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Applied and Environmental Microbiology, May 2000, p. 1939-1946, Vol. 66, No. 5
0099-2240/00/$04.00+0
Genotypic and Phenotypic Diversity of phlD-Containing
Pseudomonas Strains Isolated from the Rhizosphere of
Wheat
B. B.
McSpadden
Gardener,1,*
K. L.
Schroeder,1
S. E.
Kalloger,1
J. M.
Raaijmakers,2
L. S.
Thomashow,1 and
D. M.
Weller1
Root Disease and Biological Control Research
Unit, USDA Agricultural Research Service, Washington State
University, Pullman, Washington,1 and
Department of Phytopathology, Wageningen Agricultural
University, Wageningen, The Netherlands2
Received 8 October 1999/Accepted 2 March 2000
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ABSTRACT |
Production of 2,4-diacetylphloroglucinol (2,4-DAPG) in the
rhizosphere by strains of fluorescent Pseudomonas spp.
results in the suppression of root diseases caused by certain fungal
plant pathogens. In this study, fluorescent Pseudomonas
strains containing phlD, which is directly involved in the
biosynthesis of 2,4-DAPG, were isolated from the rhizosphere of wheat
grown in soils from wheat-growing regions of the United States and The
Netherlands. To assess the genotypic and phenotypic diversity present
in this collection, 138 isolates were compared to 4 previously
described 2,4-DAPG producers. Thirteen distinct genotypes, one of which represented over 30% of the isolates, were differentiated by
whole-cell BOX-PCR. Representatives of this group were isolated
from eight different soils taken from four different geographic
locations. ERIC-PCR gave similar results overall, differentiating 15 distinct genotypes among all of the isolates. In most cases, a single
genotype predominated among isolates obtained from each soil. Thirty
isolates, representing all of the distinct genotypes and geographic
locations, were further characterized. Restriction analysis of
amplified 16S rRNA gene sequences revealed only three distinct
phylogenetic groups, one of which accounted for 87% of the isolates.
Phenotypic analyses based on carbon source utilization profiles
revealed that all of the strains utilized 49 substrates and were unable to grow on 12 others. Individually, strains could utilize about two-thirds of the 95 substrates present in Biolog SF-N plates. Multivariate analyses of utilization profiles revealed phenotypic groupings consistent with those defined by the genotypic analyses.
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INTRODUCTION |
Strains of fluorescent
Pseudomonas spp. have been studied extensively as biological
control agents for soilborne plant pathogens (6, 14, 16, 29,
32). Strains that produce the antifungal compound
2,4-diacetylphloroglucinol (2,4-DAPG) play an important role in the
suppression of some root diseases when introduced into the rhizosphere
via seed or soil treatments (13, 26, 30). The genetic locus
responsible for the biosynthesis of 2,4-DAPG contains phlD,
a unique bacterial gene with similarity to plant genes encoding
chlacone/stilbene synthases (2, 3). The phlD gene
has been used as a genetic marker to detect 2,4-DAPG-producing Pseudomonas strains from several soils because the
occurrence of the gene is correlated with the production of 2,4-DAPG in
vitro (12, 18).
The fungus Gaeumannomyces graminis var. tritici
attacks the roots of wheat and barley, causing the disease take-all
(1, 8). Continuous cropping of wheat in fields that have
take-all results in a gradual decline in the incidence and severity of the disease, a phenomenon called take-all decline (TAD) (1, 10,
27). The production of phloroglucinol derivatives by
phlD-containing (phlD+)
Pseudomonas strains plays an important role in TAD (18,
19, 24). Populations sizes of phlD+
Pseudomonas strains were significantly larger in the rhizosphere of wheat grown in TAD (i.e., suppressive) soils than in non-TAD (i.e.,
conducive) soils from Washington State (18). In addition, phlD+ strains were associated with the roots of
wheat grown in soils from other regions of the United States and The
Netherlands with a history of wheat monoculture (21, 24).
