 745 rhizobial isolates among 21 genomic species identified by RFLP analysis of PCR-amplified
SSU rRNA genes
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Frequencies of the various rhizobial genomic species were
calculated for each host summed across all sites (Table
3) and
for each site regardless of their
host origin (Table
4). All
genomic species isolated more than once were found on several
hosts and at more than one site, including species J, which was
isolated only twice. For all but one host and at all sites, one
genomic species dominated the rhizobial community. At 10 of 12
sites and for most hosts (21 of 32) this was genomic species A.
Genomic species P dominated among the rhizobia found on four of
the
nine host species from Ben Boyd National Park site (
Aotus ericoides,
Dillwynia glaberrima,
Dillwynia
sericea, and
Pultenaea daphnoides), and B was the
dominant genomic species isolated on
the roots of four of the
five hosts collected exclusively at Lobs
Hole site (
Daviesia
latifolia,
Gompholobium huegelii,
Hovea
linearis,
Hovea purpurea, and
Mirbelia
oxylobioides). Genomic species co-occurring
either on plants of
the same host or at the same site were each
present at much lower
frequencies than the dominant rhizobial
species and were generally
widely distributed among a number of
hosts and occurred at several
sites. Only in the case of three
minor genomic species (D, F,
and H) was one particular host predominantly
nodulated (Table
3).
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TABLE 3.
Frequenciesa of 21 rhizobial
genomic species among legume host species from which nodules
were collected
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The 13 minor rhizobial genomic species were excluded from the
association analyses because of the paucity of data. There was
a highly
significant association between the remaining eight rhizobial
genomic species and both the 32 hosts (maximum-likelihood
chi-square
= 790.79 with 217 df;
P < 0.001) and
the 12 sampling sites (maximum-likelihood
chi-square = 973.98 with
77 df;
P < 0.001). This association was
clearly
visible on the projection of rhizobial genomic species
and
either legume hosts or sampling sites along the two first
axes
generated by correspondence analyses (Fig.
4A and
B). Hosts
for which several rhizobial
genomic species were relatively abundant
had an intermediate
position between the two or three major rhizobial
species, such as
Bossiaea foliosa,
Platylobium formosum, and
G. huegelii between A and B;
Daviesia leptophylla
and
Daviesia buxifolia between A and D; and
Pultenaea capitellata between A, and F and
P (Fig.
4A).
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FIG. 4.
Position of the eight more-abundant rhizobial
genomic species and legume hosts or sites along the
first and second principal axes. (A) Association between
genomic species (solid circles) and legume hosts (open
circles). Abbreviations: Ao, A. obliquinervia; Ae,
A. ericoides; Bb, Bossiaea buxifolia; Be,
Bossiaea ensata; Bf, B. foliosa; Db, D. buxifolia; Dla, D. latifolia; Dle, D. leptophylla; Dm, Daviesia mimosoides; Du, D. ulicifolia; Dwb, Dillwynia brunioides; Dwg, D. glaberrima; Dwra, Dillwynia ramosissima; Dwre,
Dillwynia retorta; Dws, D. sericea; Gh, G. huegelii; Gl, G. lotifolia; Hv, Hardenbergia
violacea; Hl, H. linearis; Hp, H. purpurea; Ia, Indigofera australis; Mo, M. oxylobioides; Mr, Mirbelia rubiifolia; Oe,
Oxylobium ellipticum; Php, P. phylicoides; Pf, P. formosum; Poa, Podolobium
alpestre; Poi, Podolobium ilicifolium; Pc, P. capitellata; Pd, P. daphnoides; Pp, Pultenaea
procumbens; Ps, Pultenaea scabra. (B) Association
between genomic species (solid circles) and legume
hosts (open triangles). Abbreviations: BBNP, Ben Boyd National Park;
BM, Black Mountain; BR, Boboyan Road; GR, Gunning Road; IB, Island
Bend; LFPR, Lowden Forest Park Road; LH, Lobs Hole; MF, Mount Franklin;
MR, Mundoonen Range; TF, Tianjiara Falls; TR, Turpentine Road; TSR,
Two Sticks Road.
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The number and frequency of rhizobial genomic species varied
among host species belonging to the same genus (Table
3). Although
genomic species A was prevalent for all species of
Bossiaea and
Pultenaea, the relative distribution
of the rhizobia between A
and all other genomic species varied
significantly from one taxon
to another within either genus
(chi-square = 41.4 with 18 df [
P < 0.005] and
chi-square = 32.3 with 15 df [
P < 0.001],
respectively).
In contrast, in the case of the two
Hovea
species, this was not
significant (chi-square = 4.1 with 3 df;
P = 0.25 [B is the prevalent
rhizobial genomic
species in this case]). The genera
Daviesia,
Dillwynia, and
Mirbelia exhibited species
differences in both
the predominant rhizobial genomic
species and the distribution
of rhizobial genomic species. No
clear specificity could be observed
at a higher taxonomic rank between
the two subfamilies represented
among the sampled hosts. The strains
isolated from the only member
of the subfamily Mimosoideae represented
in the present sample,
Acacia obliquinervia, belonged to two
of the most abundant rhizobial
genomic species, A and F, also
isolated from various Papilionoideae
species. For this host, however, F
represented more than half
of the strains (58.3%) and A represented
only 16.7%.
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DISCUSSION |
Rhizobial diversity.
Early reports indicated that symbionts of
native legumes in Australia were typical slow-growing bacteria with the
characteristics of Bradyrhizobium species (31, 40, 47,
50). Later studies, however, have suggested a higher level of
diversity. Ninety-eight percent of the strains isolated from acacias at
Fowlers Gap, near Broken Hill in arid northwestern New South Wales
(NSW), were fast growers (3), and slow- and fast-growing
rhizobia were shown to occur in temperate Australia (4, 41,
63). Nevertheless, a high predominance of
Bradyrhizobium species has generally been observed in
temperate southeastern Australia (3, 4, 41, 63) as well as
in other parts of Australia, e.g., Queensland (57) and
Western Australia (40). Barnet and Catt (3)
isolated atypical very slow growing rhizobia with high host specificity in the alpine area of Kosciusko National Park. Very slow growing rhizobia were also isolated at Bridge Hill near Bulahdelah
(4). In contrast, Barnet et al. (5) showed that
fast-growing rhizobia isolated from Acacia spp. in NSW were
diverse and belonged to various Rhizobium species,
suggesting that some represented a new genus more closely related
to Bradyrhizobium than to Rhizobium. Unfortunately, the various previous descriptions of rhizobia occurring on Australian native plants were based solely on growth features and
the cross-inoculation concept and thus do not provide precise information on the real nature and structure of the rhizobial communities in Australian ecosystems.
From these various studies, and as the range of sites studied in the
past was limited, we expected to encounter high diversity
among
indigenous rhizobial strains in Australia. Among the 745
strains that
we typed, 21 rhizobial genomic species were identified.
This might seem low; however, whereas the results of SSU rDNA
analyses clearly show differences between strains, SSU rDNA is
not appropriate for the formal delineation of species (
44,
65).
Stackebrandt and Goebel (
62) showed that
two procaryotes are
unlikely to have more than 60 to 70%
DNA similarity, and hence
to be related at the species level, when
their SSU rDNA sequences
have less than 97% homology but that above
97% SSU rDNA homology,
the DNA similarity can vary greatly, from 10 to
100%. Thus, a
very high SSU rDNA similarity, as high as 99.8%, can be
observed
for different species (
22). In contrast,
heterogeneity between
SSU rDNA sequences has been documented
within the seven rRNA operons
of
E. coli
(
12). It will thus be necessary to perform DNA-DNA
hybridization and thermal denaturation analyses, which constitute
the
criteria commonly used to define bacterial species (
65),
to
evaluate the full extent of taxonomic diversity among our strains.
Nevertheless, with one exception, none of these genomic species
corresponded to previously described rhizobia
a pattern that has
generally been observed in other studies of wild rhizobial communities
(
10,
18,
49,
53,
68) and which suggests a very high
level of
diversity within the rhizobium taxonomy.
Geographical localization.