Both 2,4-DAPG and phlD+ pseudomonads have been
isolated directly from the rhizosphere of wheat grown in a TAD soil
(4, 20). Pasteurization of a TAD soil resulted in the
concurrent loss of phlD+ Pseudomonas and
suppressiveness to take-all, while transfer of relatively small amounts
of TAD soil into a conducive soil resulted in the establishment of
2,4-DAPG-producing populations sufficient to suppress the disease
(19).
Several distinct groups of 2,4-DAPG-producing Pseudomonas
strains have been identified (12, 26). Two groups were
differentiated on the basis of antibiotic production, one that produced
2,4-DAPG, hydrogen cyanide, and pyoluteorin (PLT) and the other that
did not produce PLT but did produce the first two substances. The former group contained strains that were genotypically quite
homogeneous, despite having been isolated from the roots of different
crop plants grown in soils from different geographic locations
(12). In contrast, eight distinct genotypes were observed
among 32 isolates known to produce 2,4-DAPG but not PLT
(12). Sharifi-Tehrani et al. reported that the
2,4-DAPG-producing, PLT-negative strains as a group were more effective
as biological control agents against Pythium ultimum on
cucumbers and Fusarium oxysporum f.sp.
radicis-lycopersici on tomatoes, and they noted that the
amount of 2,4-DAPG produced by these strains in vitro correlated with
the amount of disease suppression in the latter system (26).
These results suggest that there is diversity among 2,4-DAPG-producing
Pseudomonas spp. in nature and that strains will differ in
their effectiveness as biological control agents.
Given the association of phlD-containing
Pseudomonas spp. with the phenomenon of TAD, we initiated
this study to better define the diversity of these bacteria in
different wheat-growing regions. Knowledge of the genotypic and
phenotypic characteristics of these strains may implicate specific
subsets of 2,4-DAPG producers in TAD and will allow us to distinguish
strains that have different abilities to suppress soilborne root
pathogens of wheat.
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MATERIALS AND METHODS |
Soils and plants.
A collection of soils from
wheat-growing regions of the United States and The Netherlands was
assembled (Table 1). Soils from fields
near Lind or Quincy, Wash., that had supported long-term wheat cropping
or native vegetation were described previously (18) and were
designated 4L and W or QT, QTN, Q, Q8r, QX-87, and QV, respectively.
Other United States soils were collected during 1997. Soils designated
FTAD1R and FFL1R were from fields on the campus of North Dakota State
University (NDSU), Fargo, that had been cropped continuously to wheat
since 1882 (NDSU research plot no. 2) or to flax since 1894 (NDSU
research plot no. 30). The HT soil was from a field near Hallock,
Minn., that had been cropped continuously to wheat for 10 years. The OC
soil was from a field on the campus of Cornell University, Ithaca,
N.Y., which historically had been cropped primarily to legumes but on
which wheat had been planted in 1997. Soils CV and CC were collected near Caldwell, Kans., from a roadside site near a field with a 100-year
history of wheat monoculture. Soils JMP and D27B were from fields near
Woensdrecht, The Netherlands, that had been cropped continuously to
wheat for 27 and 14 years, respectively. Soils were air dried, sieved,
and homogenized by hand and then maintained at room temperature prior
to potting.
Triticum aestivum cultivars used in this study included
cultivar Penawawa for the experiments carried out in United States
soils and cultivar Bussard for isolations from the two Dutch soils.
Plants were incubated in a growth chamber under controlled conditions
(15°C, 12 h light:12 h darkness) or grown in the field during
the summer of 1998 (Table
1).
Strain isolation and maintenance.
Strains designations
followed those given to the soils from which the strains originated.
While most strains were isolated from wheat plants grown under
controlled conditions, some were obtained from plants grown in
the field during the summer of 1998 at Lind or Quincy, Wash. (Table 1).
Soils from Quincy, Wash., were collected at the same site, but these
soils were incubated differently prior to isolation of
phlD+ Pseudomonas spp. (see references
17 to 19 for details). For example, the Q8 and QV soils were cycled multiple times to wheat in
pots at 4-week intervals (19). Reference strains CHA0, Pf5, F113, and PILH1 were isolated in a variety of ways from other crop
plants (11, 12, 15, 25, 29).
The large majority of
phlD+ Pseudomonas strains
were isolated by procedures similar to those described by Raaijmakers
et al.