Barnet and Catt
(3) found marked geographical localization of the
various rhizobial types according to their rates of growth: fast
growers in arid northwestern NSW, typical Bradyrhizobium at
the two distant coastal heath areas (Myall Lakes National Park and
Wanda Beach) and a rain forest site in Blue Mountains National Park,
and slow and very slow growers in the alpine area of Kosciusko National
Park. In contrast, other studies of rhizobial diversity in other parts
of the world failed to identify a geographical specificity of
particular rhizobial types (52, 68). We did not observe such
a geographical partitioning of the various genomic species, and
we found no difference in the geographic distribution of
Rhizobium, Mesorhizobium, and
Bradyrhizobium species (we have to assume that fast growers
identified by Barnet and Catt belong to Rhizobium and
slow growers belong to Bradyrhizobium). On the contrary,
most genomic species were found at several sites, even in the
case of the species isolated on just a few occasions (e.g., genomic species E, J, and O). In general, one prevalent
genomic species was recovered from a particular site, with a
number of additional species present at much lower frequencies. Our
sampling, however, did not cover the same climatic range as the Barnet
and Catt study. A study conducted a few years ago at Mount Cootha in
Queensland investigated the diversity of soil bacterium communities by
using a molecular approach, in which partial SSU rDNA sequences (about
250 bases long) were generated from DNA directly extracted from
the soil (42). A phylogenetic analysis including members of
the alpha proteobacteria division revealed that some of the clones
belonged to the Rhizobium-Agrobacterium group. None was strictly identical to any SSU rRNA sequence already available in
nucleic acid databases at that time. When compared to our sequences, two clones, MC6 and MC23 (accession no. X65573 and X65578 in the
GenBank/EMBL/DDJB DNA sequence database), showed perfect identity
with the corresponding parts of the sequences of genomic species G and L, a difference of one base with species H, and a
difference of two bases with species P and E. In our phylogenetic analysis (Fig. 3), P and L clustered with the B. elkanii
clade. E, G, and H did not show any affinity to any particular
previously characterized lineage within the
Bradyrhizobium-Rhodopseudomonas subgroup and thus are
expected to constitute a new lineage (genus?) within this group.
These results can only be indicative, since a different phylogeny can
sometimes be obtained when the complete SSU rRNA gene is considered, as
in the case of Rhizobium galegae (51, 66) or
Rhizobium etli (46). However, it appears that there are strong similarities between rhizobial communities at the
sites that we sampled and that at the distant and climatically different site in Queensland. Species that we identified in the temperate zone thus appear to be widespread geographically under very
different climate conditions.
All 12 sites presented some degree of diversity of vegetation and soil
characteristics, but all had acid or near-neutral soil,
conditions
which favor
Bradyrhizobium species over
Rhizobium,
Mesorhizobium, or
Sinorhizobium species (
27,
50). Bradyrhizobia
can
survive at low pH, which is not the case for most strains
of the other
rhizobial genera (
27). The predominance of
Bradyrhizobium species among our isolates is thus not
surprising, as is the presence
of slow-growing strains in similar sites
(Kosciusko National Park,
Wanda Beach, Myall Lakes National Park, and
Blue Mountains National
Park) studied by Barnet and Catt
(
3). Among the
Rhizobium and
Mesorhizobium species that we isolated, the most abundant
(
R. tropici) was also one of the more acid tolerant
(
27). It is
thus likely that there is a relation between the
level of soil
acidity and the nature of the rhizobial species present
at our
sampling sites. This had also been observed in Africa for
Rhizobium species nodulating
Phaseolus vulgaris.
An apparently similar degree
of diversity was found at two sites
with different soil pH levels.
However,
R. tropici
predominated at the acid soil site, and
R. etli
predominated at the site with a near-neutral soil (
2,
26).
The apparent geographical specificity described by Barnet
and
Catt (
3) certainly reflects the lack of resolution of the
rhizobium identification methodology applied but also reflects
pH
differences between the sites.
Host specificity.
We did not observe any clear host
specificity at either the host species or genus level between any
particular rhizobial species and its leguminous host, as was previously
reported by several authors (24, 33, 48, 68). Likewise, no
clear specificity could be seen at a higher taxonomic rank or
within a particular plant. In particular, Rhizobium and
Mesorhizobium genomic species were never found to
nodulate exclusively either a particular host or a particular plant.
This co-occurrence of slow- and fast-growing rhizobia on the same host
genus or species appears to happen quite commonly (24, 43,
49, 54). Even when several legume species occurred at a
site, in most cases, all of the species were predominantly nodulated
with the commonest rhizobial species found at that site. Clearly,
in very many cases, this involved rhizobial genomic species A. However, without further detailed testing, this association cannot simply be ascribed to a generally better "fit" of
genomic species A to all legumes as, even at sites dominated by
other genomic species (e.g., Lobs Hole with
genomic species B and Gunning Road with genomic species
Q), genomic species A was also present.
In contrast, in the cases of
A. obliquinervia,
Goodia
lotifolia, and
Phyllota phylicoides, we did observe
some suggestion
of preference for particular rhizobial species.
Different rhizobial
genomic species were recovered from their
root nodules, but the
dominant species isolated (which was different
for each of the
three species) also differed from the prevalent species
isolated
from nodules on other legume hosts occurring at the same
sites.