(
18). Briefly, 1 g of roots was obtained from
wheat plants grown
in each soil. Root samples were maintained in
sterile test tubes
at 4°C prior to processing. For processing,
sterile distilled
water was added to each tube and bacteria were
dislodged from
the rhizosphere by 1 min of vortexing followed by a
1-min incubation
in a sonication bath. Samples were serially diluted
and plated
onto a modified King's medium B (KMB) agar,
KMB
+++, which has been shown to be selective for
fluorescent
Pseudomonas spp. (
28) (20 g of
proteose peptone [Difco Laboratories, Detroit,
Mich.] per liter),
1.2 g of KH
2PO
4, 1.5 g of
MgSO
4 · 7H
2O, 10
ml of glycerol, and
15 g of agar; supplemented with ampicillin
[40 µg/ml],
chloramphenicol [13 µg/ml], and cycloheximide [200
µg/ml]
[Sigma Chemical Co., St. Louis, Mo.]). In some instances,
one-third-strength KMB medium supplemented with the same concentrations
of antibiotics was used (1/3 KMB
+++).
Pseudomonas isolates containing the
phlD gene
were identified
as previously described (
18). Briefly,
colonies that hybridized
with the
phlD probe were isolated,
and the presence of
phlD was
confirmed by PCR amplification
with the gene-specific primers
Phl2A and Phl2B. These strains generally
produced a reddish-brown
pigment, a phenotype which has been correlated
with the presence
of 2,4-DAPG (
2). In all, 138
phlD-containing
Pseudomonas strains
were
identified. Frozen stock cultures of all strains were stored
in
Luria-Bertani broth plus 40% glycerol at
80°C.
Genotypic analyses.
Colonies growing on 1/3
KMB+++ or full-strength KMB agar medium were suspended in
sterile distilled water in polystyrene 96-well microtiter plates
(Costar, Corning, N.Y.) to an optical density at 600 nm
(OD600) of 0.15 to 0.45 and then subjected to freeze-thaw lysis. Analyses were replicated by amplifying at least two independent colonies of each strain on separate occasions.
Genomic fingerprints were obtained by amplification with BOX or
enterobacterial repetitive intergenic consensus sequence (ERIC)
primers
(
22). Individual reaction mixtures consisted of 5 µl
of
5× Gitschier buffer [83 mM
(NH
4)
2SO
4, 335 mM Tris-HCl (pH
8.8),
33.5 mM MgCl
2, 33.5 µM EDTA, 150 mM
-mercaptoethanol], 0.3 µl
of bovine serum albumin solution (14 mg/ml), 1.25 µl of dimethyl
sulfoxide, 9.64 µl of sterile distilled
water, 1.56 µl of the
deoxynucleoside triphosphates (2.0 mM each,
mixed 1:1:1:1), 1.0
µl of solutions of each primer (50 pmol/µl),
0.33 µl of
Taq DNA
polymerase (5 U/µl; Promega, Madison,
Wis.), and 5 µl of thawed
template (corresponding to 10
4
to 10
6 bacteria prior to lysis). Negative-control reaction
mixtures,
containing no cell lysate, were used for each amplified set.
Amplification
reactions were processed in PTC200 thermocyclers (MJ
Research,
Inc., Watertown, Mass.). Cycling conditions for ERIC-PCR were
as follows: 95°C for 7 min and then 30 cycles of 94°C for 1 min,
51°C for 1 min, and 65°C for 8 min, followed by a 16-min incubation
at 65°C; after cooling to 4°C, the PCR products were stored at
20°C. Cycling conditions for BOX-PCR were the same except that
the
annealing temperature was 50°C. Amplified products were separated
on
1.5% agarose gels in 0.5× Tris-borate-EDTA buffer for 5 to
6 h
at 140 V and 10°C. DNA banding patterns were visualized by
staining
with ethidium bromide and were analyzed with the software
GelCompar 4.0 (Applied Maths, Kortrijk, Belgium) by correlation-based
clustering (
22,
23). Groups were defined by the 95th
percentile
(near-minimum) similarity coefficient of replicate assays
for
identical
strains.