Thus, at Island Bend, where genomic species A was
predominant
on
B. foliosa and
Daviesia
ulicifolia, almost 60% of isolates
recovered from
A. obliquinervia were of genomic species F. Similarly,
for
G. lotifolia at Lowden Forest Park Road the dominant species
was H, and for
P. phylicoides at Tianjiara Falls it was D,
although
genomic species A was commonest on all co-occurring
species (two
and four species, respectively). Furthermore, the
correspondence
analyses (Fig.
4A and B) indicated stronger links in the
host-rhizobium
comparisons than in the site-rhizobium comparisons for
the genomic
species prevalent on each of these three legume
species. The differences
observed for
A. obliquinervia,
G. lotifolia, and
P. phylicoides are consistent
with the suggestion that one rhizobial species
is being selected by
these legume hosts regardless of the apparently
most abundant rhizobial
species at those sites. These three host
species could only be sampled
at one site each, and our results
need confirming by further sampling
at additional sites. However,
it is of particular interest to note that
A. obliquinervia is
the only member of the subfamily
Mimosoideae from which nodules
were isolated for the present
study, all others belonging to the
Papilionoideae. This
might indicate a specificity difference between
the Fabaceae
subfamilies. Further investigation is needed to evaluate
this
observation.
Considering the broad range of specificities either of rhizobial
species towards their hosts or of the legume species towards
their
symbionts, molecular identification appears to be a prerequisite
to any
study of rhizobial population structure. Indeed, it is
fundamental to
differentiate between members of the same species
and members of a
group of species, to be able to provide some
insight on the
relationship between the two partners, and to infer
the factors
determining the legume-rhizobium symbiotic association.
Interestingly, species of the
B. elkanii cluster
can be differentiated
from other
Bradyrhizobium species by a
short part of their SSU
rRNA gene sequence, corresponding to a highly
variable part of
the molecule (
30). Although it was fully
conserved between sequences
of species within the
B. elkanii subgroup, it was highly divergent
from that of other
species in the
Bradyrhizobium subgroup, to
the extent that
no homology could be safely identified. The other
species, in turn,
were characterized by a unique sequence. In
contrast, the
B. elkanii subgroup "signature" sequence showed
a reasonably good
level of similarity to SSU rDNA sequence from
species of the
Mesorhizobium subgroup, which represents a comparatively
distant lineage. The occurrence of recombination in SSU rRNA genes
has
been documented in
Aeromonas (
60), as well as
among
Rhizobium and
Agrobacterium species
(
19,
20); it is thus possible that
SSU rDNA of the ancestor
of the
B. elkanii cluster evolved by
recombination between
distant lineages, possibly representing
the same lifestyle. Further
analyses of rRNA genes might bring
insight into the mode of evolution
of the different rhizobial
lineages and the emergence and spreading of
nodulation ability.
| |
ACKNOWLEDGMENTS |
This work was supported by a CSIRO multidivisional program for
the study of Australian biodiversity.
We thank W. J. Müller for advising us in the statistical
analyses, T. Lally for assistance in the field, and M. Woods for technical assistance in the laboratory and greenhouse. We are grateful
to ACT Parks and Conservation Service and NSW National Parks and
Wildlife Service for permission to collect material in areas under
their jurisdiction. We are grateful to three anonymous reviewers whose
constructive criticisms contributed to improvement of the manuscript.
| |
FOOTNOTES |
*
Corresponding author. Present address: Department of
Genetics, University of Nottingham, Queens Medical Centre, Nottingham NG7 2UH, United Kingdom. Phone: 44 (0)115 924 9924, ext. 42598. Fax: 44 (0)115 970 9906. E-mail: bene{at}evol.nott.ac.uk.
| |
REFERENCES |
| 1.
|
Allen, O. N., and E. K. Allen.
1981.
The Leguminosae: a source book of characteristics, uses, and nodulation.
University of Wisconsin Press, Madison, Wis.
|
| 2.
|
Anyango, B.,
K. J. Wilson,
J. L. Beynon, and K. E. Giller.
1995.
Diversity of rhizobia nodulating Phaseolus vulgaris L. in two Kenyan soils with contrasting pHs.
Appl. Environ. Microbiol.
61:4016-4021[Abstract].
|
| 3.
|
Barnet, Y. M., and P. C. Catt.
1991.
Distribution and characteristics of root-nodule bacteria isolated from Australian Acacia spp.
Plant Soil
135:109-120.
|
| 4.
|
Barnet, Y. M.,
P. C. Catt, and D. H. Hearne.
1985.
Biological nitrogen fixation and root-nodule bacteria (Rhizobium sp. and Bradyrhizobium sp.) in two rehabilitating sand dune areas planted with Acacia spp.
Aust. J. Bot.
33:595-610.
|
| 5.
|
Barnet, Y. M.,
P. C. Catt,
R. Jenjareontham, and K. Mann.
1992.
Fast-growing root-nodule bacteria from Australian Acacia, p. 594.
In
R. Palacios, J. Mora, and W. E. Newton (ed.), New horizons in nitrogen fixation. Kluwer Academic Publishers, Dordrecht, The Netherlands.
|
| 6.
|
Barrera, L. L.,
M. E. Trujillo,
M. Goodfellow,
F. J. García,
I. Hernández-Lucas,
G. Dávila,
P. van Berkum, and E. Martínez-Romero.
1997.
Biodiversity of bradyrhizobia nodulating Lupinus spp.
Int. J. Syst. Bacteriol.
47:1086-1091[Abstract/Free Full Text].
|
| 7.
|
Brosius, J.,
M. L. Palmer,
P. J. Kennedy, and H. F. Noller.
1978.
Complete nucleotide sequence of a 16S ribosomal RNA gene from Escherichia coli.
Proc. Natl. Acad. Sci. USA
75:4801-4805[Abstract/Free Full Text].
|
| 8.
|
Cannon, J. R.,
N. H. Corbett,
J. Brockwell,
A. H. Gibson, and G. A. McIntyre.
1967.
Nodulation and growth of Trifolium subterraneum L. cv Mount Barker in agar culture.
Aust. J. Biol. Sci.
20:285-295.
|
| 9.
|
Chen, W. X.,
G. H. Yan, and J. L. Li.
1988.
Numerical taxonomic study of fast-growing soybean rhizobia and a proposal that Rhizobium fredii be assigned to Sinorhizobium gen. nov.
Int. J. Syst. Bacteriol.
38:392-397[Abstract/Free Full Text].
|
| 10.
|
Chen, W.-X.,
Z.-Y. Tan,
J.-L. Gao,
Y. Li, and E.-T. Wang.
1997.
Rhizobium hainanense sp. nov., isolated from tropical legumes.
Int. J. Syst. Bacteriol.
47:870-873[Abstract/Free Full Text].
|
| 11.
|
Christen, R.
1995.
VSM 4.0. A general databasing set of programs for managing ribosomal RNA sequences for molecular phylogeny, taxonomy and ecology.
http://www.obs-vlfr.fr/christen/Index.html.
|
| 12.
|
Cilia, V.,
B. Lafay, and R. Christen.
1996.
Sequence heterogeneities among 16S ribosomal RNA sequences, and their effect on phylogenetic analyses at the species level.
Mol. Biol. Evol.
13:451-461[Abstract].
|
| 13.
|
Crisp, M. D., and P. H. Weston.
1987.
Cladistic and legume systematics, with an analysis of the Bossiaeae, Brongniartieae and Mirbelieae, p. 62-130.
In
C. H. Stirton (ed.), Advances in legume systematics, part 3. Royal Botanic Garden, Kew, England.
|
| 14.
|
Davidson, B. R., and H. F. Davidson.
1993.
Leguminosae (Fabaceae) and Rhizobiaceae, p. 47-68.
In
B. R. Davidson, and H. F. Davidson (ed.), Legumes: the Australian experience. The botany, ecology and agriculture of indigenous and immigrant legumes. Research Studies Press Ltd, Taunton, Somerset, England.
|
| 15.
|
Dobson, A. J.
1990.
An introduction to generalized linear models.
Chapman and Hall, London, England.
|
| 16.
|
Doyle, J. J.
1994.
Phylogeny of the legume family: an approach to understanding the origins of nodulation.
Annu. Rev. Ecol. Syst.
25:325-349.
|
| 17.
|
Dreyfus, B.,
J. L. Garcia, and M. Gillis.
1988.
Characterization of Azorhizobium caulinodans gen. nov., sp. nov., a stem-nodulating nitrogen-fixing bacterium isolated from Sesbania rostrata.
Int. J. Syst. Bacteriol.
38:89-98.
|
| 18.
|
Dupuy, N.,
A. Willems,
B. Pot,
D. Dewettinck,
I. Vandenbruaene,
G. Maestrojuan,
B. Dreyfus,
K. Kersters,
M. D. Collins, and M. Gillis.
1994.
Phenotypic and genotypic characterization of bradyrhizobia nodulating the leguminous tree Acacia albida.
Int. J. Syst. Bacteriol.
44:461-473[Abstract/Free Full Text].
|
| 19.
|
Eardly, B. D.,
F. S. Wang, and P. van Berkum.
1996.