Amplified ribosomal DNA restriction analysis (ARDRA) was performed as
follows. Full-length 16S rRNA gene sequences were obtained
from each
strain by PCR amplification with the conserved eubacterial
primers 8F
(5'-AGA GTT TGA TCC TGG CTC AG-3') and 1492R (5'-ACG
GCT ACC TTG TTA
CGA CTT-3'), based on those defined by Weisburg
et al. (
31).
Reaction master mixes were prepared as described
above except that 25 pmol of each primer was used and 0.5 µg of
RNase was added for each
reaction. The cycling conditions were
as follows: 95°C for 5 min and
then 30 cycles of 94°C for 1 min,
50°C for 1 min, and 65°C for 4 min, followed by an 8-min incubation
at 65°C; after a 4°C soak, the
PCR products were stored at
20°C.
Restriction digestions consisted
of 10 µl of a PCR mixture in
a total volume of 30 µl with 10 U of a
single restriction enzyme
(New England Biolabs Inc., Beverly, Mass.).
Reaction mixtures
were incubated at either 37°C (
HaeIII,
HhaI,
HinfI,
MspI,
RsaI,
or
Sau96I) or 60°C (
BstUI or
TaqI) for
2 to 4 h and then stored
at
20°C. Digestion products were
subjected to electrophoresis
on 2% agarose gels in 0.5×
Tris-borate-EDTA buffer for 2 to 3
h at 140 V. Banding patterns
were visualized by ethidium bromide
staining and scored by comparison
to a 100-bp DNA
ladder.
Phenotypic tests.
Substrate utilization profiles were
generated by using Biolog SF-N plates (Biolog, Inc., Hayward, Calif.).
Colonies (
10 days old) grown on KMB agar were suspended in 1 ml of
sterile distilled water, pelleted by centrifugation, and resuspended in
0.5 ml of sterile distilled water. The concentration of washed cells
was adjusted with water to an OD600 of 0.05 to 0.10 by
using a microplate spectrophotometer (MR5000; Dynatech Inc.,
Burlington, Mass.). A 10-µl aliquot of washed cells (approximately
106 cells) was added to each well of a Biolog SF-N plate
preloaded with 90 µl of sterile distilled water per well. The plates
were incubated at room temperature in the dark. Bacterial growth was assayed spectrophotometrically at 600 nm after 3 and 7 days of incubation. Multivariate statistical analyses of the data obtained were
performed with the program MULTIV 1.2.1 (Valerio De Patta Pillar,
Department of Ecology, Universidade Federal do Rio Grande do Sul, Porto
Alegre, RS, Brazil). Groups were defined by the 95th percentile
(near-minimum) similarity coefficient of replicate assays for identical strains.
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RESULTS |
Genomic fingerprinting of strains.
Whole-cell rep-PCR
amplifications of the isolates yielded complex genomic fingerprints
consisting of 10 to 30 amplification products ranging in size from 100 to 3,000 bp. Among the 142 phlD+ isolates, 13 distinct genomic fingerprints were observed by rep-PCR performed
with the primer BOXA1R (Fig. 1). A
similar, but not identical, clustering was obtained with the ERIC
primer set (Fig. 2). Three differences
were observed. First, strains CHA0 and Pf5 were separated by ERIC- but
not by BOX-PCR (Table 2). This was likely
due to the low signal-to-noise ratio obtained by using the whole-cell
amplification protocol for these two strains, because the ERIC patterns
for these two strains clustered together when isolated genomic DNA was
used as template (data not shown). Second, the large BOX-defined D
genotype was split into three distinct clusters, D1, D2, and E, when
the ERIC data was used (Table 2). However, visual inspection of the
fingerprint patterns of these groups indicated that a majority of
amplified bands comigrated, and although the three clusters were
distinct by the statistical criterion applied, they were closely
related (>42.5% similar, compared to the 95th percentile near-minimum
definition of identity of 48.5% similarity) (Fig. 2). A similar split
occurred with the JMP isolates; ERIC genotypes G and H corresponded to
BOX genotype F (Table 2). The third difference involved the splitting
of ERIC genotype I into BOX genotypes G, H, and I and the close
association of these groups with genotype J as defined by both BOX and
ERIC fingerprint patterns (cf. Fig. 1 and 2; Table 2). Again, the similarity scores calculated for these clusters were close to the
values defining identity, except that BOX genotype H appeared very
distinct.