Corresponding 16S rRNA gene segments in Rhizobiaceae and Aeromonas yield discordant phylogenies.
Plant Soil
186:69-74.
|
| 20.
|
Eardly, B. D.,
F. S. Wang,
T. S. Whittam, and R. K. Selander.
1995.
Species limits in Rhizobium populations that nodulate the common bean (Phaseolus vulgaris).
Appl. Environ. Microbiol.
61:507-512[Abstract].
|
| 21.
|
Felsenstein, J.
1993.
PHYLIP (Phylogeny Inference Package) version 3.5c.
Department of Genetics, University of Washington, Seattle.
|
| 22.
|
Fox, G. E.,
J. D. Wisotzkey, and P. J. Jurtshuk.
1992.
How close is close: 16S rRNA sequence identity may not be sufficient to guarantee species identity.
Int. J. Syst. Bacteriol.
42:166-170[Abstract/Free Full Text].
|
| 23.
|
Frank, B.
1889.
Uber dir Pilzsymbiose der Leguminosen.
Ber. Dtsch. Bot. Ges.
7:332-346.
|
| 24.
|
Gao, J. L.,
J. G. Sun,
Y. Li,
E. T. Wang, and W. X. Chen.
1994.
Numerical taxonomy and DNA relatedness of tropical rhizobia isolated from Hainan Province, China.
Int. J. Syst. Bacteriol.
44:151-158[Abstract/Free Full Text].
|
| 25.
|
Gault, R. R.,
P. T. Byrne, and J. Brockwell.
1973.
Apparatus for surface sterilization of individual legume root nodules.
Lab. Pract.
22:292-293.
|
| 26.
|
Giller, K. E.,
B. Anyango,
J. L. Beynon, and K. J. Wilson.
1994.
The origin and diversity of rhizobia nodulating Phaseolus vulgaris L. in African soils, p. 57-62.
In
J. I. Sprent, and D. McKey (ed.), Advances in legumes systematics, part 5. The nitrogen factor. Royal Botanic Garden, Kew, England.
|
| 27.
|
Graham, P. H.,
K. J. Draeger,
M. L. Ferrey,
M. J. Conroy,
B. E. Hammer,
E. Martínez,
S. R. Aarons, and C. Quinto.
1994.
Acid pH tolerance in strains of Rhizobium and Bradyrhizobium, and initial studies on the basis for acid tolerance of Rhizobium tropici UMR1899.
Can. J. Microbiol.
40:198-207.
|
| 28.
|
Graham, P. H.,
M. J. Sadowsky,
H. H. Keyser,
Y. M. Barnet,
R. S. Bradley,
J. E. Cooper,
D. J. De Ley,
B. D. W. Jarvis,
E. B. Roslycky,
B. W. Stijdom, and J. P. W. Young.
1991.
Proposed minimal standards for the description of new genera and species of root- and stem-nodulating bacteria.
Int. J. Syst. Bacteriol.
41:582-587.
|
| 29.
|
Greenacre, M. J.
1984.
Theory and applications of correspondence analysis.
Academic Press, London, England.
|
| 30.
|
Gutell, R. R.
1994.
Collection of small subunit (16S- and 16S-like) ribosomal RNA structures.
Nucleic Acids Res.
22:3502-3507[Abstract/Free Full Text].
|
| 31.
|
Hannon, N. J.
1949.
The nitrogen economy of the Hawkesbury sandstone soils around Sydney. The role of native legumes. Honours thesis.
University of Sydney, Sydney, Australia.
|
| 32.
|
Hillis, D. M., and J. J. Bull.
1993.
An empirical test of bootstrapping as a method for assessing confidence in phylogenetic analysis.
Syst. Biol.
42:182-192.
|
| 33.
|
Jarvis, B. D. W.
1983.
Genetic diversity of Rhizobium strains which nodulate Leucaena leucocephala.
Curr. Microbiol.
8:153-158.
|
| 34.
|
Jarvis, B. D. W.,
P. Van Berkum,
W. X. Chen,
S. M. Nour,
M. P. Fernandez,
J. C. Cleyet-Marel, and M. Gillis.
1997.
Transfer of Rhizobium loti, Rhizobium huakuii, Rhizobium ciceri, Rhizobium mediterraneum, and Rhizobium tianshanense to Mesorhizobium gen. nov.
Int. J. Syst. Bacteriol.
47:895-898[Abstract/Free Full Text].
|
| 35.
|
Jin, L., and M. Nei.
1990.
Limitations of the evolutionary parsimony method of phylogenetic analysis.
Mol. Biol. Evol.
7:82-102[Abstract].
|
| 36.
|
Jordan, D. C.
1982.
Transfer of Rhizobium japonicum Buchanan 1980 to Bradyrhizobium gen. nov., a genus of slow-growing, root nodule bacteria from leguminous plants.
Int. J. Syst. Bacteriol.
32:136-139[Abstract/Free Full Text].
|
| 37.
|
Koehler, K. J., and K. Larntz.
1980.
An empirical investigation of goodness-of-fit statistics for sparse multinomials.
J. Am. Stat. Assoc.
75:336-344.
|
| 38.
|
Laguerre, G.,
M.-R. Allard,
F. Revoy, and N. Amarger.
1994.
Rapid identification of rhizobia by restriction fragment length polymorphism analysis of PCR-amplified 16S rRNA genes.
Appl. Environ. Microbiol.
60:56-63[Abstract/Free Full Text].
|
| 39.
|
Lane, D. J.,
B. Pace,
G. J. Olsen,
D. A. Stahl,
M. L. Sogin, and N. R. Pace.
1985.
Rapid determination of 16S ribosomal RNA sequences for phylogenetic analyses.
Proc. Natl. Acad. Sci. USA
82:6955-6959[Abstract/Free Full Text].
|
| 40.
|
Lange, R. T.
1961.
Nodule bacteria associated with the indigenous Leguminosae of southwestern Australia.
J. Gen. Microbiol.
26:351-359.
|
| 41.
|
Lawrie, A. C.
1983.
Relationships among rhizobia from native Australian legumes.
Appl. Environ. Microbiol.
45:1822-1828[Abstract/Free Full Text].
|
| 42.
|
Liesack, W., and E. Stackebrandt.
1992.
Occurrence of novel groups of the domain Bacteria as revealed by analysis of genetic material isolated from an Australian terrestrial environment.
J. Bacteriol.
174:5072-5078[Abstract/Free Full Text].
|
| 43.
|
Lim, G., and H. L. Ng.
1977.
Root nodules of some tropical legumes in Singapore.
Plant Soil
46:317-327.
|
| 44.
|
Lindström, K.,
P. van Berkum,
M. Gillis,
E. Martínez,
N. Novikova, and B. Jarvis.
1995.
Report from the roundtable on Rhizobium taxonomy, p. 365-370.
In
I. A. Tikhonovich, N. A. Provorov, V. I. Romanov, and W. E. Newton (ed.), Nitrogen fixation: fundamentals and applications. Kluwer Academic Publishers, Dordrecht, The Netherlands.
|
| 45.
|
Maidak, B. L.,
G. J. Olsen,
N. Larsen,
R. Ovebeek,
M. J. McCaughey, and C. R. Woese.
1996.
The Ribosomal Database Project (RDP).
Nucleic Acids Res.
24:82-85[Abstract/Free Full Text].
|
| 46.
|
Martínez-Romero, E., and J. Caballero-Mellado.
1996.
Rhizobium phylogenies and bacterial genetic diversity.
Crit. Rev. Plant Sci.
15:113-140.
|
| 47.
|
McKnight, T.
1949.
Efficiency of isolates of Rhizobium in the cowpea group, with proposed additions to this group.
Qld. J. Agric. Sci.
6:61-76.
|
| 48.
|
Moreira, F. M. S., and A. A. Franco.
1994.
Rhizobia-host interactions in tropical ecosystems in Brazil, p. 63-74.
In
J. I. Sprent, and D. McKey (ed.), Advances in legume systematics, part 5. The nitrogen factor. Royal Botanic Garden, Kew, England.
|
| 49.
|
Moreira, F. M. S.,
M. Gillis,
B. Pot,
K. Kersters, and A. A. Franco.
1993.
Characterization of rhizobia isolated from different divergence groups of tropical Leguminosae by comparative polyacrylamide gel electrophoresis of their total proteins.
Syst. Appl. Microbiol.
16:135-146.
|
| 50.
|
Norris, D. O.
1956.
Legumes and the Rhizobium symbiosis.
Emp. J. Exp. Agric.
24:247-270.
|
| 51.
|
Nour, S. M.,
J.-C. Cleyet-Marel,
D. Beck,
A. Effosse, and M. P. Fernandez.
1994.
Genotypic and phenotypic diversity of Rhizobium isolated from chickpea (Cicer arietinum L.).
Can. J. Microbiol.
40:345-354[Medline].
|
| 52.
|
Novikova, N. I.,
E. A. Pavlova,
N. I. Vorobjev, and E. V. Limeshchenko.
1994.