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FIG. 1.
Cluster analysis of genomic fingerprint patterns of
phlD-containing Pseudomonas strains generated by
PCR amplification of whole-cell template with the BOXA1R primer. Only
patterns of strains representing unique genotypes isolated from
individual soils are shown. Two independent amplifications were used
for each strain. Using GelCompar 4.0, the UPGMA algorithm was applied
to the similarity matrix generated from the tracks of the whole
patterns by using Pearson's correlation coefficient. The similarity
coefficient used to define distinct groups (see Materials and Methods)
is noted (*). Distinct groups of genotypes, labeled alphabetically A
through N, are discussed more fully in the text.
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FIG. 2.
Cluster analysis of genomic fingerprint patterns of
phlD-containing Pseudomonas strains generated by
PCR amplification of whole-cell template with the ERIC primer set. Only
patterns of strains representing unique genotypes isolated from
individual soils are shown. Two independent amplifications were used
for each strain. Using GelCompar 4.0, the UPGMA algorithm was applied
to the similarity matrix generated from the tracks of the whole
patterns by using Pearson's correlation coefficient. The similarity
coefficient used to define distinct groups (see Materials and Methods)
is noted (*). Distinct groups of genotypes, labeled alphabetically A
through N, are discussed more fully in the text.
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The largest set of genotypically similar isolates, defined as BOX group
D or ERIC groups D1, D2, and E, contained over 30%
of all isolates.
This set included isolates from eight soils (FTAD1R,
FFL1R, OC, 4L, W,
QT, Q8r, and QV) in four different locations
(Fargo, Ithaca, Lind, and
Quincy). We refer to this apparently
cosmopolitan set of isolates as
the Q8r1 supergroup. In contrast,
most of the other genotypic groups
contained isolates from a single
geographic location. Only three other
groups, ERIC I, J, and M,
contained isolates from more than one
location (Table
2).
Isolates from the same soil that displayed the same genomic fingerprint
are listed together in Table
1. In general, only
one or two
phlD+ strains were isolated from the rhizosphere
of wheat grown in
each soil under a given set of conditions. Multiple
genotypes
were obtained from Quincy and Fargo soils (Table
3), but these
were isolated from wheat
plants that had been grown under different
light, temperature, and
moisture regimes for the Quincy soils
and different cropping histories
at Fargo (Table
1). Four distinct
ERIC genotypes were isolated from the
Quincy soils; however, only
one genotype predominated under each set of
conditions. For example,
8 of 10 isolates from soil QX-87 were ERIC
genotype C, while 6
of 9 isolates from soil QT were ERIC genotype F. Similarly, Fargo
soils FTAD1R, from a long-term wheat field, and FFL1R,
from an
adjoining long-term flax field, yielded four distinct
genotypes,
but a single dominant genotype was isolated from each (ERIC
genotypes
D1 and J, respectively). Thirty type strains, representing
the
distinct rep-PCR genotypes present in each soil, were identified
for further analyses (Table
2).
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TABLE 3.
Number of isolates representing distinct genotypes
of phlD-containing Pseudomonas strains
isolated from Quincy and Fargo soils
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ARDRA.
Patterns of restriction fragment length polymorphisms
generated by using eight different restriction enzymes indicated that amplified 16S rRNA gene sequences were very similar for most of the 30 representative phlD+ strains (Table
4). Sequences from CHA0 and Pf5 differed
from those of all other phlD+ strains in four of
the eight digests. The enzyme TaqI revealed similarities
between these two strains, F113 and HT5-1. No restriction fragment
length polymorphisms were found among 16S rRNA gene sequences from
phlD+ strains upon digestion with
BstUI, HinfI, or Sau96I.
Growth on 95 different substrates.
The same
phlD+ isolates representing distinct genotypes
were examined for their ability to grow in vitro on a variety of carbon sources. After 3 days of incubation, each strain grew on 63 to 68 of
the 95 different substrates present in Biolog SF-N plates, except CHA0
and Pf5, which grew on no more than 61 of the substrates. The number of
substrates sustaining growth of each strain increased by approximately
10% after 7 days. However, not all of the strains used the same set of
substrates. Forty-nine substrates were utilized by all
phlD+ strains (Table
5); 17 of these allowed for rapid growth
to saturation (OD600 
0.375 after 3 days). However, 12 substrates did not support the growth of any strain (Table 5). Strains
CHA0 and Pf5 grew on significantly fewer substrates than the other
phlD+ strains, and they differed from the other
strains in the utilization of seven substrates (Table 5). The carbon
source utilization profiles of strains lacking phlD differed
greatly both from one another and from those of the
phlD+ strains (data not shown).
Multivariate analyses of the utilization profiles of the representative
strains were performed (Fig.
3). Four to
11 distinct
clusters were observed for the 30
phlD+ isolates, depending on the algorithm used
for clustering (Fig.
3) and the length of incubation (data not shown).
Three groups
were conserved in all of the analyses, so they can be
considered
phenotypically distinct. These groups are represented by
strains
CHA0, Pf5, and W4-4. Three other groups, represented by HT5-1,
JMP6 or JMP7, and Q2-87 or Q2-2, also were distinct in most cases.
The
remaining 21 strains were consolidated into a single cluster
by the
simple linkage algorithm but split into three distinct
groups by the
complete linkage algorithm (Fig.
3). To determine
which clustering
might better describe these strains, two types
of ordination analyses,
with different mathematical underpinnings,
were used. Both principal
coordinate and principal component analyses
more strongly supported the
model of a single distinct group (data
not shown). Data from 3-day and
7-day incubations gave very similar
results in terms of the topology of
the dendrograms; however,
the longer incubations resulted in fewer
distinct groups because
most strains had grown to their maximal density
on a greater percentage
of substrates (data not shown).
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FIG. 3.
Cluster analysis of carbon source utilization patterns
of phlD-containing Pseudomonas strains cultured
for 3 days. Only strains representing unique genotypes isolated from
individual soils were assayed. Two independent assays were performed on
each strain. Using MULTIV 1.2.1, the simple linkage (a) and complete
linkage (b) algorithms were applied to the similarity matrix generated
by using Pearson's correlation coefficient. The similarity coefficient
used to define distinct groups (see Materials and Methods) is noted
(*). Distinct groups of phenotypes, labeled alphabetically A
through G, are discussed more fully in the text.
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DISCUSSION |
While phlD+ Pseudomonas spp. have been
found in soils naturally suppressive to take-all (18, 19),
it is not clear that isolates from such soils contribute equally to the
phenomenon of TAD. As a first step toward characterizing these
populations, we studied the genotypic and phenotypic diversity of a
large collection of isolates from soils of different wheat-growing
regions of the United States and The Netherlands. Using rep-PCR,
we identified at most 17 genotypes distributed among three ARDRA
groups (Table 2). Previously, Keel et al. characterized a smaller
collection of phloroglucinol-producing strains and found seven
randomly amplified polymorphic DNA-defined genotypes belonging to
three ARDRA groups (12). The genotypic distinctions of
the 14 strains included in both studies (CHA0, Pf5, F113, PILH1, and
all QX-87 isolates) were entirely consistent with one another. Each of
the newly identified genotypes in our study occurred in soils from
locations not previously examined. Only one or two genotypes were
isolated from each soil under a particular set of conditions. One
possible explanation for this observation is that a single
genotypically distinct population of phlD+
Pseudomonas strains dominates in the rhizosphere under a
given set of environmental conditions. Among the three BOX genotypes isolated from the Quincy TAD soil, there were at least two distinctly different phenotypes based on carbon source utilization (Fig. 3 and
Table 2). It may be that the different types of 2,4-DAPG producers had
differentially adapted to changing environmental conditions and/or that
some sort of microbial succession had taken place. The dominant
genotypes obtained from the two Fargo soils, FTAD1R34 and FFL1R22, also
differed phenotypically (Table 2). These two soils differed in their
cropping history, indicating that wheat and flax roots may enrich for
different phlD+ genotypes over time.