Numerical taxonomy of Rhizobium strains from legumes of the temperate zone.
Int. J. Syst. Bacteriol.
44:734-742[Abstract/Free Full Text].
|
| 53.
|
Oyaizu, H.,
S. Matsumoto,
K. Minanisawa, and T. Gamou.
1993.
Distribution of rhizobia in leguminous plants surveyed by phylogenetic identification.
J. Gen. Appl. Microbiol.
39:339-354.
|
| 54.
|
Padmanabhan, S.,
R.-D. Hirtz, and W. J. Broughton.
1990.
Rhizobia in tropical legumes: cultural characteristics of Bradyrhizobium and Rhizobium sp.
Soil Biol. Biochem.
22:23-28.
|
| 55.
|
Perrière, G., and M. Gouy.
1996.
WWW-Query: an on-line retrieval system for biological sequence banks.
Biochimie
78:364-369[Medline].
|
| 56.
|
Polhill, R. M.,
P. H. Raven, and C. H. Stirton.
1981.
Evolution and systematics of the Leguminosae, p. 1-26.
In
R. M. Polhill, and P. H. Raven (ed.), Advances in legume systematics, part 1. Royal Botanic Garden, Kew, England.
|
| 57.
|
Prin, Y.,
A. Galiana,
M. Ducousso,
N. Dupuy,
P. de Lajudie, and M. Neyra.
1993.
Les rhizobiums d'acacia.
Bois For. Trop.
238:5-19.
|
| 58.
|
Raven, P. H., and R. M. Polhill.
1981.
Biogeography of the Leguminosae.
In
R. M. Polhill, and P. H. Raven (ed.), Advances in legume systematics, Part 6. Royal Botanic Garden, Kew, England.
|
| 59.
|
Saitou, N., and M. Nei.
1987.
The neighbor-joining method: a new method for reconstructing phylogenetic trees.
Mol. Biol. Evol.
4:406-425[Abstract].
|
| 60.
|
Sneath, P. H. A.
1993.
Evidence from Aeromonas for genetic crossing-over in ribosomal sequences.
Int. J. Syst. Bacteriol.
43:626-629[Free Full Text].
|
| 61.
|
Sritharan, V., and R. H. J. Barker.
1991.
A simple method for diagnosing M. tuberculosis infection in clinical samples using PCR.
Mol. Cell. Probes
5:385-395[Medline].
|
| 62.
|
Stackebrandt, E., and B. M. Goebel.
1994.
Taxonomic note: a place for DNA-DNA reassociation and 16S rRNA sequence analysis in the present species definition in bacteriology.
Int. J. Syst. Bacteriol.
44:846-849[Abstract/Free Full Text].
|
| 63.
|
Thompson, S. C.,
G. Gemell, and R. J. Roughley.
1984.
Host specificity for nodulation among Australian acacias, p. 27-28.
In
J. R. Kennedy, and L. Cope (ed.), 7th Australian Legume Nodulation Conference. Australian Institute of Agricultural Science Occasional Publication no. 12. Australian Institute of Agricultural Science, Sydney, Australia.
|
| 64.
|
Vincent, J. M.
1970.
A manual for the practical study of root-nodule bacteria. International biological programme handbook no. 15.
Blackwell Science Publications, Oxford, England.
|
| 65.
|
Wayne, L. G.,
D. J. Brenner,
R. R. Colwell,
P. A. D. Grimont,
O. Kandler,
M. I. Krichevsky,
L. H. Moore,
W. E. C. Moore,
R. G. E. Murray,
E. Stackebrandt,
M. P. Starr, and H. G. Trüper.
1987.
Report of the ad hoc committee on reconciliation of approaches to bacterial systematics.
Int. J. Syst. Bacteriol.
37:463-464[Free Full Text].
|
| 66.
|
Willems, A., and M. D. Collins.
1993.
Phylogenetic analysis of rhizobia and agrobacteria based on 16S rRNA gene sequences.
Int. J. Syst. Bacteriol.
43:305-313[Abstract/Free Full Text].
|
| 67.
|
Young, J. P. W., and K. E. Haukka.
1996.
Diversity and phylogeny of rhizobia.
New Phytol.
133:87-94.
|
| 68.
|
Zhang, X.,
R. Harper,
M. Karsisto, and K. Lindström.
1991.
Diversity of Rhizobium bacteria isolated from the root nodules of leguminous trees.
Int. J. Syst. Bacteriol.
41:104-113.
|
Applied and Environmental Microbiology, October 1998, p. 3989-3997, Vol. 64, No. 10
0099-2240/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
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Abstract
Introduction