The strains isolated in this study may differ in their effectiveness
against fungal root pathogens of wheat and barley. Different genotypes
of 2,4-DAPG-producing Pseudomonas strains have been reported
to differ in their capacity to inhibit the growth of Fusarium on tomatoes and of Pythium on cucumbers
(26). Differences in biocontrol capacities may be due to one
or more of the following factors. First, different genotypes produce
different amounts of 2,4-DAPG and other antifungal metabolites in vitro
(12, 26), and such differences also may occur in situ.
Second, the degree to which strains can colonize the rhizosphere may
impact their ability to suppress invading pathogens. Indeed, evidence
that DAPG producers differ substantially in their ability to colonize wheat roots when introduced as a seed or soil treatment is now mounting
(20, 21; B. B. McSpadden Gardener and D. M. Weller, unpublished data). Lastly, there may be undiscovered
phenotypic characteristics of certain genotypes that positively
contribute to disease suppression, even though the production of
2,4-DAPG alone can explain the ability of some strains to suppress
fungal root pathogens (7, 13, 30).
The genotypic data presented in this study could aid substantially in
the identification and selection of new 2,4-DAPG-producing biocontrol
agents. Currently, most biocontrol studies are initiated by collecting
large numbers of isolates and then laboriously screening them for the
ability to suppress the growth of plant pathogens (5).
Because it is reasonable to assume that isolates of the same genotype
will perform similarly, fewer isolates may need to be tested in
greenhouse and field studies. With rep-PCR, novel genotypes can be
easily recognized and selected for more-intensive analyses.
Additionally, our screening program can be focused on genotypes that
are known to be particularly effective on wheat. In our collection,
nearly one-third of the isolates were genotypically similar to strain
Q8r1-96. This strain very effectively colonizes wheat roots and can
suppress take-all when applied at low doses (19, 21).
Preliminary evidence suggests that other isolates of this genotype also
have this ability (B. B. McSpadden Gardener and D. M. Weller,
unpublished data). The widespread occurrence of this genotype may be
indicative of the generalist nature of Q8r1-like pseudomonads and their
ability to adapt successfully to the wheat rhizosphere regardless of
the prevailing soil conditions. It is possible, too, that subtle
phenotypic differences that represent site-specific adaptations may be
discovered by comparing isolates of this group.
Of the assays used in this study, rep-PCR was the most discriminating
because it revealed the largest number of distinct groups. Previously,
it was noted that rep-PCR has a greater capacity to distinguish strains
than do 16S rRNA sequence-based approaches (22). However,
there has not been a similar comparison of taxonomic abilities of
rep-PCR and phenotypic assays based on carbon source utilization. In
this instance, rep-PCR proved to be more effective in discriminating
between strains than the Biolog SF-N assays. In our study of carbon
source utilization, over 75% of the substrates used by any individual
strain could be utilized by all of the phlD+
isolates (Table 5). Most differences among strains were not in the
number or types of carbon sources used but rather in the degree to
which they supported growth. This homogeneity of growth phenotypes is
reflected in the relatively high similarity coefficients used to define
distinct clusters (Fig. 3). While only at most seven groups were
defined by using the Biolog data, these groups were all strongly
associated with particular genotypes. Thus, there is hidden genotypic
diversity within these phenotypic groups, suggesting that there is also
hidden phenotypic diversity which is not detected with the Biolog
assay. The carbon sources present in the Biolog assays were selected
for their ability to discriminate microbial taxa and included some
substrates not likely to be found in the rhizosphere. While a modified
system using substrates found in wheat rhizospheres may be more useful
in identifying strains with different abilities to colonize this
environment, it remains to be seen whether such an in vitro test can
detect the same degree of diversity as the genomic fingerprint assays.
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FOOTNOTES |
*
Corresponding author. Mailing address: Root Disease and
Biological Control Research Unit, USDA Agricultural Research Service, Pullman, WA 99164-6430. Phone: (509) 335-1116. Fax: (509) 335-7674. E-mail: mcspadde{at}mail.wsu.edu.
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Applied and Environmental Microbiology, May 2000, p. 1939-1946, Vol. 66, No. 5
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Abstract
Introduction
